**Meet the editor**

Prof. Anica Dricu was awarded his PhD in Experimental Pathology by the University of Medicine " Karolinska Institute" , Stockholm, in 1997. She became Docent in Molecular Oncology at the University of Medicine "Karolinska Institute", Stockholm, in 2007. Dr. Anica Dricu was appointed Professor at the University of Medicine and Pharmacy Craiova, in 2009. As a faculty member

at the University of Medicine and Pharmacy Craiova, Prof. Anica Dricu is involved in teaching medical students, residents, clinical fellows, graduate and undergraduate students. In addition to her research and teaching activities, Prof. Dricu is actively involved in service at national and institutional levels; she is currently, director of the Doctoral School and head of the Division of Biochemistry at the University of Medicine and Pharmacy Craiova and she also serves as a member of the National Council for Titles, Diplomas and Certificates. She is a editorial member of several medical journal. Her special interests include: stem cell research, experimental pathology, molecular oncology, medical biochemistry.

Contents

**Preface VII**

Byron Baron

Kubareva

**Section 1 Gene Expression and Methylation 1**

**Factors and DNA Methylation 3**

**and Prokaryotic System 27**

Marina L. Mokrishcheva

Henk J Blom and Rita Castro

Chapter 5 **Methylation in Tumorigenesis 119**

**Processes 117**

A. Brown

Chapter 1 **Breaking the Silence: The Interplay Between Transcription**

**Section 2 DNA-Methyltransferases: Structure and Function in Eukaryotic**

A. Yu. Ryazanova, L. A. Abrosimova, T. S. Oretskaya and E. A.

Dmitry V. Nikitin, Attila Kertesz-Farkas, Alexander S. Solonin and

Ruben Esse, Paula Leandro, Isabel Rivera, Isabel Tavares de Almeida,

Melissa A. Edwards, Pashayar P. Lookian, Drew R. Neavin and Mark

Chapter 2 **Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization 29**

Chapter 3 **Bifunctional Prokaryotic DNA-Methyltransferases 71**

Chapter 4 **Deciphering Protein Arginine Methylation in Mammals 91**

**Section 3 Protein Arginine Methylation in Mammals 89**

**Section 4 Cancer Research Through Study of Methylation Cell**

### Contents

#### **Preface XI**


Chapter 6 **Circulating Methylated DNA as Biomarkers for Cancer Detection 137**

Hongchuan Jin, Yanning Ma, Qi Shen and Xian Wang

Chapter 7 **DNA Methylation, Stem Cells and Cancer 153** Anica Dricu, Stefana Oana Purcaru, Alice Sandra Buteica, Daniela Elise Tache, Oana Daianu, Bogdan Stoleru, Amelia Mihaela Dobrescu, Tiberiu Daianu and Ligia Gabriela Tataranu

Preface

including cancer.

There is a widespread interest in the today scientific literature for methylation field, which started to be published in the early 60's and continues to be a future line of research. This book represents a comprehensively reviewed literature on the importance of methylation processes in human health and disease. The book, covers the basic mechanism of DNA and protein methylation, along with the role of mRNA cap methylation in viral replication, gene expression and viral pathogenesis. Human health risks from metals methylation in the natu‐ ral environment has been well describe in the literature. As a consequence, the formation processes, the biotic and abiotic degradation and the accumulation of the methylated metals

DNA methylation is a well-characterized process, allowing cells to control gene expression, while the study of histone methylation is more recent. The enzymes responsible for histone methylation (histone methyltransferases and histone demethylases) are important for tran‐ scriptional regulation in both normal and abnormal states, representing an important target for drug discovery. The interconection between DNA methylation and other regulatory mol‐ ecules such as: enzymes, transcription factors, proteins and growth factors is discussed, pro‐ viding key information about the mechanisms that trigger cell proliferation, differentiation, aging and malignant transformation. This textbook strongly point out the importance of me‐ thylated DNA as a biological marker of cancer an also gives the reader insights into the re‐ cently emerged treatment modalities targeting methylation mechanism, in various diseases

The textbook addresses the following topics: Gene expression and methylation, DNA-meth‐ yltransferases: structure and function in eukaryotic and prokaryotic system; Protein arginine methylation in mammals; Cancer research through study of methylation cell processes; Bac‐

The book aims at the advanced undergraduate and graduate biomedical students and re‐ searchers working in the epigenetic area, providing readers with both classical and relevant recent discoveries that have been made in the research field of methylation and also point‐

**Prof. Anica Dricu**

Faculty of Medicine Craiova, Romania

University of Medicine and Pharmacy

teria, viruses and metals methylation: risk and benefit for human health.

ing out pathways where questions remain.

and metalloids in the aquatic environment is reviwed in the book.


### Preface

Chapter 6 **Circulating Methylated DNA as Biomarkers for Cancer**

Chapter 7 **DNA Methylation, Stem Cells and Cancer 153**

Chapter 8 **DNA Methylation in the Pathogenesis of Head and**

Hongchuan Jin, Yanning Ma, Qi Shen and Xian Wang

Anica Dricu, Stefana Oana Purcaru, Alice Sandra Buteica, Daniela Elise Tache, Oana Daianu, Bogdan Stoleru, Amelia Mihaela Dobrescu, Tiberiu Daianu and Ligia Gabriela Tataranu

Zvonko Magić, Gordana Supić, Mirjana Branković-Magić and

**Section 5 Bacteria, Viruses and Metals Methylation: Risk and Benefit for**

Chapter 9 **Host-Mimicking Strategies in DNA Methylation for Improved**

Chapter 10 **Messenger RNA Cap Methylation in Vesicular Stomatitis Virus, a Prototype of Non‐Segmented Negative‐Sense**

**Detection 137**

**VI** Contents

**Neck Cancer 185**

**Human Health 217**

Hirokazu Suzuki

**RNA Virus 237**

Jianrong Li and Yu Zhang

**Aquatic Systems 271** Robert P. Mason

Chapter 11 **The Methylation of Metals and Metalloids in**

**Bacterial Transformation 219**

Nebojša Jović

There is a widespread interest in the today scientific literature for methylation field, which started to be published in the early 60's and continues to be a future line of research. This book represents a comprehensively reviewed literature on the importance of methylation processes in human health and disease. The book, covers the basic mechanism of DNA and protein methylation, along with the role of mRNA cap methylation in viral replication, gene expression and viral pathogenesis. Human health risks from metals methylation in the natu‐ ral environment has been well describe in the literature. As a consequence, the formation processes, the biotic and abiotic degradation and the accumulation of the methylated metals and metalloids in the aquatic environment is reviwed in the book.

DNA methylation is a well-characterized process, allowing cells to control gene expression, while the study of histone methylation is more recent. The enzymes responsible for histone methylation (histone methyltransferases and histone demethylases) are important for tran‐ scriptional regulation in both normal and abnormal states, representing an important target for drug discovery. The interconection between DNA methylation and other regulatory mol‐ ecules such as: enzymes, transcription factors, proteins and growth factors is discussed, pro‐ viding key information about the mechanisms that trigger cell proliferation, differentiation, aging and malignant transformation. This textbook strongly point out the importance of me‐ thylated DNA as a biological marker of cancer an also gives the reader insights into the re‐ cently emerged treatment modalities targeting methylation mechanism, in various diseases including cancer.

The textbook addresses the following topics: Gene expression and methylation, DNA-meth‐ yltransferases: structure and function in eukaryotic and prokaryotic system; Protein arginine methylation in mammals; Cancer research through study of methylation cell processes; Bac‐ teria, viruses and metals methylation: risk and benefit for human health.

The book aims at the advanced undergraduate and graduate biomedical students and re‐ searchers working in the epigenetic area, providing readers with both classical and relevant recent discoveries that have been made in the research field of methylation and also point‐ ing out pathways where questions remain.

> **Prof. Anica Dricu** University of Medicine and Pharmacy Faculty of Medicine Craiova, Romania

**Section 1**

**Gene Expression and Methylation**

**Gene Expression and Methylation**

**Chapter 1**

**Breaking the Silence: The Interplay Between**

**Transcription Factors and DNA Methylation**

DNA methylation is best known for its role in gene silencing through a methyl group (CH3) being added to the 5' carbon of cytosine bases (giving 5-methylcytosine) in the promoters of genes leading to supression of transcription [1]. However this is far from the whole story.

*De novo* methylation, which involves the addition of a methyl group to unmodified DNA, is described as an epigenetic change because it is a chemical modification to DNA not a change brought about by a DNA mutation. Unlike mutations, methylation changes are potentially reversible. Epigenetic changes also include changes to DNA-associated molecules such as histone modifications, chromatin-remodelling complexes and other small non-coding RNAs including miRNAs and siRNAs [2]. These changes have key roles in imprinting (gene-ex‐ pression dependent on parental origin), X chromosome inactivation and heterochromatin

DNA methylation leading to silencing is a very important survival mechanism used on re‐ petitive sequences in the human genome, which come from DNA and RNA viruses or from mRNA and tRNA molecules that are able to replicate independently of the host genome. Such elements need to be controlled from spreading throughout the genome, by being si‐ lenced through CpG methylation, as they cause genetic instability and activation of onco‐ genes [6-10]. Such elements can be categorised into three groups: SINEs (Small Interspersed Nuclear Elements), LINEs (Long Interspersed Nuclear Elements) and LTRs (Long Terminal Repeats) [6,11-13]. Repetitive sequences are recognised by Lymphoid-Specific Helicase (LSH) also known as the 'heterochromatin guardian' [14,15], which additionally acts on sin‐

> © 2013 Baron; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

Additional information is available at the end of the chapter

Byron Baron

**1. Introduction**

http://dx.doi.org/10.5772/55021

formation among others [3-5].

gle-copy genes [16].

### **Breaking the Silence: The Interplay Between Transcription Factors and DNA Methylation**

#### Byron Baron

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/55021

#### **1. Introduction**

DNA methylation is best known for its role in gene silencing through a methyl group (CH3) being added to the 5' carbon of cytosine bases (giving 5-methylcytosine) in the promoters of genes leading to supression of transcription [1]. However this is far from the whole story.

*De novo* methylation, which involves the addition of a methyl group to unmodified DNA, is described as an epigenetic change because it is a chemical modification to DNA not a change brought about by a DNA mutation. Unlike mutations, methylation changes are potentially reversible. Epigenetic changes also include changes to DNA-associated molecules such as histone modifications, chromatin-remodelling complexes and other small non-coding RNAs including miRNAs and siRNAs [2]. These changes have key roles in imprinting (gene-ex‐ pression dependent on parental origin), X chromosome inactivation and heterochromatin formation among others [3-5].

DNA methylation leading to silencing is a very important survival mechanism used on re‐ petitive sequences in the human genome, which come from DNA and RNA viruses or from mRNA and tRNA molecules that are able to replicate independently of the host genome. Such elements need to be controlled from spreading throughout the genome, by being si‐ lenced through CpG methylation, as they cause genetic instability and activation of onco‐ genes [6-10]. Such elements can be categorised into three groups: SINEs (Small Interspersed Nuclear Elements), LINEs (Long Interspersed Nuclear Elements) and LTRs (Long Terminal Repeats) [6,11-13]. Repetitive sequences are recognised by Lymphoid-Specific Helicase (LSH) also known as the 'heterochromatin guardian' [14,15], which additionally acts on sin‐ gle-copy genes [16].

© 2013 Baron; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

DNA methylation generally occurs when a cytosine is adjacent 5' to a guanine, called a CpG di‐ nucleotide. Such dinucleotides are spread all over the genome and over 70% of CpGs are me‐ thylated. Clusters of CpGs, called CpG Islands (CGI), consist of stretches of 200–4000bp that are 60 to 70% G/C rich, found in TATAless promoters and/or first exons of genes [17-19].

DNA methylation shows different effects on gene expression, brought about by an interplay of several different mechanisms, which can be grouped into three categories [2,54]: i. effects on di‐ rect transcription factor binding at CpG dinucleotides; ii. binding of specific methylation-rec‐ ognition factors (such as MeCP1 and MeCP2) to methylated DNA; iii. changes in chromatin

Breaking the Silence: The Interplay Between Transcription Factors and DNA Methylation

http://dx.doi.org/10.5772/55021

5

Key stages in development make use of methylation to switch on/off and regulate gene ex‐ pression. DNA methylation was shown to be essential for embryonic development through homozygous deletion of the mouse Mtase gene which leads to embryonic lethality [52]. Germline cells show 4% less methylation in CGI promoters, including almost all CGI pro‐

Immediately after fertilisation but before the first cell division, the paternal DNA undergoes active demethylation throughout the genome [55-58]. After the first cell cycle, the maternal DNA undergoes passive demethylation as a result of a lack of methylation maintenance af‐ ter mitosis [56,59], and this genome-wide demethylation continues, except for the imprinted

After implantation, the genome (except for CGIs) undergoes *de novo* methylation [54]. Active demethylation subsequently occurs during early embryogenesis [62] with tissue-specific genes undergoing demethylation in their respective tissues, creating a methylation pattern

Somatic cells go through the process of aging as they divide and replicate. Aging is charac‐ terised by a genome-wide loss and a regional gain of DNA methylation [63]. CGI promoters present an increase in DNA methylation in normal tissues of older individuals at several sites throughout the genome [64,65]. This causes genomic instability and deregulation of tis‐ sue-specific and imprinted genes as well as silencing of tumour suppressor genes (control‐ ling cell cycle, apoptosis or DNA repair) through hypermethylation of promoter CGIs [5,66]. The age-related change in methylation was shown in a genome-wide CGI methylation study comparing small intestine (and other tissues) from 3-month-old and 35-month-old mice, which presented linear age-related increased methylation in 21% and decreased methylation in 13% of tested CGIs with strong tissue-specificity [67]. Furthermore, human intestinal agerelated aberrant methylation was shown to share similarities to mouse [67]. Although the majority of CGIs methylated in tumours are also methylated in a selection of normal tissues during aging, particular tumours exhibit methylation in specific promoters and are thus said

Aging appears to exhibit common methylation features with carcinogenesis and in fact these processes share a large number of hypermethylated genes such as ER, IGF2, N33 and MyoD in colon cancer, NKX2-5 in prostate cancer and several Polycomb-group protein target genes, which suggests they probably have common epigenetic mechanisms driving them [68-70].

which is maintained in the adult, giving each cell type a unique epigenome. [54].

structure.

**2. Methylation in development and aging**

genes, until the formation of the blastocyst [60,61].

to display a CpG island methylator phenotype (CIMP) [65].

moters of germline-specific genes, compared to somatic cells [21].

In the human genome almost 50% of transcription start sites (TSS) [20], and about 70% of all genes contain CGIs [21,22]. CGIs present in the promoters or first exons of ubiquitously ex‐ pressed housekeeping and tightly regulated developmental genes are usually hypomethy‐ lated, irrispective of transcription activity [1,19,21,23-29] and become silenced when they are hypermethylated [20]. On the contrary, promoters of some tissue-specific genes, with low CpG density, are commonly methylated without loss of transcription activity [21,26,30].

Many active promoters were shown to contain a low percentage of methylation (4 - 7%) in‐ dicating that supression through DNA hypermethylation is density-dependent [21]. The op‐ posite was shown for the cAMP-responsive element (CRE)-binding sites, which are found in the promoters of numerous tissue-specific genes, including hormone-coding and viral genes [31]. Methylation of the CpG at the centre of the CRE sequence inhibits transcription, by in‐ hibiting transcription factor binding, indicating that methylation at specific CpG sites can contribute to the regulation of gene expression [32].

Low-density gene body methylation has been observed in actively transcribed genes and is implicated in reducing 'transcriptional noise' – the inappropriate gene transcription from al‐ ternative start sites or in cells where it is meant to be silenced [33]. Moreover it is thought to inhibit antisense transcription, to direct RNA splicing and to have a role in replication tim‐ ing [34-37]. Methylation is thought to play a role in transcriptional elongation, termination and splicing regulation due to higher CpG methylation in exons compared to introns [38,39] and the transacription start and termination regions lacking methylation [40,41].

CpG dinucleotides are not the only sequences that can be methylated, although non-CpG methylation was thought to be infrequent until the methylome of embryonic stem (ES) cells revealed that such non-CpG methylation, generally occuring in a CHG and CHH context, constitues 25% of total methylated sites in the genome [40]. Non-CpG methylation was also reported in some genes from mouse ES cells [42,43]. The distribution of such non-CG meth‐ ylation was high in gene bodies and low in promoters and regulatory sequences with almost complete loss during differentiation [40].

DNA methyltransferases (DNMTs) are enzymes that catalyse the addition of methyl groups to cytosine residues in DNA. Mammals have three important DNMTs: DNMT1 is responsi‐ ble for the maintenance of existing methylation patterns following DNA replication, while DNMT3a and DNMT3b are *de novo* methyltransferases [1,44-46]. As a result of DNA replica‐ tion, fully methylated DNA becomes hemi-methylated and DNMT1 binds hemi-methylated DNA to add a methyl group to the 5′ carbon of cytosines [2].

Overall, most DNA methylation changes can be observed invariantly in all tissues [47]. However, the small portion of tissue-specific methylation has a profound effect on cellular activity including cell differentiation, disease and cancer [48-53].

DNA methylation shows different effects on gene expression, brought about by an interplay of several different mechanisms, which can be grouped into three categories [2,54]: i. effects on di‐ rect transcription factor binding at CpG dinucleotides; ii. binding of specific methylation-rec‐ ognition factors (such as MeCP1 and MeCP2) to methylated DNA; iii. changes in chromatin structure.

#### **2. Methylation in development and aging**

DNA methylation generally occurs when a cytosine is adjacent 5' to a guanine, called a CpG di‐ nucleotide. Such dinucleotides are spread all over the genome and over 70% of CpGs are me‐ thylated. Clusters of CpGs, called CpG Islands (CGI), consist of stretches of 200–4000bp that are

In the human genome almost 50% of transcription start sites (TSS) [20], and about 70% of all genes contain CGIs [21,22]. CGIs present in the promoters or first exons of ubiquitously ex‐ pressed housekeeping and tightly regulated developmental genes are usually hypomethy‐ lated, irrispective of transcription activity [1,19,21,23-29] and become silenced when they are hypermethylated [20]. On the contrary, promoters of some tissue-specific genes, with low CpG density, are commonly methylated without loss of transcription activity [21,26,30].

Many active promoters were shown to contain a low percentage of methylation (4 - 7%) in‐ dicating that supression through DNA hypermethylation is density-dependent [21]. The op‐ posite was shown for the cAMP-responsive element (CRE)-binding sites, which are found in the promoters of numerous tissue-specific genes, including hormone-coding and viral genes [31]. Methylation of the CpG at the centre of the CRE sequence inhibits transcription, by in‐ hibiting transcription factor binding, indicating that methylation at specific CpG sites can

Low-density gene body methylation has been observed in actively transcribed genes and is implicated in reducing 'transcriptional noise' – the inappropriate gene transcription from al‐ ternative start sites or in cells where it is meant to be silenced [33]. Moreover it is thought to inhibit antisense transcription, to direct RNA splicing and to have a role in replication tim‐ ing [34-37]. Methylation is thought to play a role in transcriptional elongation, termination and splicing regulation due to higher CpG methylation in exons compared to introns [38,39]

CpG dinucleotides are not the only sequences that can be methylated, although non-CpG methylation was thought to be infrequent until the methylome of embryonic stem (ES) cells revealed that such non-CpG methylation, generally occuring in a CHG and CHH context, constitues 25% of total methylated sites in the genome [40]. Non-CpG methylation was also reported in some genes from mouse ES cells [42,43]. The distribution of such non-CG meth‐ ylation was high in gene bodies and low in promoters and regulatory sequences with almost

DNA methyltransferases (DNMTs) are enzymes that catalyse the addition of methyl groups to cytosine residues in DNA. Mammals have three important DNMTs: DNMT1 is responsi‐ ble for the maintenance of existing methylation patterns following DNA replication, while DNMT3a and DNMT3b are *de novo* methyltransferases [1,44-46]. As a result of DNA replica‐ tion, fully methylated DNA becomes hemi-methylated and DNMT1 binds hemi-methylated

Overall, most DNA methylation changes can be observed invariantly in all tissues [47]. However, the small portion of tissue-specific methylation has a profound effect on cellular

and the transacription start and termination regions lacking methylation [40,41].

60 to 70% G/C rich, found in TATAless promoters and/or first exons of genes [17-19].

contribute to the regulation of gene expression [32].

4 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

complete loss during differentiation [40].

DNA to add a methyl group to the 5′ carbon of cytosines [2].

activity including cell differentiation, disease and cancer [48-53].

Key stages in development make use of methylation to switch on/off and regulate gene ex‐ pression. DNA methylation was shown to be essential for embryonic development through homozygous deletion of the mouse Mtase gene which leads to embryonic lethality [52]. Germline cells show 4% less methylation in CGI promoters, including almost all CGI pro‐ moters of germline-specific genes, compared to somatic cells [21].

Immediately after fertilisation but before the first cell division, the paternal DNA undergoes active demethylation throughout the genome [55-58]. After the first cell cycle, the maternal DNA undergoes passive demethylation as a result of a lack of methylation maintenance af‐ ter mitosis [56,59], and this genome-wide demethylation continues, except for the imprinted genes, until the formation of the blastocyst [60,61].

After implantation, the genome (except for CGIs) undergoes *de novo* methylation [54]. Active demethylation subsequently occurs during early embryogenesis [62] with tissue-specific genes undergoing demethylation in their respective tissues, creating a methylation pattern which is maintained in the adult, giving each cell type a unique epigenome. [54].

Somatic cells go through the process of aging as they divide and replicate. Aging is charac‐ terised by a genome-wide loss and a regional gain of DNA methylation [63]. CGI promoters present an increase in DNA methylation in normal tissues of older individuals at several sites throughout the genome [64,65]. This causes genomic instability and deregulation of tis‐ sue-specific and imprinted genes as well as silencing of tumour suppressor genes (control‐ ling cell cycle, apoptosis or DNA repair) through hypermethylation of promoter CGIs [5,66].

The age-related change in methylation was shown in a genome-wide CGI methylation study comparing small intestine (and other tissues) from 3-month-old and 35-month-old mice, which presented linear age-related increased methylation in 21% and decreased methylation in 13% of tested CGIs with strong tissue-specificity [67]. Furthermore, human intestinal agerelated aberrant methylation was shown to share similarities to mouse [67]. Although the majority of CGIs methylated in tumours are also methylated in a selection of normal tissues during aging, particular tumours exhibit methylation in specific promoters and are thus said to display a CpG island methylator phenotype (CIMP) [65].

Aging appears to exhibit common methylation features with carcinogenesis and in fact these processes share a large number of hypermethylated genes such as ER, IGF2, N33 and MyoD in colon cancer, NKX2-5 in prostate cancer and several Polycomb-group protein target genes, which suggests they probably have common epigenetic mechanisms driving them [68-70].

#### **3. Methylation in carcinogenesis**

DNA methylation can either affects key genes which act as a driving force in cancer forma‐ tion or else be a downstream effect of cancer progression [71,72]. According to the widely accepted 'two-hit' hypothesis of carcinogenesis [73], loss of function of both alleles for a giv‐ en gene, such as a tumour suppressor gene, is required for malignant transformation. The first hit is typically in the form of a mutation while the second hit tends to be due to aberrant methylation leading to gene suppression. While in familial cancers only one allele needs to be aberrantly methylated to result in carcinogenesis [74,75], both alleles have to be silenced by methylation in non-familial cancers [76,77]. Interestingly, cancer cells appear to use DNMT3b in addition to DNMT1 to maintain hypermethylation [78,79].

**4. Effect of CpG methylation on transcription factor binding**

GABP [110], NF-κB [111-100], HiNF-P [112] and MSPF [113].

in cells where the gene should not be expressed [102].

the genes involved [114].

activation e.g. STAT proteins) [115].

The methylation of CpGs in transcription factor binding sites in general leads to transcrip‐ tion suppression and gene silencing by directly inhibiting the binding of specific transcrip‐ tion factors. Transcription factors that have CpGs in their recognition sequences and are thus methylation-sensitive include AP-2 [98-100], Ah receptor [101], CREB/ATF [32,100,102], E2F [103], EBP-80 [104], ETS factors [105], MLTF [106], MTF-1 [107], c-Myc, c-Myn [108-109],

Breaking the Silence: The Interplay Between Transcription Factors and DNA Methylation

http://dx.doi.org/10.5772/55021

7

There are also some transcription factors that are not sensitive to methylation e.g. Sp1, CTF and YY1 [100]. Thus methylation does not hinder binding of gene-specific transcription fac‐ tors, but rather prevents the binding of ubiquitous factors, and subsequently transcription,

A model of CpG *de novo* methylation through over-expression of DNMT1 revealed that de‐ spite the overall increase in CGI methylation, there was a differential response of specific sites. The vast majority of CGIs were resistant to *de novo* methylation, while seven novel se‐ quence patterns proved to be particularly susceptible to aberrant methylation [114]. This es‐ sentially means that the sequence in itself plays a role in the methylation state of CGIs. The result of this study implies that specific CGI patterns have an intrinsic susceptibility to aber‐ rant methylation, which means that the genes regulated by promoters containing such CGIs are more susceptible to *de novo* methylation and could lead to various cancers depending on

Various studies have identified three main groups of transcription factors as being impor‐ tant in human cancer: steroid receptors (e.g. oestrogen receptors in breast cancer and andro‐ gen receptors in prostate cancer), resident nuclear factors (always in the nucleus e.g. c-JUN) [115,116] and latent cytoplasmic factors (translocated from the cytopasm to the nucleus after

Resident nuclear proteins are proteins ubiquitously present in the nucleus irrespective of cell type which include bZip proteins e.g. c-JUN, c-FOS, ATFs, CREBs and CREMs, the cEBP fami‐ ly, the ETS proteins and the MAD-box family [117]. The different families vary greatly in over‐ all structure and interaction profiles but have the common functional feature of promoting transcription by co-operating with other transcription factors through tandem recognition se‐ quences in promoters as well as by interacting with co-activator proteins [116,118-124]. Resi‐ dent nuclear transcription factors drive carcinogenesis by direct over-expression or as highly active fusion proteins e.g. MYC acting with MAX [125-127]. The two families of resident nucle‐ ar transcription factors that are most prominent in human cancers are the ETS family proteins and proteins composing the AP-1 transcription complexes. ETS family proteins are of particu‐ lar interest because they promote transcription of a wide range of genes by providing a DNA-

binding domain through fusion with other proteins or by mutation [123,128,129].

Latent cytoplasmic proteins are found in the cytoplasm of cells and rely on protein−protein in‐ teraction at the cell surface to produce a cascade which activates them as they are directed to the nucleus where they affect transcription by binding to activation sites in the promoters of indu‐

Hypermethylation and suppression of promoter CGIs through d*e novo* methylation is welldocumented for numerous cancer, affecting mostly general but occasionally tumour-specific genes [3,4,66,80,81]. A study of over 1000 CGIs from almost 100 human primary tumours de‐ duced that on average 600 CGIs out of an estimated 45,000 spread throughout the genome were aberrantly methylated in cancers. It was shown that while some CGI methylation pat‐ terns were common to all test tumours, others were highly specific to a specific tumourtype, implying that the methylation of certain groups of CGIs may have implications in the formation, malignancy and progression of specific tumour types [82].

CGI shores (the 2kb region at the boundary of CGIs) are methylated in a tissue-specific man‐ ner to regulate gene expression but become hypermethylated in cancer [83-85]. Methylation boundaries flanking the CGIs in the E-cad and VHL tumour suppressor genes were found to be over-ridden by *de novo* methylation, resulting in transcription supression and consequen‐ tially oncogenesis [86]. On the other hand, the location and function of non-CG methylation in cancer is still mostly unknown [87-88].

Aberrant methylation has been linked to cancer cell energetics. Most cancer cells exhibit the Warburg effect i.e. produce energy mainly through a high level of glycolysis followed by lactic acid fermentation in the cytosol even under aerobic conditions, rather than through a low level of glycolysis followed by oxidative phosphorylation in the mitochondria as is the case in normal cells [89].

In one study it was found that fructose-1,6-bisphosphatase-1 (FBP1), which reduces glycoly‐ sis, is down-regulated by the NF-κB pathway partly through hypermethylation of the FBP1 promoter [90]. It was proposed that NF-κB could interact with co-repressors such as Histone deacetylases 1 and 2 (HDAC1 and HDAC2) to suppress gene expression [91,92] and subse‐ quently the HDACs could interact with DNMT1, which gives hypermethylation of the pro‐ moter resulting in gene silencing [93-96].

In another study it was proposed that environmental toxins bring about oxidative-stress which affects genome-wide methylation by activating the Ten-Eleven Translocation (TET) proteins (which convert methylcytosine to 5-hydroxymethylcytosine) and chromatin modi‐ fying proteins which interfere with oxidative phoshphorylation [97].

#### **4. Effect of CpG methylation on transcription factor binding**

**3. Methylation in carcinogenesis**

6 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

in cancer is still mostly unknown [87-88].

moter resulting in gene silencing [93-96].

case in normal cells [89].

DNA methylation can either affects key genes which act as a driving force in cancer forma‐ tion or else be a downstream effect of cancer progression [71,72]. According to the widely accepted 'two-hit' hypothesis of carcinogenesis [73], loss of function of both alleles for a giv‐ en gene, such as a tumour suppressor gene, is required for malignant transformation. The first hit is typically in the form of a mutation while the second hit tends to be due to aberrant methylation leading to gene suppression. While in familial cancers only one allele needs to be aberrantly methylated to result in carcinogenesis [74,75], both alleles have to be silenced by methylation in non-familial cancers [76,77]. Interestingly, cancer cells appear to use

Hypermethylation and suppression of promoter CGIs through d*e novo* methylation is welldocumented for numerous cancer, affecting mostly general but occasionally tumour-specific genes [3,4,66,80,81]. A study of over 1000 CGIs from almost 100 human primary tumours de‐ duced that on average 600 CGIs out of an estimated 45,000 spread throughout the genome were aberrantly methylated in cancers. It was shown that while some CGI methylation pat‐ terns were common to all test tumours, others were highly specific to a specific tumourtype, implying that the methylation of certain groups of CGIs may have implications in the

CGI shores (the 2kb region at the boundary of CGIs) are methylated in a tissue-specific man‐ ner to regulate gene expression but become hypermethylated in cancer [83-85]. Methylation boundaries flanking the CGIs in the E-cad and VHL tumour suppressor genes were found to be over-ridden by *de novo* methylation, resulting in transcription supression and consequen‐ tially oncogenesis [86]. On the other hand, the location and function of non-CG methylation

Aberrant methylation has been linked to cancer cell energetics. Most cancer cells exhibit the Warburg effect i.e. produce energy mainly through a high level of glycolysis followed by lactic acid fermentation in the cytosol even under aerobic conditions, rather than through a low level of glycolysis followed by oxidative phosphorylation in the mitochondria as is the

In one study it was found that fructose-1,6-bisphosphatase-1 (FBP1), which reduces glycoly‐ sis, is down-regulated by the NF-κB pathway partly through hypermethylation of the FBP1 promoter [90]. It was proposed that NF-κB could interact with co-repressors such as Histone deacetylases 1 and 2 (HDAC1 and HDAC2) to suppress gene expression [91,92] and subse‐ quently the HDACs could interact with DNMT1, which gives hypermethylation of the pro‐

In another study it was proposed that environmental toxins bring about oxidative-stress which affects genome-wide methylation by activating the Ten-Eleven Translocation (TET) proteins (which convert methylcytosine to 5-hydroxymethylcytosine) and chromatin modi‐

fying proteins which interfere with oxidative phoshphorylation [97].

DNMT3b in addition to DNMT1 to maintain hypermethylation [78,79].

formation, malignancy and progression of specific tumour types [82].

The methylation of CpGs in transcription factor binding sites in general leads to transcrip‐ tion suppression and gene silencing by directly inhibiting the binding of specific transcrip‐ tion factors. Transcription factors that have CpGs in their recognition sequences and are thus methylation-sensitive include AP-2 [98-100], Ah receptor [101], CREB/ATF [32,100,102], E2F [103], EBP-80 [104], ETS factors [105], MLTF [106], MTF-1 [107], c-Myc, c-Myn [108-109], GABP [110], NF-κB [111-100], HiNF-P [112] and MSPF [113].

There are also some transcription factors that are not sensitive to methylation e.g. Sp1, CTF and YY1 [100]. Thus methylation does not hinder binding of gene-specific transcription fac‐ tors, but rather prevents the binding of ubiquitous factors, and subsequently transcription, in cells where the gene should not be expressed [102].

A model of CpG *de novo* methylation through over-expression of DNMT1 revealed that de‐ spite the overall increase in CGI methylation, there was a differential response of specific sites. The vast majority of CGIs were resistant to *de novo* methylation, while seven novel se‐ quence patterns proved to be particularly susceptible to aberrant methylation [114]. This es‐ sentially means that the sequence in itself plays a role in the methylation state of CGIs. The result of this study implies that specific CGI patterns have an intrinsic susceptibility to aber‐ rant methylation, which means that the genes regulated by promoters containing such CGIs are more susceptible to *de novo* methylation and could lead to various cancers depending on the genes involved [114].

Various studies have identified three main groups of transcription factors as being impor‐ tant in human cancer: steroid receptors (e.g. oestrogen receptors in breast cancer and andro‐ gen receptors in prostate cancer), resident nuclear factors (always in the nucleus e.g. c-JUN) [115,116] and latent cytoplasmic factors (translocated from the cytopasm to the nucleus after activation e.g. STAT proteins) [115].

Resident nuclear proteins are proteins ubiquitously present in the nucleus irrespective of cell type which include bZip proteins e.g. c-JUN, c-FOS, ATFs, CREBs and CREMs, the cEBP fami‐ ly, the ETS proteins and the MAD-box family [117]. The different families vary greatly in over‐ all structure and interaction profiles but have the common functional feature of promoting transcription by co-operating with other transcription factors through tandem recognition se‐ quences in promoters as well as by interacting with co-activator proteins [116,118-124]. Resi‐ dent nuclear transcription factors drive carcinogenesis by direct over-expression or as highly active fusion proteins e.g. MYC acting with MAX [125-127]. The two families of resident nucle‐ ar transcription factors that are most prominent in human cancers are the ETS family proteins and proteins composing the AP-1 transcription complexes. ETS family proteins are of particu‐ lar interest because they promote transcription of a wide range of genes by providing a DNAbinding domain through fusion with other proteins or by mutation [123,128,129].

Latent cytoplasmic proteins are found in the cytoplasm of cells and rely on protein−protein in‐ teraction at the cell surface to produce a cascade which activates them as they are directed to the nucleus where they affect transcription by binding to activation sites in the promoters of indu‐ cible genes and interacting with transcription initiation factors. They can be activated either di‐ rectly by tyrosine or serine kinases at the cell surface or by complex processes which include kinases along the pathway [117]. STATs (signal transducers and activators of transcription) are activated by JAK (a tyrosine kinase family) which is activated by various receptors [130,131].

[142-145], and play a role in malignancy e.g. Sp1 has been shown to be the key factor in epi‐

Breaking the Silence: The Interplay Between Transcription Factors and DNA Methylation

http://dx.doi.org/10.5772/55021

9

Multiple Sp1 binding sites are found in the CGI-promoters of housekeeping genes [148,149] as well as CGIs downstream of the TSS [150]. Sp1 sites in gene promoters have been shown to pro‐ tect CGIs from *de novo* methylation and maintain expression of downstream genes [151,153] e.g. Sp1-binding site protect the APRT gene from *de novo* methylation in humans and mice [154,155]. However, Sp1 binding is not methylation-sensitive [151,156,157] and resistance to *de novo* methylation by DNMT1 is not correlated to the frequency of Sp1 sites in CGIs [114].

Sp1 co-operates with the GABP complex to activate genes which include the folate receptor b [158], CD18 [159], utrophin [141,160], heparanase-1 [161], the pem pd homeobox gene [162], the mouse thymidylate synthase promoter [163] and mouse DNA polymerase alpha

GABP is a transcription factor composed of two distinct subunits: GABPα and GABPβ. GABPα, also known as Nuclear Respiratory Factor 2 (NRF-2) or Adenovirus E4 Transcrip‐ tion Factor 1 (E4TF1-60), is a member of the E26 Transformation-Specific (ETS) family of proteins [166-169]. However unlike other ETS factors GABPα forms an obligate heteromeric protein complex with GABPβ [170-172]. Together they generally form a heterotetramer con‐ sisting of 2α and 2β subunits [173,174] and the presence of sites for GABP binding contain‐ ing 2 tandem ETS consensus motifs has been reported [175]. On the other hand, single GABP binding sites tend to combine with another site that recognises a different transcrip‐ tion factor e.g. NRF-1, Sp1 or YY1 [175]. GABP is able to recruit co-activators such as PCG1

and p300/CBP that create a chromatin environment favouring transcription [176,177].

GABPα (like all other ETS factors) binds to purine-rich sequences containing a 5'- GGAA/ T-3' core by means of a highly conserved DNA-binding domain made up of an 85 amino acid sequence rich in tryptophan which forms a winged-helix-turn-helix structure, charac‐ teristic of the ETS protein family near its carboxy terminal [166,167,170,172,178-181]. The do‐ main through which GABPα binds to the ankyrin repeats of GABPβ is found just downstream of the DNA-binding domain [167,168]. GABPα also has another two domains, the helical bundle pointed (PNT) domain found in its mid-region, which consists of five αhelices [182,183] and the On-SighT (OST) domain near the amino-terminus (residues 35−121), which folds as a 5-stranded β-sheet crossed by a distorted helix and contains two predominant clusters of negatively-charged residues, which might be used to interact with

The role of GABP is very versatile and its ability to co-operate with other transcription fac‐ tors gives it a key role in transcription regulation. GABP and PU.1 compete for binding to the promoter of the b2-integrin gene, yet co-operate to increase gene transcription [185]. GABP also acts as a repressor of mouse ribosomal protein gene transcription [186], appa‐ rently by interfering with the formation of the transcriptional initiation complex [187].

thelial carcinomas [146,147].

primase with E2F [164,165].

**7. GA-Binding Protein (GABP)**

positively-charged proteins [184].

#### **5. Protection mechanisms against methylation**

It has been generally accepted that methylation-resistant CGIs are associated with broad ex‐ pression or housekeeping genes while the majority of methylation-prone CGIs are associat‐ ed with tissue-specific and thus restricted-expression genes [132]. Exceptions to this pattern have also been found, including WNT10B, NPTXR and POP3. Thus the hypothesis that ac‐ tive transcription has an indirect protective effect against aberrant methylation of CGIs [1,133] has been repeatedly proven to be valid though not absolute [114].

A number of mechanisms have been put forward to explain the relationship between aber‐ rant *de novo* methylation and cancer. One hypothesis proposed that an initial random meth‐ ylation event is selected for as proliferation progresses [80]. Another hypothesis proposed the recruitment of DNA methyltransferases to methylation-sensitive sequences by cis-acting factors [134,135], histone methyltransferases such as G9a [136,137], or EZH2 [138]. Yet an‐ other hypothesis proposed the loss of chromatin boundaries or the absence of 'protective' transcription factors, leading to the spread of DNA methylation in CGIs [139].

The most recent hypothesis proposes the protective character of co-operative binding of transcription factors in maintaining CGIs unmethylated [140]. CGIs showed an unexpected resistance to *de novo* methylation when DNMT1 was over-expressed. The general pattern that emerged was that most *de novo* methylated CGIs were characterised by an absence of intandem transcription factor binding sites and an absence of bound transcription factors. Thus protection from *de novo* methylation requires the presence of tandem transcription fac‐ tor binding sites that are stably co-bound by at least one general transcription factor, with the second factor being either a general or a tissue–specific transcription factor. Among the most prominent transcription factors found to be linked with aberrant methylation were GABP, SP1, NFY, NRF1 and YY1 [140].

This study re-confirmed that methylation-resistant CGIs were bound by combinations of ubiquitous transcription factors which regulated genes of basic cellular functions, while methylation-prone CGIs were mostly associated with development, differentiation and cell communication, which are frequently regulated by tissue–specific transcription factors [140].

#### **6. Specificity protein 1 (Sp1)**

Sp1 is an Sp/KLF (Krüppel-like factor) family member containing a zinc-finger DNA-bind‐ ing domain [141]. Many KLF proteins regulate cellular proliferation and differentiation [142-145], and play a role in malignancy e.g. Sp1 has been shown to be the key factor in epi‐ thelial carcinomas [146,147].

Multiple Sp1 binding sites are found in the CGI-promoters of housekeeping genes [148,149] as well as CGIs downstream of the TSS [150]. Sp1 sites in gene promoters have been shown to pro‐ tect CGIs from *de novo* methylation and maintain expression of downstream genes [151,153] e.g. Sp1-binding site protect the APRT gene from *de novo* methylation in humans and mice [154,155]. However, Sp1 binding is not methylation-sensitive [151,156,157] and resistance to *de novo* methylation by DNMT1 is not correlated to the frequency of Sp1 sites in CGIs [114].

Sp1 co-operates with the GABP complex to activate genes which include the folate receptor b [158], CD18 [159], utrophin [141,160], heparanase-1 [161], the pem pd homeobox gene [162], the mouse thymidylate synthase promoter [163] and mouse DNA polymerase alpha primase with E2F [164,165].

#### **7. GA-Binding Protein (GABP)**

cible genes and interacting with transcription initiation factors. They can be activated either di‐ rectly by tyrosine or serine kinases at the cell surface or by complex processes which include kinases along the pathway [117]. STATs (signal transducers and activators of transcription) are activated by JAK (a tyrosine kinase family) which is activated by various receptors [130,131].

It has been generally accepted that methylation-resistant CGIs are associated with broad ex‐ pression or housekeeping genes while the majority of methylation-prone CGIs are associat‐ ed with tissue-specific and thus restricted-expression genes [132]. Exceptions to this pattern have also been found, including WNT10B, NPTXR and POP3. Thus the hypothesis that ac‐ tive transcription has an indirect protective effect against aberrant methylation of CGIs

A number of mechanisms have been put forward to explain the relationship between aber‐ rant *de novo* methylation and cancer. One hypothesis proposed that an initial random meth‐ ylation event is selected for as proliferation progresses [80]. Another hypothesis proposed the recruitment of DNA methyltransferases to methylation-sensitive sequences by cis-acting factors [134,135], histone methyltransferases such as G9a [136,137], or EZH2 [138]. Yet an‐ other hypothesis proposed the loss of chromatin boundaries or the absence of 'protective'

The most recent hypothesis proposes the protective character of co-operative binding of transcription factors in maintaining CGIs unmethylated [140]. CGIs showed an unexpected resistance to *de novo* methylation when DNMT1 was over-expressed. The general pattern that emerged was that most *de novo* methylated CGIs were characterised by an absence of intandem transcription factor binding sites and an absence of bound transcription factors. Thus protection from *de novo* methylation requires the presence of tandem transcription fac‐ tor binding sites that are stably co-bound by at least one general transcription factor, with the second factor being either a general or a tissue–specific transcription factor. Among the most prominent transcription factors found to be linked with aberrant methylation were

This study re-confirmed that methylation-resistant CGIs were bound by combinations of ubiquitous transcription factors which regulated genes of basic cellular functions, while methylation-prone CGIs were mostly associated with development, differentiation and cell communication, which are frequently regulated by tissue–specific transcription factors [140].

Sp1 is an Sp/KLF (Krüppel-like factor) family member containing a zinc-finger DNA-bind‐ ing domain [141]. Many KLF proteins regulate cellular proliferation and differentiation

**5. Protection mechanisms against methylation**

8 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

GABP, SP1, NFY, NRF1 and YY1 [140].

**6. Specificity protein 1 (Sp1)**

[1,133] has been repeatedly proven to be valid though not absolute [114].

transcription factors, leading to the spread of DNA methylation in CGIs [139].

GABP is a transcription factor composed of two distinct subunits: GABPα and GABPβ. GABPα, also known as Nuclear Respiratory Factor 2 (NRF-2) or Adenovirus E4 Transcrip‐ tion Factor 1 (E4TF1-60), is a member of the E26 Transformation-Specific (ETS) family of proteins [166-169]. However unlike other ETS factors GABPα forms an obligate heteromeric protein complex with GABPβ [170-172]. Together they generally form a heterotetramer con‐ sisting of 2α and 2β subunits [173,174] and the presence of sites for GABP binding contain‐ ing 2 tandem ETS consensus motifs has been reported [175]. On the other hand, single GABP binding sites tend to combine with another site that recognises a different transcrip‐ tion factor e.g. NRF-1, Sp1 or YY1 [175]. GABP is able to recruit co-activators such as PCG1 and p300/CBP that create a chromatin environment favouring transcription [176,177].

GABPα (like all other ETS factors) binds to purine-rich sequences containing a 5'- GGAA/ T-3' core by means of a highly conserved DNA-binding domain made up of an 85 amino acid sequence rich in tryptophan which forms a winged-helix-turn-helix structure, charac‐ teristic of the ETS protein family near its carboxy terminal [166,167,170,172,178-181]. The do‐ main through which GABPα binds to the ankyrin repeats of GABPβ is found just downstream of the DNA-binding domain [167,168]. GABPα also has another two domains, the helical bundle pointed (PNT) domain found in its mid-region, which consists of five αhelices [182,183] and the On-SighT (OST) domain near the amino-terminus (residues 35−121), which folds as a 5-stranded β-sheet crossed by a distorted helix and contains two predominant clusters of negatively-charged residues, which might be used to interact with positively-charged proteins [184].

The role of GABP is very versatile and its ability to co-operate with other transcription fac‐ tors gives it a key role in transcription regulation. GABP and PU.1 compete for binding to the promoter of the b2-integrin gene, yet co-operate to increase gene transcription [185]. GABP also acts as a repressor of mouse ribosomal protein gene transcription [186], appa‐ rently by interfering with the formation of the transcriptional initiation complex [187].

GABP is a methylation-sensitive transcription factor [110] and its modulation is best seen in the transactivation of the *Cyp*2d-9 promoter for the male-specific steroid 16a-hydroxylase in mouse liver where GABP does not bind to the promoter when the CpG site at -97 is methylated [187]. Interestingly, CpG sites located at -93 and -85, outside of the GABP recognition sequence in the Thyroid Stimulating Hormone Receptor (TSHR) gene promoter when methylated, affect the binding of GABP to the promoter, leading to a reduction in basal transcription [187].

**9. Conclusion**

in protein-targetting treatments.

Address all correspondence to: angenlabs@gmail.com

School of Medicine, Ube-shi, Yamaguchi-ken, Japan

cinogenesis 2000: 21, 461–467.

ty goes awry, Dev. Cell 2010: 19, 698-711.

**Author details**

Malta, Msida, Malta

Byron Baron1,2

**References**

21

763–770.

Research into DNA methylation, particularly at CGIs has come a long way and it is now known that gene silencing, albeit essential, is not the only purpose of methylation processes. In particular, the interactions of transcription factors with promoters have been shown to modulate the function of genes through their methylation-sensitivity and may thus be re‐ garded as viable targets for therapeutics. Unfortunately the biochemical mechanisms and principles required to successfully inhibit protein–protein interactions require further study and clarification [203-206]. Additionally, delivery systems for such cellular treatments also need further study and improvement. However as more focus is put on molecular medicine and with the shift towards personalised medicine, there will surely be significant advances

Breaking the Silence: The Interplay Between Transcription Factors and DNA Methylation

http://dx.doi.org/10.5772/55021

11

1 Department of Anatomy and Cell Biology, Faculty of Medicine and Surgery, University of

2 Department of Biochemistry and Functional Proteomics, Yamaguchi University Graduate

[1] Bird A. DNA methylation patterns and epigenetic memory. Genes Dev. 2002: 16, 6–

[2] Chatterjee R, Vinson C. CpG methylation recruits sequence specific transcription fac‐ tors essential for tissue specific gene expression. Biochim. Biophys. Acta 2012: 1819,

[3] Robertson KD, Jones PA. DNA methylation: past, present and future directions. Car‐

[4] Esteller M. CpG island hypermethylation and tumour suppressor genes: a booming

[5] Berdasco M, Esteller M. Aberrant epigenetic landscape in cancer: how cellular identi‐

present, a brighter future. Oncogene 2002: 21, 5427–5440.

#### **8. Therapeutic applications**

As more such data is accumulated, it presents methylation as a very interesting and promis‐ ing tumour-specific therapeutic target especially since the lack of methylation of CGIs in normal cells makes it a safe therapy. Demethylation is known to reactivate the expression of many genes silenced in cultured tumour cells [82]. While high doses of DNMT inhibitors can inhibit DNA synthesis and eventually lead to cell death by cytotoxicity, administration of low doses of these drugs over a prolonged period has a therapeutic effect [188-191]. In fact, the United States Food and Drug Administration has approved the DNMT inhibitors, 5-azacytidine and its derivative 5-aza-2′-deoxycytidine (decitabine), for therapy of patients with solid tumours, myelodysplastic syndrome (which can lead to the development of acute leukemia) and myelogenous leukemia [192].

5-azacitidine acts by becoming phosphorylated and being incorporated into RNA, where it suppresses RNA synthesis and produces a cytotoxic effect [3,193]. It is converted by ribonu‐ cleotide reductase to 5-aza-2'-deoxycytidine diphosphate and subsequently phosphorylated. The triphosphate form is then incorporated into DNA in place of cytosine. The substitution of the 5' nitrogen atom in place of the carbon, traps the DNMTs on the substituted DNA strand and methylation is inhibited [194].

Several more stable analogues such as arabinofuranosyl-5-azacytosine [195], pseudo-isocyti‐ dine [196], 5-fluorocytidine [196], pyrimidone [197] and dihydro-5-azacytidine [198] have been tested, and others are undergoing clinical trials [199,200].

Targetting overactive transcription factors is another interesting tumour-specific therapeu‐ tic strategy. Many human cancers appear to have a small number of specific overactive transcription factors which are valid candidate targets to at least control further malignan‐ cy and metastasis. Such tumour-specific transcription factors are ideal targets because they are less numerous and more significant than other possible protein targets in the tran‐ scription activation pathway.

However it is not a simple task to target transcription factors in a controlled manner particu‐ larly if attempting to inhibit the interaction of DNA-binding proteins with their recognition sequences [201,202]. Inhibition of a DNA-binding transcription factor can alternatively be done in one of two ways: lowering the overall level of intracellular transcription factor through siRNA or directing methylation to the recognition sequence of the DNA-binding protein. Both options are extremely difficult to carry out *in vivo* even if their *in vitro* counter‐ part has proven to be successful.

#### **9. Conclusion**

GABP is a methylation-sensitive transcription factor [110] and its modulation is best seen in the transactivation of the *Cyp*2d-9 promoter for the male-specific steroid 16a-hydroxylase in mouse liver where GABP does not bind to the promoter when the CpG site at -97 is methylated [187]. Interestingly, CpG sites located at -93 and -85, outside of the GABP recognition sequence in the Thyroid Stimulating Hormone Receptor (TSHR) gene promoter when methylated, affect

As more such data is accumulated, it presents methylation as a very interesting and promis‐ ing tumour-specific therapeutic target especially since the lack of methylation of CGIs in normal cells makes it a safe therapy. Demethylation is known to reactivate the expression of many genes silenced in cultured tumour cells [82]. While high doses of DNMT inhibitors can inhibit DNA synthesis and eventually lead to cell death by cytotoxicity, administration of low doses of these drugs over a prolonged period has a therapeutic effect [188-191]. In fact, the United States Food and Drug Administration has approved the DNMT inhibitors, 5-azacytidine and its derivative 5-aza-2′-deoxycytidine (decitabine), for therapy of patients with solid tumours, myelodysplastic syndrome (which can lead to the development of acute

5-azacitidine acts by becoming phosphorylated and being incorporated into RNA, where it suppresses RNA synthesis and produces a cytotoxic effect [3,193]. It is converted by ribonu‐ cleotide reductase to 5-aza-2'-deoxycytidine diphosphate and subsequently phosphorylated. The triphosphate form is then incorporated into DNA in place of cytosine. The substitution of the 5' nitrogen atom in place of the carbon, traps the DNMTs on the substituted DNA

Several more stable analogues such as arabinofuranosyl-5-azacytosine [195], pseudo-isocyti‐ dine [196], 5-fluorocytidine [196], pyrimidone [197] and dihydro-5-azacytidine [198] have

Targetting overactive transcription factors is another interesting tumour-specific therapeu‐ tic strategy. Many human cancers appear to have a small number of specific overactive transcription factors which are valid candidate targets to at least control further malignan‐ cy and metastasis. Such tumour-specific transcription factors are ideal targets because they are less numerous and more significant than other possible protein targets in the tran‐

However it is not a simple task to target transcription factors in a controlled manner particu‐ larly if attempting to inhibit the interaction of DNA-binding proteins with their recognition sequences [201,202]. Inhibition of a DNA-binding transcription factor can alternatively be done in one of two ways: lowering the overall level of intracellular transcription factor through siRNA or directing methylation to the recognition sequence of the DNA-binding protein. Both options are extremely difficult to carry out *in vivo* even if their *in vitro* counter‐

the binding of GABP to the promoter, leading to a reduction in basal transcription [187].

**8. Therapeutic applications**

10 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

leukemia) and myelogenous leukemia [192].

strand and methylation is inhibited [194].

scription activation pathway.

part has proven to be successful.

been tested, and others are undergoing clinical trials [199,200].

Research into DNA methylation, particularly at CGIs has come a long way and it is now known that gene silencing, albeit essential, is not the only purpose of methylation processes. In particular, the interactions of transcription factors with promoters have been shown to modulate the function of genes through their methylation-sensitivity and may thus be re‐ garded as viable targets for therapeutics. Unfortunately the biochemical mechanisms and principles required to successfully inhibit protein–protein interactions require further study and clarification [203-206]. Additionally, delivery systems for such cellular treatments also need further study and improvement. However as more focus is put on molecular medicine and with the shift towards personalised medicine, there will surely be significant advances in protein-targetting treatments.

#### **Author details**

Byron Baron1,2

Address all correspondence to: angenlabs@gmail.com

1 Department of Anatomy and Cell Biology, Faculty of Medicine and Surgery, University of Malta, Msida, Malta

2 Department of Biochemistry and Functional Proteomics, Yamaguchi University Graduate School of Medicine, Ube-shi, Yamaguchi-ken, Japan

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Breaking the Silence: The Interplay Between Transcription Factors and DNA Methylation

http://dx.doi.org/10.5772/55021

13

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**Section 2**

**DNA-Methyltransferases: Structure and**

**Function in Eukaryotic and Prokaryotic System**


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**Chapter 2**

**Diverse Domains of (Cytosine-5)-DNA**

**Characterization**

http://dx.doi.org/10.5772/52046

Watson–Crick interactions [1].

**1. Introduction**

A. Yu. Ryazanova, L. A. Abrosimova, T. S. Oretskaya and E. A. Kubareva

Additional information is available at the end of the chapter

**Methyltransferases: Structural and Functional**

(Cytosine-5)-DNA methyltransferases (C5-DNA MTases) are enzymes which catalyze meth‐ yl group transfer from *S*-adenosyl-*L*-methionine (AdoMet) to C5 atom of cytosine residue in DNA. As a result, AdoMet is converted into *S*-adenosyl-*L*-homocysteine (AdoHcy). The rec‐ ognition sites of C5-DNA MTases are usually short palindromic sequences (2–6 bp) in dou‐ ble-stranded DNA. One or both DNA strands can be methylated. The introduced methyl group is localized in the major groove of the DNA double helix and thus does not disrupt

In prokaryotes, DNA methylation underlies several important processes, *e.g.* host and for‐ eign DNA distinction as well as maternal and daughter strand discrimination that is vital for correction of replication errors in the newly synthesized DNA strand. DNA methylation is also responsible for DNA replication control and its interconnection with cell cycle [1]. The majority of the known DNA MTases are components of restriction–modification (R–M) sys‐ tems which protect bacterial cells from bacteriophage infection. A typical R–M system con‐ sists of a MTase which modifies certain DNA sequences and a restriction endonuclease (RE)

In eukaryotes, DNA methylation has diverse functions such as control of gene expression, regulation of genome imprinting, X-chromosome inactivation, genome defense from endog‐

and reproduction in any medium, provided the original work is properly cited.

© 2013 Ryazanova et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

which hydrolyses DNA if these sequences remain unmethylated [2].

**Chapter 2**

## **Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization**

A. Yu. Ryazanova, L. A. Abrosimova, T. S. Oretskaya and E. A. Kubareva

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52046

#### **1. Introduction**

(Cytosine-5)-DNA methyltransferases (C5-DNA MTases) are enzymes which catalyze meth‐ yl group transfer from *S*-adenosyl-*L*-methionine (AdoMet) to C5 atom of cytosine residue in DNA. As a result, AdoMet is converted into *S*-adenosyl-*L*-homocysteine (AdoHcy). The rec‐ ognition sites of C5-DNA MTases are usually short palindromic sequences (2–6 bp) in dou‐ ble-stranded DNA. One or both DNA strands can be methylated. The introduced methyl group is localized in the major groove of the DNA double helix and thus does not disrupt Watson–Crick interactions [1].

In prokaryotes, DNA methylation underlies several important processes, *e.g.* host and for‐ eign DNA distinction as well as maternal and daughter strand discrimination that is vital for correction of replication errors in the newly synthesized DNA strand. DNA methylation is also responsible for DNA replication control and its interconnection with cell cycle [1]. The majority of the known DNA MTases are components of restriction–modification (R–M) sys‐ tems which protect bacterial cells from bacteriophage infection. A typical R–M system con‐ sists of a MTase which modifies certain DNA sequences and a restriction endonuclease (RE) which hydrolyses DNA if these sequences remain unmethylated [2].

In eukaryotes, DNA methylation has diverse functions such as control of gene expression, regulation of genome imprinting, X-chromosome inactivation, genome defense from endog‐

© 2013 Ryazanova et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

enous retroviruses and transposons, participation in development of immune system and in brain functioning. Anomalous methylation patterns in humans are associated with psycho‐ ses, immune system diseases and different cancers [3].

**Domain name and number Domain description**

Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization

http://dx.doi.org/10.5772/52046

31

MethyltransfD12 (PF02086) D12 class N6-adenine-specific DNA methyltransferase

**DNA\_methylase (PF00145)** C5-cytosine-specific DNA methyltransferase Eco57I (PF07669) Eco57I restriction–modification methyltransferase

Methyltransf\_26 (PF13659) Methyltransferase domain

N6\_Mtase (PF02384) N6-DNA methyltransferase N6\_N4\_Mtase (PF01555) DNA methyltransferase

Cons\_hypoth95 (PF03602) Conserved hypothetical protein 95 EcoRI\_methylase (PF13651) Adenine-specific methyltransferase EcoRI Dam (PF05869) DNA N6-adenine-methyltransferase (Dam)

RE\_Eco47II (PF09553) Eco47II restriction endonuclease RE\_HaeII (PF09554) HaeII restriction endonuclease RE\_HaeIII (PF09556) HaeIII restriction endonuclease RE\_HpaII (PF09561) HpaII restriction endonuclease

Vsr (PF03852) DNA mismatch endonuclease Vsr DUF559 (PF04480) Domain of unknown function

BsuBI\_PstI\_RE (PF06616) BsuBI/PstI restriction endonuclease C-terminus

TaqI\_C (PF12950) TaqI-like C-terminal specificity domain

HTH\_26 (PF13443) Cro/C1-type HTH DNA-binding domain

HNH\_3(PF13392) HNH endonuclease

HTH\_3 (PF01381) Helix–turn–helix (HTH)

HTH\_23 (PF13384) Homeodomain-like domain

MerR (PF00376) MerR family regulatory protein MerR\_1 (PF13411) MerR HTH family regulatory protein DUF1870 (PF08965) Domain of unknown function PHD (PF00628) PHD-finger (plant homeo domain)

SNF2\_N (PF00176) SNF2 family N-terminal domain

HTH\_17 (PF12728) HTH HTH\_19 (PF12844) HTH

HTH\_31 (PF13560) HTH

**Domains of other DNA-operating enzymes**

**Methyltransferase domains**

**Endonuclease domains**

**Transcription regulators**

Different C5-DNA MTases share high similarity both in the primary and in the tertiary structure that enables their easy identification by bioinformatic tools. At the moment (July 2012), the Pfam database (http://pfam.sanger.ac.uk/) contains 5065 protein sequences that possess the characteristic domain of C5-DNA MTases (PF00145). Among this moiety, 3072 sequences (61%) contain the only domain PF00145 while the others have a duplication of this domain and/or other additional domains. The diversity of such domains fused in a sin‐ gle polypeptide with the C5-DNA MTase domain is rather wide (Table 1): there are MTases, RE, transcription factors, chromatin-associated domains *etc*.

To date, the structural characteristics of the catalytic domain from different MTases, the de‐ tails of the catalytic mechanism and the biological functions of C5-DNA MTases from differ‐ ent organisms are summarized in a variety of reviews (for example, see [1, 4-8]). Therefore, these aspects are discussed here rather briefly. The present review is focused on the addi‐ tional activities of C5-DNA MTases, on the structure and functions of the domains which are additional to the catalytic one. The data about C5-DNA MTases have not yet been sum‐ marized from this point of view.

### **2. The methyltransferase domain in prokaryotic and eukaryotic (cytosine-5)-DNA methyltransferases**

The most studied enzyme among the prokaryotic C5-DNA MTases is MTase HhaI (M.HhaI) from *Haemophilus haemolyticus*. It methylates the inner cytosine residue in the sequence 5′- G*C*GC-3′/3′-CG*C*G-5′ (italicised). M.HhaI consists of only the MTase domain (Figure 1). The structural organization and catalytic mechanism of C5-DNA MTases were extensively stud‐ ied using this enzyme as a model.

The catalytic domain of C5-DNA MTases consists of two subdomains, a large one and a small one, separated by a DNA-binding cleft. The tertiary structure of the large (catalytic) subdomain has a common structural core – a β-sheet that consists of 7 β-strands and is flanked by 3 α-helices from each side. Six of seven β-strands have a parallel orientation, while the 7th β-strand is located between the 5th and the 6th β-strands in an antiparallel orien‐ tation (Figure 1, b). Thus, the large subdomain consists of 2 parts: the first one (β1–β3) forms the AdoMet binding site while the second one (β4–β7) forms the binding site for the target cytosine. The small subdomain contains a TRD region (target recognition domain) that has a unique sequence in each MTase and is responsible for the substrate specificity. The small subdomains of C5-DNA MTases vary substantially in size and spatial structure [1].

enous retroviruses and transposons, participation in development of immune system and in brain functioning. Anomalous methylation patterns in humans are associated with psycho‐

Different C5-DNA MTases share high similarity both in the primary and in the tertiary structure that enables their easy identification by bioinformatic tools. At the moment (July 2012), the Pfam database (http://pfam.sanger.ac.uk/) contains 5065 protein sequences that possess the characteristic domain of C5-DNA MTases (PF00145). Among this moiety, 3072 sequences (61%) contain the only domain PF00145 while the others have a duplication of this domain and/or other additional domains. The diversity of such domains fused in a sin‐ gle polypeptide with the C5-DNA MTase domain is rather wide (Table 1): there are MTases,

To date, the structural characteristics of the catalytic domain from different MTases, the de‐ tails of the catalytic mechanism and the biological functions of C5-DNA MTases from differ‐ ent organisms are summarized in a variety of reviews (for example, see [1, 4-8]). Therefore, these aspects are discussed here rather briefly. The present review is focused on the addi‐ tional activities of C5-DNA MTases, on the structure and functions of the domains which are additional to the catalytic one. The data about C5-DNA MTases have not yet been sum‐

**2. The methyltransferase domain in prokaryotic and eukaryotic**

The most studied enzyme among the prokaryotic C5-DNA MTases is MTase HhaI (M.HhaI) from *Haemophilus haemolyticus*. It methylates the inner cytosine residue in the sequence 5′- G*C*GC-3′/3′-CG*C*G-5′ (italicised). M.HhaI consists of only the MTase domain (Figure 1). The structural organization and catalytic mechanism of C5-DNA MTases were extensively stud‐

The catalytic domain of C5-DNA MTases consists of two subdomains, a large one and a small one, separated by a DNA-binding cleft. The tertiary structure of the large (catalytic) subdomain has a common structural core – a β-sheet that consists of 7 β-strands and is flanked by 3 α-helices from each side. Six of seven β-strands have a parallel orientation, while the 7th β-strand is located between the 5th and the 6th β-strands in an antiparallel orien‐ tation (Figure 1, b). Thus, the large subdomain consists of 2 parts: the first one (β1–β3) forms the AdoMet binding site while the second one (β4–β7) forms the binding site for the target cytosine. The small subdomain contains a TRD region (target recognition domain) that has a unique sequence in each MTase and is responsible for the substrate specificity. The small

subdomains of C5-DNA MTases vary substantially in size and spatial structure [1].

ses, immune system diseases and different cancers [3].

30 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

RE, transcription factors, chromatin-associated domains *etc*.

marized from this point of view.

ied using this enzyme as a model.

**(cytosine-5)-DNA methyltransferases**



(a) (b)

man numerals.

[14-23] (Figure 2).

**Figure 1.** *a*) Secondary and tertiary structure typical for the C5-DNA MTase domain (M.HhaI in complex with DNA and AdoHcy, PDB code: 3mht). α-Helices are depicted in pink, β-strands are in yellow, DNA is in blue. AdoHcy is in black, spacefill representation. (*b*) The structural core and the order of the conservative motifs in the large subdomain of C5- DNA MTases. α-Helices are depicted as boxes, β-strands – as arrows. The conservative motifs are numbered using Ro‐

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Mammalian DNA MTase Dnmt1 is responsible for maintenance methylation. Its main recog‐ nition site is monomethylated 5′-CG-3′/3′-GC-5′ DNA fragment. The structures of M.HhaI and Dnmt1 in their complexes with DNA were compared [9]. Their catalytic subdomains are rather similar (the root mean square deviation of Cα atoms is 2.0 Å over 218 aligned resi‐ dues). However, the TRD primary and tertiary structures differ significantly between Dnmt1 and M.HhaI. The larger part of the Dnmt1 TRD is structurally isolated and stabilized by Zn2+ ion, the latter one being coordinated by three Cys and one His residues. A β-hairpin in the C-terminal part of the Dnmt1 TRD forms hydrophobic contacts with the catalytic sub‐ domain and the BAH1 domain (see section 4.1.5). The side chains of a few residues (presum‐ ably arginine) in the catalytic subdomain make contacts with the phosphate groups that flank unmethylated 5′-CG-3′/3′-GC-5′ sites. In the M.HhaI structure, the DNA is located in the cleft between the catalytic subdomain and the TRD, whereas the DNA in Dnmt1 com‐ plex is distant from the Dnmt1 catalytic center. This is likely to be connected with the fact that the activity of the Dnmt1 catalytic domain is regulated by the N-terminal part of the

To catalyze the methylation reaction, a MTase binds DNA containing its recognition site and the cofactor AdoMet. Specific DNA–protein contacts are formed between the MTase and heterocyclic bases of the recognition site except the cytosine to be methylated. The target cy‐ tosine is methylated according to SN2 mechanism. The whole methylation process can be divided into 3 steps: a cytosine flipping out of the DNA double helix, a formation of a cova‐ lent enzyme–substrate intermediate and a methyl group transfer to the cytosine residue

protein. An isolated Dnmt1 catalytic domain proved to be inactive [10-13].

**3. The mechanism of DNA methylation**

**Table 1.** Domains existing in a single polypeptide chain with the C5-DNA MTase domain (PF00145) according to the Pfam database.

Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization http://dx.doi.org/10.5772/52046 33

**Figure 1.** *a*) Secondary and tertiary structure typical for the C5-DNA MTase domain (M.HhaI in complex with DNA and AdoHcy, PDB code: 3mht). α-Helices are depicted in pink, β-strands are in yellow, DNA is in blue. AdoHcy is in black, spacefill representation. (*b*) The structural core and the order of the conservative motifs in the large subdomain of C5- DNA MTases. α-Helices are depicted as boxes, β-strands – as arrows. The conservative motifs are numbered using Ro‐ man numerals.

Mammalian DNA MTase Dnmt1 is responsible for maintenance methylation. Its main recog‐ nition site is monomethylated 5′-CG-3′/3′-GC-5′ DNA fragment. The structures of M.HhaI and Dnmt1 in their complexes with DNA were compared [9]. Their catalytic subdomains are rather similar (the root mean square deviation of Cα atoms is 2.0 Å over 218 aligned resi‐ dues). However, the TRD primary and tertiary structures differ significantly between Dnmt1 and M.HhaI. The larger part of the Dnmt1 TRD is structurally isolated and stabilized by Zn2+ ion, the latter one being coordinated by three Cys and one His residues. A β-hairpin in the C-terminal part of the Dnmt1 TRD forms hydrophobic contacts with the catalytic sub‐ domain and the BAH1 domain (see section 4.1.5). The side chains of a few residues (presum‐ ably arginine) in the catalytic subdomain make contacts with the phosphate groups that flank unmethylated 5′-CG-3′/3′-GC-5′ sites. In the M.HhaI structure, the DNA is located in the cleft between the catalytic subdomain and the TRD, whereas the DNA in Dnmt1 com‐ plex is distant from the Dnmt1 catalytic center. This is likely to be connected with the fact that the activity of the Dnmt1 catalytic domain is regulated by the N-terminal part of the protein. An isolated Dnmt1 catalytic domain proved to be inactive [10-13].

#### **3. The mechanism of DNA methylation**

Helicase\_C (PF00271) Helicase conserved C-terminal domain

MutH (PF02976) DNA mismatch repair enzyme MutH

DYW\_deaminase (PF14432) DYW family of nucleic acid deaminases

Transposase\_20 (PF02371) Transposase IS116/IS110/IS902 family

Chromo (PF00385) CHRromatin Organisation MOdifier

DMAP\_binding (PF06464) DMAP1-binding domain

zf-CXXC (PF02008) CXXC zinc finger domain

Pkinase\_Tyr (PF07714) Protein tyrosine kinase YTH (PF04146) YT521-B-like domain

AOX (PF01786) Alternative oxidase

Cullin (PF00888) Cullin family

Hint\_2 (PF13403) Hint domain

PPR (PF01535) PPR repeat (pentatricopeptide repeat)

PALP (PF00291) Pyridoxal-phosphate dependent enzyme

CH (PF00307) Calponin homology (CH) domain Dabb (PF07876) Stress responsive A/B barrel domain

DUF1152 (PF06626) Domain of unknown function DUF3444 (PF11926) Domain of unknown function

Cyt-b5 (PF00173) Cytochrome b5-like heme/steroid binding domain

**Table 1.** Domains existing in a single polypeptide chain with the C5-DNA MTase domain (PF00145) according to the

DNA\_pol3\_beta\_2 (PF02767) DNA polymerase III beta subunit, central domain DNA\_pol3\_beta\_3 (PF02768) DNA polymerase III beta subunit, C-terminal domain HhH-GPD (PF00730) HhH-GPD superfamily base excision DNA repair protein

PWWP (PF00855) PWWP domain (conserved Pro-Trp-Trp-Pro motif)

DNMT1-RFD (PF12047) Cytosine specific DNA methyltransferase replication foci domain

RCC1\_2 (PF13540) Regulator of chromosome condensation (RCC1) repeat

BAH (PF01426) BAH domain (bromo-adjacent homology)

RVT\_1 (PF00078) Reverse transcriptase (RNA-dependent DNA polymerase)

Terminase\_6 (PF03237) Terminase-like family

32 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

DEDD\_Tnp\_IS110 (PF01548) Transposase

**Chromatin-associated domains**

**Others**

Pfam database.

To catalyze the methylation reaction, a MTase binds DNA containing its recognition site and the cofactor AdoMet. Specific DNA–protein contacts are formed between the MTase and heterocyclic bases of the recognition site except the cytosine to be methylated. The target cy‐ tosine is methylated according to SN2 mechanism. The whole methylation process can be divided into 3 steps: a cytosine flipping out of the DNA double helix, a formation of a cova‐ lent enzyme–substrate intermediate and a methyl group transfer to the cytosine residue [14-23] (Figure 2).

VI) protonates the cytosine N3 atom thus facilitating the nucleophilic attack by the thiol group. In the conjugate, the negative charge of the cytosine C5 atom causes its alkylation by the Ado‐ Met methyl group. Proton elimination from the C5 atom of the methylated cytosine and βelimination of the Cys residue result in a breakdown of the covalent DNA–protein complex and restore aromaticity of the modified cytosine base (Figure 2, a). The rate constant for meth‐ yl group transfer catalyzed by M.HhaI is about 0.14–0.26 s–1 [17, 18]. The following release of

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35

C5-DNA MTases HhaI and HpaII catalyze the methylation reaction in a distributive man‐ ner, *i.e.* dissociate from the DNA substrate after every catalytic act [1, 18]. Each one of these MTases is a component of a R–M system where the cognate RE searches for its recognition sites *via* scanning DNA by a linear diffusion mechanism. The MTase distributivity provides for the corresponding RE a possibility to bind an unmodified site before the MTase and to cleave it. On the other hand, C5-DNA MTase SssI [26] has no cognate RE and is able to methylate several recognition sites located in one DNA substrate in a processive manner, *i.e.*

The affinity of prokaryotic C5-DNA MTases to their DNA ligands in the presence of non‐ reactive cofactor analogs increases in the following manner: dimethylated DNA << unme‐ thylated DNA < monomethylated DNA [1, 2, 27, 28]. A similar correlation is observed for the eukaryotic enzyme Dnmt1. The Dnmt1 affinity to monomethylated 5′-CG-3′/3′-GC-5′ sites is 2–200 times higher than to unmethylated ones depending on the experimental condi‐ tions [13, 29-35]. Dnmt1 modifies monomethylated sites processively (more than 50 sites per one binding act on average). Unmethylated sites (mainly 5′-CCGG-3′) are methylated by

Mammalian C5-DNA MTases Dnmt3a and Dnmt3b methylate presumably unmodified sites in DNA and are responsible for *de novo* DNA methylation during embryonic development. Dinucleotides 5'-CG-3'/3′-GC-5′ are the main recognition sites of Dnmt3a and Dnmt3b. These enzymes are also able to modify dinucleotides 5′-CА-3′ but 10–100 times less efficiently [37]. The efficiency of methylation by Dnmt3a and Dnmt3b is also dependent on the sequences flanking the recognition site: 5′-RCGY-3′ is the most frequently methylated site whereas 5′- YCGR-3′ is methylated with a lower efficiency (R and Y are purine and pyrimidine nucleo‐ sides respectively). The difference in the methylation rate of these sites can exceed 500 times [38]. The primary structures of the Dnmt3a and Dnmt3b catalytic domains share identity of 84%. In contrast to Dnmt1, isolated catalytic domains of Dnmt3a and Dnmt3b retain their ac‐ tivity [12, 39]. Dnmt3a methylates DNA in a distributive manner while Dnmt3b modifies DNA processively since the DNA binding center of Dnmt3b is more positively charged than DNA binding center of Dnmt3a [37, 39]. Interestingly, a Cys residue substitution in the catalytic ProCys motif of an isolated Dnmt3a catalytic domain does not totally abolish the enzyme ac‐ tivity but merely decreases it 2–6 times. This Cys residue is shown to take part in the DNA– Dnmt3a conjugate formation. However, the Dnmt3a catalytic domain looses its activity com‐ pletely after a substitution of a Glu residue in the GluAsnVal motif. The active center of Dnmt3a seems to have an unusual conformation and the Cys residue perhaps does not have its optimal orientation. Therefore, the nucleophilic attack onto the cytosine C6 atom can be

the reaction products is a rate-limiting step of the M.HhaI catalytic cycle [17, 18].

without dissociation from DNA after each catalytic act.

Dnmt1 in a distributive manner [36].

**Figure 2.** *a*) Mechanism of cytosine C5 atom methylation by a C5-DNA MTase. (*b*) Mechanism of cytosine deamination as a possible side reaction of the methylation process.

The target nucleotide flipping can occur spontaneously or with the help of an enzyme [20]. The catalytic loop (residues 81–100 in M.HhaI) shifts substantially towards the DNA almost simultaneously with the target cytosine flipping. As a result, the flipped out base occurs in close proximity with the cofactor molecule inside of a closed catalytic cavity [23-25].

A nucleophilic attack of M.HhaI Cys81 thiol group on the cytosine C6 atom results in a forma‐ tion of M.HhaI–DNA conjugate. The Cys81 residue is a part of a conservative ProCys dipep‐ tide of the protein (motif IV, Figure 2). The Glu119 residue from GluAsnVal tripeptide (motif VI) protonates the cytosine N3 atom thus facilitating the nucleophilic attack by the thiol group. In the conjugate, the negative charge of the cytosine C5 atom causes its alkylation by the Ado‐ Met methyl group. Proton elimination from the C5 atom of the methylated cytosine and βelimination of the Cys residue result in a breakdown of the covalent DNA–protein complex and restore aromaticity of the modified cytosine base (Figure 2, a). The rate constant for meth‐ yl group transfer catalyzed by M.HhaI is about 0.14–0.26 s–1 [17, 18]. The following release of the reaction products is a rate-limiting step of the M.HhaI catalytic cycle [17, 18].

C5-DNA MTases HhaI and HpaII catalyze the methylation reaction in a distributive man‐ ner, *i.e.* dissociate from the DNA substrate after every catalytic act [1, 18]. Each one of these MTases is a component of a R–M system where the cognate RE searches for its recognition sites *via* scanning DNA by a linear diffusion mechanism. The MTase distributivity provides for the corresponding RE a possibility to bind an unmodified site before the MTase and to cleave it. On the other hand, C5-DNA MTase SssI [26] has no cognate RE and is able to methylate several recognition sites located in one DNA substrate in a processive manner, *i.e.* without dissociation from DNA after each catalytic act.

The affinity of prokaryotic C5-DNA MTases to their DNA ligands in the presence of non‐ reactive cofactor analogs increases in the following manner: dimethylated DNA << unme‐ thylated DNA < monomethylated DNA [1, 2, 27, 28]. A similar correlation is observed for the eukaryotic enzyme Dnmt1. The Dnmt1 affinity to monomethylated 5′-CG-3′/3′-GC-5′ sites is 2–200 times higher than to unmethylated ones depending on the experimental condi‐ tions [13, 29-35]. Dnmt1 modifies monomethylated sites processively (more than 50 sites per one binding act on average). Unmethylated sites (mainly 5′-CCGG-3′) are methylated by Dnmt1 in a distributive manner [36].

Mammalian C5-DNA MTases Dnmt3a and Dnmt3b methylate presumably unmodified sites in DNA and are responsible for *de novo* DNA methylation during embryonic development. Dinucleotides 5'-CG-3'/3′-GC-5′ are the main recognition sites of Dnmt3a and Dnmt3b. These enzymes are also able to modify dinucleotides 5′-CА-3′ but 10–100 times less efficiently [37]. The efficiency of methylation by Dnmt3a and Dnmt3b is also dependent on the sequences flanking the recognition site: 5′-RCGY-3′ is the most frequently methylated site whereas 5′- YCGR-3′ is methylated with a lower efficiency (R and Y are purine and pyrimidine nucleo‐ sides respectively). The difference in the methylation rate of these sites can exceed 500 times [38]. The primary structures of the Dnmt3a and Dnmt3b catalytic domains share identity of 84%. In contrast to Dnmt1, isolated catalytic domains of Dnmt3a and Dnmt3b retain their ac‐ tivity [12, 39]. Dnmt3a methylates DNA in a distributive manner while Dnmt3b modifies DNA processively since the DNA binding center of Dnmt3b is more positively charged than DNA binding center of Dnmt3a [37, 39]. Interestingly, a Cys residue substitution in the catalytic ProCys motif of an isolated Dnmt3a catalytic domain does not totally abolish the enzyme ac‐ tivity but merely decreases it 2–6 times. This Cys residue is shown to take part in the DNA– Dnmt3a conjugate formation. However, the Dnmt3a catalytic domain looses its activity com‐ pletely after a substitution of a Glu residue in the GluAsnVal motif. The active center of Dnmt3a seems to have an unusual conformation and the Cys residue perhaps does not have its optimal orientation. Therefore, the nucleophilic attack onto the cytosine C6 atom can be

(a) (b)

(a) (b)

**Figure 2.** *a*) Mechanism of cytosine C5 atom methylation by a C5-DNA MTase. (*b*) Mechanism of cytosine deamination

The target nucleotide flipping can occur spontaneously or with the help of an enzyme [20]. The catalytic loop (residues 81–100 in M.HhaI) shifts substantially towards the DNA almost simultaneously with the target cytosine flipping. As a result, the flipped out base occurs in

A nucleophilic attack of M.HhaI Cys81 thiol group on the cytosine C6 atom results in a forma‐ tion of M.HhaI–DNA conjugate. The Cys81 residue is a part of a conservative ProCys dipep‐ tide of the protein (motif IV, Figure 2). The Glu119 residue from GluAsnVal tripeptide (motif

close proximity with the cofactor molecule inside of a closed catalytic cavity [23-25].

as a possible side reaction of the methylation process.

34 Methylation - From DNA, RNA and Histones to Diseases and Treatment

Methylation - From DNA, RNA and Histones to Diseases and Treatment

performed by some other residue or hydroxyl ion. Post-translational modifications or interac‐ tion with other proteins are likely to be needed for the Dnmt3a activation [40].

was demonstrated for M.Alw26I from *Acinetobacter lwoffi* RFL26 (recognition site 5′- GT*C*TC-3′/3′-C*A*GAG-5′) and for M.Esp3I from *Hafnia alvei* RFL3 (recognition site 5′- CGT*C*TC-3′/3′-GC*A*GAG-5′) [44]. Each of these proteins contains the N6-DNA MTase domain in its N-terminal part and the C5-DNA MTase domain – in its C-terminal part [45]. The ability of each domain to methylate its recognition site in the absence of the second do‐

Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization

A gene coding for another enzyme consisting of two MTase domains has been constructed from two genes of *Helicobacter pylori* 26695 [46]. These genes are located in tandem orienta‐ tion and code for DNA MTases M.HpyAVIB and M.HpyAVIA. One nucleotide insertion be‐ fore the stop codon results in a formation of a fused gene. A similar mutation has been occurred naturally in *H. pylori* D27 strain [47]. M.HpyAVIB methylates the cytosine C5 atom and M.HpyAVIA methylates the adenine N6 atom in the sequence 5′-*C*CTC-3′/3′-GG*A*G-5′ (italicised). The obtained bifunctional protein contains the C5-DNA MTase domain and the N6-DNA MTase domain in its N-terminal and C-terminal parts, respectively. The both do‐ mains recognize in DNA the same sites as the initial proteins. The methylation kinetics and the properties of point mutants demonstrate that these domains function independently from each other. Each one of them contains its own catalytic and AdoMet binding motifs

Prokaryotic C5-DNA MTases M.HpaII, M.HhaI, M.SssI, Dcm (from *E*. *coli*), M.EcoRII, and

→ T mutagenesis *in vitro* [48-55]. Some of them demonstrate the mutagenic activity (M.HpaII, M.EcoRII, and Dcm) also *in vivo* increasing the mutagenesis rate up to 50 times [50, 54, 56]. Interestingly, prokaryotic M.MspI does not stimulate cytosine deamination *in vi‐ tro* but its mutagenic effect is comparable with the effect of other C5-DNA MTases when M.MspI is expressed in *E*. *coli* cells [57]. Moreover, М.EcoRII is shown to catalyze 5-methyl‐

The enzymatic catalysis of cytosine deamination is a side reaction of the methylation proc‐ ess. According to the standard mechanism (Figure 2), the cysteine thiol group performs a nucleophilic attack onto the cytosine C6 atom and at the same time the cytosine N3 atom gets protonated that altogether leads to the DNA–enzyme conjugate formation. As a result, the cytosine base aromaticity is disrupted and the C5 atom becomes negatively charged. However, the following step (the methyl group transfer to the C5 atom) becomes impossible in the absence of AdoMet. If water penetrates into the enzyme active center, hydroxylation of the cytosine C4 atom is likely to occur. These processes initiate the deamination reaction (Figure 2, b). Afterwards, the cytosine amino group is substituted with a carboxyl group and the base is converted into uracil [58, 59]. The presence of AdoMet or AdoHcy prevents water penetration into the active center and therefore inhibits deamination. A point mutant of М.HpaII incapable of AdoMet binding is a very effective catalyst of C → dU conversion [56]. The AdoMet analogs such as sinefungin and 5′-amino-5′-deoxyadenosine can increase the rate of enzymatic deamination even in the presence of AdoMet and AdoHcy [53]. They seem

C (5-methyl-2'-deoxycytidine)

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37

main has not been investigated.

**4.2. Deamination of cytosine and 5-methylcytosine**

cytosine conversion into thymine [52].

M.SsoII are shown to increase the rate of C → dU → T and m<sup>5</sup>

[46].

Murine Dnmt3a is able to perform automethylation, a methyl group transfer to the Cys resi‐ due of its own catalytic center in the presence of AdoMet. This reaction is irreversible and rather slow but its can be activated by Dnmt3L. In the presence of a duplex containing 5′- CG-3′/3'-GC-5' sites Dnmt3a methylates the substrate DNA but not its own Cys residue. The automethylation seems to have a regulatory function that enables to inactivate excessive en‐ zyme molecules in a cell. On the other hand, it can be just a side reaction which takes place in the absence of DNA [41].

A protein called Dnmt3L is also a member of Dnmt3 family. It is catalytically inactive but plays an important role as a stimulator of the Dnmt3a and Dnmt3b activity. A structure of a complex consisting of Dnmt3a and Dnmt3L C-terminal domain has been determined by Xray crystallography (PDB-code 2qrv) [42]. The complex is a heterotetramer where the subu‐ nits are localized in the following order: Dnmt3L–Dnmt3a–Dnmt3a–Dnmt3L. Two active centers of Dnmt3a are localized nearby and probably can methylate two 5′-CG-3′/3′-GC-5′ sites simultaneously. These recognition sites should be separated by 8–10 bp (about one turn of DNA double helix). Twelve murine genes which undergo maternal imprinting contain 5′- CG-3′/3′-GC-5′ sites localized in such a manner. Moreover, highly methylated regions of hu‐ man chromosome 21 possess 5′-CG-3′/3′-GC-5′ sites separated by 9, 18 and 27 bp more frequently in comparison to the unmethylated regions [43]. The effective methylation of these regions could be determined by the proper distribution of 5′-CG-3′/3′-GC-5′ sites. A substitution of Dnmt3a and Dnmt3L residues which do not take part in catalysis but are im‐ portant for the interface formation (Dnmt3a–Dnmt3L or Dnmt3a–Dnmt3a interfaces) results in a suppression of Dnmt3a activity. This fact confirms the importance of the appropriate complex formation between Dnmt3a and Dnmt3L [42]. The orientation of the Dnmt3a– Dnmt3L complex relative to the substrate DNA is still unclear.

### **4. Additional functions of prokaryotic (cytosine-5)-DNA methyltransferases**

#### **4.1. DNA methyltransferases with multiple methyltransferase domains**

Some C5-DNA MTases contain more than one MTase domain. The "additional" domain can belong to (cytosine-5)-DNA or (adenine-N6)-DNA MTases. Up to date, the Pfam database contains 5065 protein sequences that possess the characteristic domain of C5-DNA MTases (PF00145). Among them, 676 sequences (13%) contain two PF00145 domains and 42 sequen‐ ces (1%) – even three PF00145 domains. Such structures might have arisen as fusions of two adjacent genes encoding different MTases. However, none of these proteins has been stud‐ ied experimentally.

The ability of a single molecule to methylate both the cytosine C5 atom and the adenine N6 atom is typical for MTases that recognize asymmetric DNA sequences. This phenomenon was demonstrated for M.Alw26I from *Acinetobacter lwoffi* RFL26 (recognition site 5′- GT*C*TC-3′/3′-C*A*GAG-5′) and for M.Esp3I from *Hafnia alvei* RFL3 (recognition site 5′- CGT*C*TC-3′/3′-GC*A*GAG-5′) [44]. Each of these proteins contains the N6-DNA MTase domain in its N-terminal part and the C5-DNA MTase domain – in its C-terminal part [45]. The ability of each domain to methylate its recognition site in the absence of the second do‐ main has not been investigated.

A gene coding for another enzyme consisting of two MTase domains has been constructed from two genes of *Helicobacter pylori* 26695 [46]. These genes are located in tandem orienta‐ tion and code for DNA MTases M.HpyAVIB and M.HpyAVIA. One nucleotide insertion be‐ fore the stop codon results in a formation of a fused gene. A similar mutation has been occurred naturally in *H. pylori* D27 strain [47]. M.HpyAVIB methylates the cytosine C5 atom and M.HpyAVIA methylates the adenine N6 atom in the sequence 5′-*C*CTC-3′/3′-GG*A*G-5′ (italicised). The obtained bifunctional protein contains the C5-DNA MTase domain and the N6-DNA MTase domain in its N-terminal and C-terminal parts, respectively. The both do‐ mains recognize in DNA the same sites as the initial proteins. The methylation kinetics and the properties of point mutants demonstrate that these domains function independently from each other. Each one of them contains its own catalytic and AdoMet binding motifs [46].

#### **4.2. Deamination of cytosine and 5-methylcytosine**

performed by some other residue or hydroxyl ion. Post-translational modifications or interac‐

Murine Dnmt3a is able to perform automethylation, a methyl group transfer to the Cys resi‐ due of its own catalytic center in the presence of AdoMet. This reaction is irreversible and rather slow but its can be activated by Dnmt3L. In the presence of a duplex containing 5′- CG-3′/3'-GC-5' sites Dnmt3a methylates the substrate DNA but not its own Cys residue. The automethylation seems to have a regulatory function that enables to inactivate excessive en‐ zyme molecules in a cell. On the other hand, it can be just a side reaction which takes place

A protein called Dnmt3L is also a member of Dnmt3 family. It is catalytically inactive but plays an important role as a stimulator of the Dnmt3a and Dnmt3b activity. A structure of a complex consisting of Dnmt3a and Dnmt3L C-terminal domain has been determined by Xray crystallography (PDB-code 2qrv) [42]. The complex is a heterotetramer where the subu‐ nits are localized in the following order: Dnmt3L–Dnmt3a–Dnmt3a–Dnmt3L. Two active centers of Dnmt3a are localized nearby and probably can methylate two 5′-CG-3′/3′-GC-5′ sites simultaneously. These recognition sites should be separated by 8–10 bp (about one turn of DNA double helix). Twelve murine genes which undergo maternal imprinting contain 5′- CG-3′/3′-GC-5′ sites localized in such a manner. Moreover, highly methylated regions of hu‐ man chromosome 21 possess 5′-CG-3′/3′-GC-5′ sites separated by 9, 18 and 27 bp more frequently in comparison to the unmethylated regions [43]. The effective methylation of these regions could be determined by the proper distribution of 5′-CG-3′/3′-GC-5′ sites. A substitution of Dnmt3a and Dnmt3L residues which do not take part in catalysis but are im‐ portant for the interface formation (Dnmt3a–Dnmt3L or Dnmt3a–Dnmt3a interfaces) results in a suppression of Dnmt3a activity. This fact confirms the importance of the appropriate complex formation between Dnmt3a and Dnmt3L [42]. The orientation of the Dnmt3a–

tion with other proteins are likely to be needed for the Dnmt3a activation [40].

36 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Dnmt3L complex relative to the substrate DNA is still unclear.

**4. Additional functions of prokaryotic (cytosine-5)-DNA**

**4.1. DNA methyltransferases with multiple methyltransferase domains**

Some C5-DNA MTases contain more than one MTase domain. The "additional" domain can belong to (cytosine-5)-DNA or (adenine-N6)-DNA MTases. Up to date, the Pfam database contains 5065 protein sequences that possess the characteristic domain of C5-DNA MTases (PF00145). Among them, 676 sequences (13%) contain two PF00145 domains and 42 sequen‐ ces (1%) – even three PF00145 domains. Such structures might have arisen as fusions of two adjacent genes encoding different MTases. However, none of these proteins has been stud‐

The ability of a single molecule to methylate both the cytosine C5 atom and the adenine N6 atom is typical for MTases that recognize asymmetric DNA sequences. This phenomenon

in the absence of DNA [41].

**methyltransferases**

ied experimentally.

Prokaryotic C5-DNA MTases M.HpaII, M.HhaI, M.SssI, Dcm (from *E*. *coli*), M.EcoRII, and M.SsoII are shown to increase the rate of C → dU → T and m<sup>5</sup> C (5-methyl-2'-deoxycytidine) → T mutagenesis *in vitro* [48-55]. Some of them demonstrate the mutagenic activity (M.HpaII, M.EcoRII, and Dcm) also *in vivo* increasing the mutagenesis rate up to 50 times [50, 54, 56]. Interestingly, prokaryotic M.MspI does not stimulate cytosine deamination *in vi‐ tro* but its mutagenic effect is comparable with the effect of other C5-DNA MTases when M.MspI is expressed in *E*. *coli* cells [57]. Moreover, М.EcoRII is shown to catalyze 5-methyl‐ cytosine conversion into thymine [52].

The enzymatic catalysis of cytosine deamination is a side reaction of the methylation proc‐ ess. According to the standard mechanism (Figure 2), the cysteine thiol group performs a nucleophilic attack onto the cytosine C6 atom and at the same time the cytosine N3 atom gets protonated that altogether leads to the DNA–enzyme conjugate formation. As a result, the cytosine base aromaticity is disrupted and the C5 atom becomes negatively charged. However, the following step (the methyl group transfer to the C5 atom) becomes impossible in the absence of AdoMet. If water penetrates into the enzyme active center, hydroxylation of the cytosine C4 atom is likely to occur. These processes initiate the deamination reaction (Figure 2, b). Afterwards, the cytosine amino group is substituted with a carboxyl group and the base is converted into uracil [58, 59]. The presence of AdoMet or AdoHcy prevents water penetration into the active center and therefore inhibits deamination. A point mutant of М.HpaII incapable of AdoMet binding is a very effective catalyst of C → dU conversion [56].

The AdoMet analogs such as sinefungin and 5′-amino-5′-deoxyadenosine can increase the rate of enzymatic deamination even in the presence of AdoMet and AdoHcy [53]. They seem to trigger other reaction mechanisms which are not completely clarified yet. The supposed mechanisms include a water molecule direct attack onto the cytosine C4 atom. It becomes possible after the N3 atom protonation by a MTase which is a step of the methylation reac‐ tion. Two alternative mechanisms are suggested (Figure 3). According to the first one (mech‐ anism 1), the hydroxyl group of 5′-amino-5′-deoxyadenosine activates a water molecule producing a hydroxyl ion which attacks the cytosine C4 atom (Figure 3). According to the mechanism 2, 5′-amino-5′-deoxyadenosine acts as an acid and protonates the cytosine N4 atom (Figure 3) thus facilitating the amino group elimination in the form of ammonium ion [59]. The mechanisms 1 and 2 are based on a water molecule direct attack onto the C4 atom and differ considerably from the others as they do not require a MTase interaction with the cytosine C6 atom. Therefore, a mutant form of a MTase which does not catalyze the methyl‐ ation reaction should be able to catalyze these side reactions. Indeed, it has been shown ex‐ perimentally for an М.EcoRII point mutant where catalytic Cys was substituted with Ala [59].

karyotic MTases catalyze effectively the deamination reaction in the absence of the cofactor

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The different rates of the covalent enzyme–substrate complex formation in the absence of the cofactor could reflect different physiological functions of prokaryotic and eukaryotic C5- DNA MTases. Limited nutrition decreases AdoMet amounts in prokaryotic cells. The muta‐ tions derived from the cytosine deamination might not be lethal for a bacterial cell and could help to prevent hydrolysis of cellular DNA by phage endonucleases. So, the ability of a C5- DNA MTase to catalyze deamination can turn out a physiological advantage for a bacterial cell [5]. On the contrary, mammalian cells are not likely to benefit from this kind of mutagen‐ esis and therefore have developed mechanisms which provide low mutagenesis rate.

Two prokaryotic C5-DNA MTases are shown to have a topoisomerase activity, namely М.SssI from *Spiroplasma* MQI and M.MspI from *Moraxella species*. М.SssI is not a part of R–M system (there is no corresponding RE in *Spiroplasma* genome). As well as eukaryotic MTases, М.SssI modifies cytosine C5 atom in 5′-CG-3′/3′-GC-5′ sequences [61]. In the presence of 10 mM Mg2+, М.SssI provides relaxation of negatively supercoiled plasmids which leads to ac‐ cumulation of plasmids with different degree of supercoiling. The obtained set of plasmids is similar to the products of topoisomerase I from calf thymus. An ATP addition does not influence the topoisomerase activity of М.SssI. Since type II topoisomerases need ATP for the enzymatic activity, М.SssI can be regarded as a type I topoisomerase [62]. The MTase and the topoisomerase activities of М.SssI are functionally independent. The methylation process requires AdoMet, while the topoisomerase reaction demands Mg2+ ions. The M.SssI conservative motifs IV and VIII share a certain similarity with topoisomerase sequences. In particular, the motif IV contains Tyr that is an important catalytic residue in topoisomerases. A more detailed analysis of M.SssI regions responsible for the topoisomerase activity has

Different speculations are proposed to explain why these two activities are combined in one M.SssI molecule. Firstly, the topoisomerase activity alters the supercoiling degree of plas‐ mid DNA and thus could facilitate or complicate the cytosine flipping out of the DNA helix. Secondly, the methylation of 5′-CG-3'/3′-GC-5′ sites by M.SssI could change the DNA struc‐ ture. For example, a negatively supercoiled DNA region with large amount of methylated 5′- CG-3′/3′-GC-5′ sites is likely to be converted into Z-form. For the B-form restoration, topoisomerase activity is necessary. Thirdly, the change in DNA topology perhaps influen‐ ces the level of gene expression in *Spiroplasma*. Finally, the two different activities can be com‐ bined in one protein for the purpose of genome economy. *Spiroplasma* belongs to mycoplasmas – cellular organisms which have the smallest genome (from 600 to 1800 kbp). For a compari‐ son, the T4 bacteriophage genome consists of 165 kbp and the *E. coli* genome – of 4600 kbp [62].

M.MspI is a part of MspI R–M system and methylates the first cytosine residue in the se‐ quence 5′-*C*CGG-3′/3′-GGC*C*-5′ [63]. The unique property of this MTase is its ability to bend DNA at 142 ± 4° upon its binding to the methylation site. This was demonstrated using 127

[48, 53, 57].

**4.3. Topoisomerase activity**

not been conducted.

**Figure 3.** The supposed mechanisms of cytosine deamination catalyzed by an AdoMet analogue (sinefungin or 5′ amino-5′-deoxyadenosine). The AdoMet analogue might act as a base (mechanism 1) or as an acid (mechanism 2).

Both the methylation and the deamination reactions require flipping out of the cytosine resi‐ due. Thus, the longer cytosine base remains flipped out the faster these reactions go. The both reactions share a common intermediate – the enzyme–substrate conjugate (Figure 2). The usage of tritiated cytosine allows an estimation of tritium to hydrogen exchange rate at the cytosine C5 atom which can serve as a measure of the conjugate formation rate. In the absence of the cofactor, such an exchange catalyzed by murine Dnmt1 is slower than the one catalyzed by M.HhaI [60]. Thus, Dnmt1 forms the conjugate in the absence of AdoMet with a lower probability and therefore is less mutagenic than M.HhaI [60]. On the contrary, pro‐ karyotic MTases catalyze effectively the deamination reaction in the absence of the cofactor [48, 53, 57].

The different rates of the covalent enzyme–substrate complex formation in the absence of the cofactor could reflect different physiological functions of prokaryotic and eukaryotic C5- DNA MTases. Limited nutrition decreases AdoMet amounts in prokaryotic cells. The muta‐ tions derived from the cytosine deamination might not be lethal for a bacterial cell and could help to prevent hydrolysis of cellular DNA by phage endonucleases. So, the ability of a C5- DNA MTase to catalyze deamination can turn out a physiological advantage for a bacterial cell [5]. On the contrary, mammalian cells are not likely to benefit from this kind of mutagen‐ esis and therefore have developed mechanisms which provide low mutagenesis rate.

#### **4.3. Topoisomerase activity**

to trigger other reaction mechanisms which are not completely clarified yet. The supposed mechanisms include a water molecule direct attack onto the cytosine C4 atom. It becomes possible after the N3 atom protonation by a MTase which is a step of the methylation reac‐ tion. Two alternative mechanisms are suggested (Figure 3). According to the first one (mech‐ anism 1), the hydroxyl group of 5′-amino-5′-deoxyadenosine activates a water molecule producing a hydroxyl ion which attacks the cytosine C4 atom (Figure 3). According to the mechanism 2, 5′-amino-5′-deoxyadenosine acts as an acid and protonates the cytosine N4 atom (Figure 3) thus facilitating the amino group elimination in the form of ammonium ion [59]. The mechanisms 1 and 2 are based on a water molecule direct attack onto the C4 atom and differ considerably from the others as they do not require a MTase interaction with the cytosine C6 atom. Therefore, a mutant form of a MTase which does not catalyze the methyl‐ ation reaction should be able to catalyze these side reactions. Indeed, it has been shown ex‐ perimentally for an М.EcoRII point mutant where catalytic Cys was substituted with Ala

38 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 3.** The supposed mechanisms of cytosine deamination catalyzed by an AdoMet analogue (sinefungin or 5′ amino-5′-deoxyadenosine). The AdoMet analogue might act as a base (mechanism 1) or as an acid (mechanism 2).

Both the methylation and the deamination reactions require flipping out of the cytosine resi‐ due. Thus, the longer cytosine base remains flipped out the faster these reactions go. The both reactions share a common intermediate – the enzyme–substrate conjugate (Figure 2). The usage of tritiated cytosine allows an estimation of tritium to hydrogen exchange rate at the cytosine C5 atom which can serve as a measure of the conjugate formation rate. In the absence of the cofactor, such an exchange catalyzed by murine Dnmt1 is slower than the one catalyzed by M.HhaI [60]. Thus, Dnmt1 forms the conjugate in the absence of AdoMet with a lower probability and therefore is less mutagenic than M.HhaI [60]. On the contrary, pro‐

[59].

Two prokaryotic C5-DNA MTases are shown to have a topoisomerase activity, namely М.SssI from *Spiroplasma* MQI and M.MspI from *Moraxella species*. М.SssI is not a part of R–M system (there is no corresponding RE in *Spiroplasma* genome). As well as eukaryotic MTases, М.SssI modifies cytosine C5 atom in 5′-CG-3′/3′-GC-5′ sequences [61]. In the presence of 10 mM Mg2+, М.SssI provides relaxation of negatively supercoiled plasmids which leads to ac‐ cumulation of plasmids with different degree of supercoiling. The obtained set of plasmids is similar to the products of topoisomerase I from calf thymus. An ATP addition does not influence the topoisomerase activity of М.SssI. Since type II topoisomerases need ATP for the enzymatic activity, М.SssI can be regarded as a type I topoisomerase [62]. The MTase and the topoisomerase activities of М.SssI are functionally independent. The methylation process requires AdoMet, while the topoisomerase reaction demands Mg2+ ions. The M.SssI conservative motifs IV and VIII share a certain similarity with topoisomerase sequences. In particular, the motif IV contains Tyr that is an important catalytic residue in topoisomerases. A more detailed analysis of M.SssI regions responsible for the topoisomerase activity has not been conducted.

Different speculations are proposed to explain why these two activities are combined in one M.SssI molecule. Firstly, the topoisomerase activity alters the supercoiling degree of plas‐ mid DNA and thus could facilitate or complicate the cytosine flipping out of the DNA helix. Secondly, the methylation of 5′-CG-3'/3′-GC-5′ sites by M.SssI could change the DNA struc‐ ture. For example, a negatively supercoiled DNA region with large amount of methylated 5′- CG-3′/3′-GC-5′ sites is likely to be converted into Z-form. For the B-form restoration, topoisomerase activity is necessary. Thirdly, the change in DNA topology perhaps influen‐ ces the level of gene expression in *Spiroplasma*. Finally, the two different activities can be com‐ bined in one protein for the purpose of genome economy. *Spiroplasma* belongs to mycoplasmas – cellular organisms which have the smallest genome (from 600 to 1800 kbp). For a compari‐ son, the T4 bacteriophage genome consists of 165 kbp and the *E. coli* genome – of 4600 kbp [62].

M.MspI is a part of MspI R–M system and methylates the first cytosine residue in the se‐ quence 5′-*C*CGG-3′/3′-GGC*C*-5′ [63]. The unique property of this MTase is its ability to bend DNA at 142 ± 4° upon its binding to the methylation site. This was demonstrated using 127 bp DNA duplex and has not been shown for any other MTase [64]. Unlike M.SssI, M.MspI has an N-terminal part responsible for the topoisomerase activity. This part consists of 107 residues and is located before the conservative motif I. There are two regions of M.MspI that share similarity with topoisomerase sequences: the residues 32–98 and the conservative mo‐ tif VIII. In contrast to M.SssI, there is no similarity between the M.MspI conservative motif IV and the topoisomerase sequences. A mutant form of M.MspI lacking the first 34 residues retains the ability to methylate DNA but loses its topoisomerase activity. Mutant proteins M.MspI(W34A) and M.MspI(Y74A) also do not have topoisomerase activity but are still able to methylate the DNA substrate [65]. Additionally, the M.MspI C-terminal part contains a region (245–287 а.a.) that shares similarity with DNA ligase I active center. This is in accord‐ ance with the fact that the topoisomerase I activity includes ligation of the DNA strands [65].

quence 5′-GGNCC-3′/3′-CCNGG-5′ (N = A, G, C, T). It remains unclear which one cytosine is modified. The М.Eco47II N-terminal part is predicted to contain an HTH motif and is demonstrated to be responsible for the transcription regulation but not for the MTase activi‐ ty. Mutations introduced into the catalytic center of the enzyme result in suppression of the

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In the MspI R–M system, the *mspIM* and *mspIR* genes are transcribed divergently from the complementary DNA strands and their promoter regions (–35 elements) are separated by 6 bp. The regulatory site of M.MspI is located in the promoter region of the *mspIM* gene and contains a 12 bp inverted repeat. M.MspI protects from DNase I hydrolysis the region from –34 to +17 position in the "top" DNA strand and the region from –33 to +17 position in the "bottom" DNA strand (the numbers indicate the position relatively to the start point of the *mspIM* gene transcription). So, M.MspI interaction with the regulatory region prevents RNA polymerase binding to the promoter region and blocks transcription initiation from the *mspIM* gene. At the same time, M.MspI does not interact with the promoter region of the

R–M system ScrFI from *Lactococcus lactis subsp. cremoris* also contains a C5-DNA MTase that regulates gene expression in its R–M system [66]. The RE gene (*scrFIR*) is flanked by two genes that code for MTases: *scrFIBM* and *scrFIAM.* The *scrFIAM* gene has its own promoter while the *scrFIBM* and *scrFIR* genes are transcribed together with a gene of unknown func‐ tion – *orfX* (Figure 4). Both MTases from the ScrFI R–M system recognize and methylate the cytosine base in the sequence 5′-CCNGG-3′/3′-GGNCC-5′ (it remains unknown which one cytosine is methylated). The biological sense of existing of two MTase genes is not clear. The N-terminal part of M.ScrFIA is predicted to contain an HTH motif [72]. M.ScrFIA is also shown to bind to the regulatory region – a 15 bp inverted repeat located before the tran‐ scription start point for the *scrFIAM* gene. M.ScrFIA binds to this region and inhibits the ex‐

**Figure 4.** Genetic organization of the ScrFI R–M system. Thin arrows indicate the transcription initiation points.

R–M system LlaJI from *Lactococcus lactis* contains two C5-DNA MTases – M1.LlaJI and M2.LlaJI. These MTases have the same recognition site 5′-GA*C*GC-3′/3′-CTG*C*G-5′ but methylate different cytosine bases (italicised) in the "top" and the "bottom" DNA strands respectively [73, 74]. Genes coding for these MTases and two RE compose one operon and are transcribed from the same promoter (Figure 5, a). The promoter region contains a 24 bp inverted repeat that is a regulatory site for the MTases. This palindrome sequence contains two methylation sites of the LlaJI MTases one of which overlaps with –35 promoter element (Figure 5, b). At first, M2.LlaJI modifies both sites that enables binding of M1.LlaJI and methylation of cytosine bases in the opposite strand. Interaction of M1.LlaJI with the unme‐ thylated substrate has not been demonstrated [73]. M2.LlaJI acts only as a modifying en‐

methylation activity but do not disrupt the regulatory function [68].

*mspIR* gene and does not interfere with the expression of RE MspI [71].

pression of its own gene 72].

#### **4.4. (Cytosine-5)-DNA methyltransferases as transcription factors**

According to the Table 1, some C5-DNA MTases contain domains that can function as tran‐ scription factors. These domains are located in the N-terminal parts of the proteins and are followed by the MTase domains. The main structural element of these domains is a charac‐ teristic helix-turn-helix (HTH) motif that is also present in many transcription factors. To date, the Pfam database contains 68 sequences that include a domain with HTH motif fol‐ lowed by the C5-DNA MTase domain. Among them, 25 sequences belong to the HTH\_3 family (PF01381). The ability to downregulate expression of its own gene was shown experi‐ mentally only for 7 DNA MTases (M.MspI, M.EcoRII, M.ScrFIA, M1.LlaJI, M.Eco47II, M.SsoII, and M.Ecl18kI) [66-68]. Among them, M.SsoII and M.Ecl18kI are the most remarka‐ ble ones as they not only suppress the transcription of their own genes but also stimulate the transcription of their cognate RE genes.

M.EcoRII from *E. coli* R245 strain methylates C5 atom of the second cytosine in the sequence 5′-C*C*WGG-3′/3′-GGW*C*C-5′ (W = А or Т). *In vitro* experiments demonstrate that M.EcoRII can bind both its methylation site and the promoter region of its own gene. The enzyme's binding site has been determined by footprinting with DNase I: M.EcoRII protects 47 nu‐ cleotides in the "top" strand and 49 nucleotides in the "bottom" strand from DNase I hy‐ drolysis [69]. Thus, the binding site of M.EcoRII is located upstream of the MTase gene coding region and overlaps with its –10 and –35 promoter elements. This localization of M.EcoRII prevents RNA polymerase binding to the promoter and results in a suppression of the MTase gene transcription. The M.EcoRII binding site in the promoter region contains an imperfect inverted repeat (with 2 nucleotide substitutions). The repeat consists of two 11 bp sequences spaced by 12 bp. This kind of symmetry supposes the MTase to bind the promot‐ er region as a dimer though this protein is a monomer in solution. Investigation of M.EcoRII catalytically inactive forms shows that the efficiency of the MTase binding to the promoter region does not depend on its ability to methylate substrate. These facts demonstrate that M.EcoRII consists of two domains: a catalytic domain and a domain responsible for the in‐ teraction with the promoter region [70].

R–M system Eco47II from *E. coli* RFL47 strain contains a MTase that is also able to downre‐ gulate the expression of its own gene. М.Eco47II methylates the cytosine C5 atom in the se‐ quence 5′-GGNCC-3′/3′-CCNGG-5′ (N = A, G, C, T). It remains unclear which one cytosine is modified. The М.Eco47II N-terminal part is predicted to contain an HTH motif and is demonstrated to be responsible for the transcription regulation but not for the MTase activi‐ ty. Mutations introduced into the catalytic center of the enzyme result in suppression of the methylation activity but do not disrupt the regulatory function [68].

bp DNA duplex and has not been shown for any other MTase [64]. Unlike M.SssI, M.MspI has an N-terminal part responsible for the topoisomerase activity. This part consists of 107 residues and is located before the conservative motif I. There are two regions of M.MspI that share similarity with topoisomerase sequences: the residues 32–98 and the conservative mo‐ tif VIII. In contrast to M.SssI, there is no similarity between the M.MspI conservative motif IV and the topoisomerase sequences. A mutant form of M.MspI lacking the first 34 residues retains the ability to methylate DNA but loses its topoisomerase activity. Mutant proteins M.MspI(W34A) and M.MspI(Y74A) also do not have topoisomerase activity but are still able to methylate the DNA substrate [65]. Additionally, the M.MspI C-terminal part contains a region (245–287 а.a.) that shares similarity with DNA ligase I active center. This is in accord‐ ance with the fact that the topoisomerase I activity includes ligation of the DNA strands [65].

According to the Table 1, some C5-DNA MTases contain domains that can function as tran‐ scription factors. These domains are located in the N-terminal parts of the proteins and are followed by the MTase domains. The main structural element of these domains is a charac‐ teristic helix-turn-helix (HTH) motif that is also present in many transcription factors. To date, the Pfam database contains 68 sequences that include a domain with HTH motif fol‐ lowed by the C5-DNA MTase domain. Among them, 25 sequences belong to the HTH\_3 family (PF01381). The ability to downregulate expression of its own gene was shown experi‐ mentally only for 7 DNA MTases (M.MspI, M.EcoRII, M.ScrFIA, M1.LlaJI, M.Eco47II, M.SsoII, and M.Ecl18kI) [66-68]. Among them, M.SsoII and M.Ecl18kI are the most remarka‐ ble ones as they not only suppress the transcription of their own genes but also stimulate the

M.EcoRII from *E. coli* R245 strain methylates C5 atom of the second cytosine in the sequence 5′-C*C*WGG-3′/3′-GGW*C*C-5′ (W = А or Т). *In vitro* experiments demonstrate that M.EcoRII can bind both its methylation site and the promoter region of its own gene. The enzyme's binding site has been determined by footprinting with DNase I: M.EcoRII protects 47 nu‐ cleotides in the "top" strand and 49 nucleotides in the "bottom" strand from DNase I hy‐ drolysis [69]. Thus, the binding site of M.EcoRII is located upstream of the MTase gene coding region and overlaps with its –10 and –35 promoter elements. This localization of M.EcoRII prevents RNA polymerase binding to the promoter and results in a suppression of the MTase gene transcription. The M.EcoRII binding site in the promoter region contains an imperfect inverted repeat (with 2 nucleotide substitutions). The repeat consists of two 11 bp sequences spaced by 12 bp. This kind of symmetry supposes the MTase to bind the promot‐ er region as a dimer though this protein is a monomer in solution. Investigation of M.EcoRII catalytically inactive forms shows that the efficiency of the MTase binding to the promoter region does not depend on its ability to methylate substrate. These facts demonstrate that M.EcoRII consists of two domains: a catalytic domain and a domain responsible for the in‐

R–M system Eco47II from *E. coli* RFL47 strain contains a MTase that is also able to downre‐ gulate the expression of its own gene. М.Eco47II methylates the cytosine C5 atom in the se‐

**4.4. (Cytosine-5)-DNA methyltransferases as transcription factors**

40 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

transcription of their cognate RE genes.

teraction with the promoter region [70].

In the MspI R–M system, the *mspIM* and *mspIR* genes are transcribed divergently from the complementary DNA strands and their promoter regions (–35 elements) are separated by 6 bp. The regulatory site of M.MspI is located in the promoter region of the *mspIM* gene and contains a 12 bp inverted repeat. M.MspI protects from DNase I hydrolysis the region from –34 to +17 position in the "top" DNA strand and the region from –33 to +17 position in the "bottom" DNA strand (the numbers indicate the position relatively to the start point of the *mspIM* gene transcription). So, M.MspI interaction with the regulatory region prevents RNA polymerase binding to the promoter region and blocks transcription initiation from the *mspIM* gene. At the same time, M.MspI does not interact with the promoter region of the *mspIR* gene and does not interfere with the expression of RE MspI [71].

R–M system ScrFI from *Lactococcus lactis subsp. cremoris* also contains a C5-DNA MTase that regulates gene expression in its R–M system [66]. The RE gene (*scrFIR*) is flanked by two genes that code for MTases: *scrFIBM* and *scrFIAM.* The *scrFIAM* gene has its own promoter while the *scrFIBM* and *scrFIR* genes are transcribed together with a gene of unknown func‐ tion – *orfX* (Figure 4). Both MTases from the ScrFI R–M system recognize and methylate the cytosine base in the sequence 5′-CCNGG-3′/3′-GGNCC-5′ (it remains unknown which one cytosine is methylated). The biological sense of existing of two MTase genes is not clear. The N-terminal part of M.ScrFIA is predicted to contain an HTH motif [72]. M.ScrFIA is also shown to bind to the regulatory region – a 15 bp inverted repeat located before the tran‐ scription start point for the *scrFIAM* gene. M.ScrFIA binds to this region and inhibits the ex‐ pression of its own gene 72].

**Figure 4.** Genetic organization of the ScrFI R–M system. Thin arrows indicate the transcription initiation points.

R–M system LlaJI from *Lactococcus lactis* contains two C5-DNA MTases – M1.LlaJI and M2.LlaJI. These MTases have the same recognition site 5′-GA*C*GC-3′/3′-CTG*C*G-5′ but methylate different cytosine bases (italicised) in the "top" and the "bottom" DNA strands respectively [73, 74]. Genes coding for these MTases and two RE compose one operon and are transcribed from the same promoter (Figure 5, a). The promoter region contains a 24 bp inverted repeat that is a regulatory site for the MTases. This palindrome sequence contains two methylation sites of the LlaJI MTases one of which overlaps with –35 promoter element (Figure 5, b). At first, M2.LlaJI modifies both sites that enables binding of M1.LlaJI and methylation of cytosine bases in the opposite strand. Interaction of M1.LlaJI with the unme‐ thylated substrate has not been demonstrated [73]. M2.LlaJI acts only as a modifying en‐ zyme while Ml.LlaJI has an additional capability of binding to the inverted repeat that contains –35 promoter element. The N-terminal part of Ml.LlaJI is predicted to contain an HTH motif. The M1.LlaJI binding to the inverted repeat results in suppression of gene tran‐ scription for the whole LlaJI operon. This mechanism seems to be unique since methylation usually decreases the binding efficiency of a repressor with its operator [74]. The complicat‐ ed regulatory mechanism is likely to enable fine tuning of transcriptional level in LlaJI R–M system [74].

sion level. The same effect is observed after the transformation with a plasmid encoding M.NlaX – a protein homologous to the M.SsoII domain responsible for methylation. On the contrary, a plasmid encoding a fusion of the M.SsoII first 72 residues with the full-length M.NlaX gives the same effect as the plasmid encoding the full-length M.SsoII [75]. These ex‐ periments demonstrate that the M.SsoII ability to regulate transcription in the SsoII R–M system is determined by its N-terminal part (72 residues). This part is predicted to contain

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M.SsoII is a typical C5-DNA MTase which modifies the second cytosine in the sequence 5′- C*C*NGG-3′/3′-GGN*C*C-5′. Moreover, M.SsoII interacts with the intergenic region of the SsoII R–M system protecting from DNase I hydrolysis 48 nucleotides in the top strand and 52 nu‐ cleotides in the bottom strand [75]. The intergenic region contains a 15 bp inverted repeat (regulatory site) [76]. Seven guanine bases interacting with M.SsoII are identified using pro‐ tection footprinting; six of them are located inside the regulatory site symmetrically relative to the central А•Т pair (Figure 7) [76]. Interference footprinting experiments with formic acid, hydrazine, dimethyl sulfate, and *N*-ethyl-*N*-nitrosourea show 6 guanine, 2 adenine, and 4 thymine residues as well as 6 phosphate groups interacting with M.SsoII. These nu‐ cleotides form two symmetrically located clusters: 5'-GGA-3' and 5'-TGT-3' in each DNA strand of the regulatory site (Figure 7) [77]. Such a symmetrical interaction with the both halves of the palindromic site is typical for many regulatory proteins which bind to the op‐ erator sequence in a dimeric form and contain an α-helix that interacts with the DNA major groove recognizing the specific sequence. M.SsoII as a transcription factor is supposed to

**Figure 6.** Genetic organization of the SsoII R–M system. The RE and MTase genes are depicted with green and yellow arrows respectively. The initiation codons are underlined. The region protected by M.SsoII from DNase I hydrolysis is highlighted with grey. The regulatory site is in red. The transcription initiation points of the RE and MTase genes are

an HTH motif.

have the same mechanism of functioning.

marked by black arrows, their promoter elements are in blue.

**Figure 5.** *a*) Genetic organization of the LlaJI R–M system. The thin arrow marks the transcription initiation point of the operon. (*b*) A fragment of the promoter region of the LlaJI R–M system. Two black arrows indicate the 24 bp in‐ verted repeat. The methylation sites are highlighted with yellow. The transcription start point is marked by "+1", the promoter –10 and –35 elements are in blue. The start codon is in red.

Transcription regulation has been studied most thoroughly in the R–M systems SsoII from *Shigella sonnei* 47 and Ecl18kI from *Enterobacter сloacea* 18k. The nucleotide sequences of these systems share 99% identity. The sequences of the intergenic regions are completely identical while the proteins differ in 1 amino acid residue. We will refer to such R–M systems as to SsoII-like ones. The genes of the SsoII R–M system are located divergently and spaced by an intergenic region of 109 bp (Figure 6). To investigate the regulation mechanism of the sys‐ tem, two plasmids have been constructed, with both possible combinations of the intergenic region and a *lacZ* gene which encodes β-galactosidase. Thus, the expression of the *lacZ* gene is under control of a promoter of the *ssoIIM* gene (pACYC-SsoIIM) or the *ssoIIR* gene (pA‐ CYC-SsoIIR). The β-galactosidase expression level is found to be 540 times higher in the cells containing the pACYC-SsoIIM plasmid in comparison to the other plasmid. Therefore, the expression from the *ssoIIM* promoter is much higher than from the *ssoIIR* promoter. When transformed with an additional plasmid where the *ssoIIM* gene is under its own promoter, the cells containing pACYC-SsoIIM demonstrate a 20-fold decrease of the β-galactosidase expression while the cells containing pACYC-SsoIIR display an 8-fold increase of it [75]. Thus, M.SsoII is shown to downregulate the expression of its own gene and to stimulate the expression of the cognate RE gene. The transformation of the cells with a plasmid encoding a mutant M.SsoII without its first 72 residues does not influence the β-galactosidase expres‐ sion level. The same effect is observed after the transformation with a plasmid encoding M.NlaX – a protein homologous to the M.SsoII domain responsible for methylation. On the contrary, a plasmid encoding a fusion of the M.SsoII first 72 residues with the full-length M.NlaX gives the same effect as the plasmid encoding the full-length M.SsoII [75]. These ex‐ periments demonstrate that the M.SsoII ability to regulate transcription in the SsoII R–M system is determined by its N-terminal part (72 residues). This part is predicted to contain an HTH motif.

zyme while Ml.LlaJI has an additional capability of binding to the inverted repeat that contains –35 promoter element. The N-terminal part of Ml.LlaJI is predicted to contain an HTH motif. The M1.LlaJI binding to the inverted repeat results in suppression of gene tran‐ scription for the whole LlaJI operon. This mechanism seems to be unique since methylation usually decreases the binding efficiency of a repressor with its operator [74]. The complicat‐ ed regulatory mechanism is likely to enable fine tuning of transcriptional level in LlaJI R–M

(a)

(b)

promoter –10 and –35 elements are in blue. The start codon is in red.

42 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 5.** *a*) Genetic organization of the LlaJI R–M system. The thin arrow marks the transcription initiation point of the operon. (*b*) A fragment of the promoter region of the LlaJI R–M system. Two black arrows indicate the 24 bp in‐ verted repeat. The methylation sites are highlighted with yellow. The transcription start point is marked by "+1", the

Transcription regulation has been studied most thoroughly in the R–M systems SsoII from *Shigella sonnei* 47 and Ecl18kI from *Enterobacter сloacea* 18k. The nucleotide sequences of these systems share 99% identity. The sequences of the intergenic regions are completely identical while the proteins differ in 1 amino acid residue. We will refer to such R–M systems as to SsoII-like ones. The genes of the SsoII R–M system are located divergently and spaced by an intergenic region of 109 bp (Figure 6). To investigate the regulation mechanism of the sys‐ tem, two plasmids have been constructed, with both possible combinations of the intergenic region and a *lacZ* gene which encodes β-galactosidase. Thus, the expression of the *lacZ* gene is under control of a promoter of the *ssoIIM* gene (pACYC-SsoIIM) or the *ssoIIR* gene (pA‐ CYC-SsoIIR). The β-galactosidase expression level is found to be 540 times higher in the cells containing the pACYC-SsoIIM plasmid in comparison to the other plasmid. Therefore, the expression from the *ssoIIM* promoter is much higher than from the *ssoIIR* promoter. When transformed with an additional plasmid where the *ssoIIM* gene is under its own promoter, the cells containing pACYC-SsoIIM demonstrate a 20-fold decrease of the β-galactosidase expression while the cells containing pACYC-SsoIIR display an 8-fold increase of it [75]. Thus, M.SsoII is shown to downregulate the expression of its own gene and to stimulate the expression of the cognate RE gene. The transformation of the cells with a plasmid encoding a mutant M.SsoII without its first 72 residues does not influence the β-galactosidase expres‐

system [74].

M.SsoII is a typical C5-DNA MTase which modifies the second cytosine in the sequence 5′- C*C*NGG-3′/3′-GGN*C*C-5′. Moreover, M.SsoII interacts with the intergenic region of the SsoII R–M system protecting from DNase I hydrolysis 48 nucleotides in the top strand and 52 nu‐ cleotides in the bottom strand [75]. The intergenic region contains a 15 bp inverted repeat (regulatory site) [76]. Seven guanine bases interacting with M.SsoII are identified using pro‐ tection footprinting; six of them are located inside the regulatory site symmetrically relative to the central А•Т pair (Figure 7) [76]. Interference footprinting experiments with formic acid, hydrazine, dimethyl sulfate, and *N*-ethyl-*N*-nitrosourea show 6 guanine, 2 adenine, and 4 thymine residues as well as 6 phosphate groups interacting with M.SsoII. These nu‐ cleotides form two symmetrically located clusters: 5'-GGA-3' and 5'-TGT-3' in each DNA strand of the regulatory site (Figure 7) [77]. Such a symmetrical interaction with the both halves of the palindromic site is typical for many regulatory proteins which bind to the op‐ erator sequence in a dimeric form and contain an α-helix that interacts with the DNA major groove recognizing the specific sequence. M.SsoII as a transcription factor is supposed to have the same mechanism of functioning.

**Figure 6.** Genetic organization of the SsoII R–M system. The RE and MTase genes are depicted with green and yellow arrows respectively. The initiation codons are underlined. The region protected by M.SsoII from DNase I hydrolysis is highlighted with grey. The regulatory site is in red. The transcription initiation points of the RE and MTase genes are marked by black arrows, their promoter elements are in blue.

Arg35 or Arg38 substitution with Ala in M.Ecl18kI significantly impairs the protein binding to the regulatory site [79]. According to the computer simulation performed for M.SsoII, these residues belong to the second recognizing helix of the HTH motif [78]. Arg38 is sup‐ posed to form contacts with guanine bases of 5'-GGA-3' trinucleotide in one DNA strand while Arg35 can interact with thymine bases and DNA backbone of 5'-TGT-3' trinucleotide in the other strand (Figure 7). Amino acid substitutions in the M.Ecl18kI N-terminal part in‐ fluence the ability of this protein to regulate transcription in the R–M system. However, there is no correlation between M.Ecl18kI affinity to the regulatory site and the amounts of the RNA transcripts [80]. Amino acid substitutions in the M.Ecl18kI N-terminal part in‐ crease the methylation activity of this enzyme in most of cases. There is also an inverse rela‐ tionship: an M.SsoII point mutant which has Cys142 substituted with Ala is catalytically inactive but demonstrates an increased affinity to the regulatory site and effectively regu‐ lates transcription in the SsoII R–M system [79]. Thus, the interconnection between the two DNA binding sites is experimentally demonstrated for the SsoII-like DNA MTases. More‐ over, it has been shown recently that M.SsoII binding to the regulatory site prevents its in‐ teraction with the methylation site. Thus, the two functions of the protein are mutually exclusive [67, 81].

fere with RNA polymerase binding to the RE gene promoter. Thus, the RE gene is activated

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All known eukaryotic DNA MTases methylate C5 atom of cytosine. Eukaryotic MTases are

Murine and human Dnmt1 consist of 1620 and 1616 amino acid residues respectively. The pri‐

mary structures of these proteins share 85% identity. Dnmt1 molecule contains the follow‐

ing domains and functionally important regions (listed starting from the N-end, Figure 8):

**1.** charge-rich domain or DMAP1-binding domain (DMAP1 is DNA methyltransferase as‐

**4.** RFTS domain (replication foci targeting sequence), also called TS (targeting sequence),

**6.** BAH1 and BAH2 (bromo-adjacent homology or bromo-associated homology) domains

**7.** KG linker (consists of Lys and Gly residues) which connects N- and C-terminal parts of

The N-terminal part of Dnmt1 contains several domains that regulate the activity of the cata‐ lytic domain. Such a structural organization supposes that the Dnmt1 gene has arisen as fu‐

sion of a MTase gene with nonhomologous genes of other DNA-binding proteins [11, 83].

which are parts of the so-called PBHD domain (polybromo homology domain);

**3.** at least three functionally independent nuclear localization signals, NLS [82];

RFD (replication foci domain) or TRF (targeting to replication foci) domain;

**5.** cysteine-rich Zn2+-binding domain, also called CXXC domain;

indirectly *via* averting of RNA polymerase binding with the MTase gene promoter [80].

**5. Domains of eukaryotic DNA methyltransferases**

usually multidomain proteins (Figure 8).

**5.1. Functional domains of mammalian Dnmt1**

sociated protein, a transcription repressor);

**2.** PCNA-binding domain, PBD;

Dnmt1;

**8.** C-terminal catalytic domain.

**Figure 7.** Scheme of the contacts formed between the two N-terminal domains of M.SsoII and the regulatory DNA site. The heterocyclic bases and the phosphate groups interacting with M.SsoII identified by footprinting are marked by red and blue respectively.

The transcription start point of the RE gene is located in the beginning of the MTase gene in the SsoII and Ecl18kI R–M systems (Figure 6) [67]. The transcription start point of the MTase gene is located inside the region protected by M.SsoII from DNase I hydrolysis, 5 bp away from the regulatory site. The suppression of the MTase gene transcription is based on the competitive binding of the MTase and RNA polymerase with the intergenic region of the SsoII (or Ecl18kI) R–M system. The MTase interaction with the regulatory site does not inter‐ fere with RNA polymerase binding to the RE gene promoter. Thus, the RE gene is activated indirectly *via* averting of RNA polymerase binding with the MTase gene promoter [80].

#### **5. Domains of eukaryotic DNA methyltransferases**

All known eukaryotic DNA MTases methylate C5 atom of cytosine. Eukaryotic MTases are usually multidomain proteins (Figure 8).

#### **5.1. Functional domains of mammalian Dnmt1**

Murine and human Dnmt1 consist of 1620 and 1616 amino acid residues respectively. The pri‐ mary structures of these proteins share 85% identity. Dnmt1 molecule contains the follow‐ ing domains and functionally important regions (listed starting from the N-end, Figure 8):


Arg35 or Arg38 substitution with Ala in M.Ecl18kI significantly impairs the protein binding to the regulatory site [79]. According to the computer simulation performed for M.SsoII, these residues belong to the second recognizing helix of the HTH motif [78]. Arg38 is sup‐ posed to form contacts with guanine bases of 5'-GGA-3' trinucleotide in one DNA strand while Arg35 can interact with thymine bases and DNA backbone of 5'-TGT-3' trinucleotide in the other strand (Figure 7). Amino acid substitutions in the M.Ecl18kI N-terminal part in‐ fluence the ability of this protein to regulate transcription in the R–M system. However, there is no correlation between M.Ecl18kI affinity to the regulatory site and the amounts of the RNA transcripts [80]. Amino acid substitutions in the M.Ecl18kI N-terminal part in‐ crease the methylation activity of this enzyme in most of cases. There is also an inverse rela‐ tionship: an M.SsoII point mutant which has Cys142 substituted with Ala is catalytically inactive but demonstrates an increased affinity to the regulatory site and effectively regu‐ lates transcription in the SsoII R–M system [79]. Thus, the interconnection between the two DNA binding sites is experimentally demonstrated for the SsoII-like DNA MTases. More‐ over, it has been shown recently that M.SsoII binding to the regulatory site prevents its in‐ teraction with the methylation site. Thus, the two functions of the protein are mutually

44 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 7.** Scheme of the contacts formed between the two N-terminal domains of M.SsoII and the regulatory DNA site. The heterocyclic bases and the phosphate groups interacting with M.SsoII identified by footprinting are marked

The transcription start point of the RE gene is located in the beginning of the MTase gene in the SsoII and Ecl18kI R–M systems (Figure 6) [67]. The transcription start point of the MTase gene is located inside the region protected by M.SsoII from DNase I hydrolysis, 5 bp away from the regulatory site. The suppression of the MTase gene transcription is based on the competitive binding of the MTase and RNA polymerase with the intergenic region of the SsoII (or Ecl18kI) R–M system. The MTase interaction with the regulatory site does not inter‐

exclusive [67, 81].

by red and blue respectively.


The N-terminal part of Dnmt1 contains several domains that regulate the activity of the cata‐ lytic domain. Such a structural organization supposes that the Dnmt1 gene has arisen as fu‐ sion of a MTase gene with nonhomologous genes of other DNA-binding proteins [11, 83].

**Figure 8.** Domain architecture of mammalian С5-DNA MTases. The C-terminal domains are marked by a light blue fill‐ ing. The other domains are signed. Roman numerals indicate the conservative motifs characteristic for C5-DNA MTases in the C-terminal domains. The borders of the Dnmt1 domains are shown according to [9].

(a) (b)

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**Figure 9.** Spatial structure of murine Dnmt1. (*a*) Dnmt1(650–1602) complexed with AdoHcy and a 19 bp DNA duplex containing two unmethylated 5′-CG-3′/3′-GC-5′ sites (PDB code: 3pt6). (*b*) Dnmt1(291–1620) in apo-form (PDB code: 3av4). The RFTS, CXXC, BAH1, BAH2, and MTase domains are colored in magenta, cyan, green, orange, and blue, re‐ spectively. Zn2+ ions are in red and AdoHcy is in yellow, both in space-filling representation. DNA is in grey, in sticks representation. The autoinhibitory linker CXXC–BAH1 is colored in red. The borders of the domains are shown accord‐

The first 120 residues of Dnmt1 can bind the transcription repressor DMAP1. This interac‐ tion provides DMAP1 presence in replication foci during the whole S phase of cell cycle. Moreover, Dnmt1 interacts directly with histone deacetylase 2 (HDAC2) during late S phase. Dnmt1, HDAC2, and DMAP1 form a complex *in vivo*. Since the direct interaction between HDAC2 and DMAP1 has not been demonstrated, Dnmt1 is likely to serve as a basis for this

A special form of Dnmt1, Dnmt1o, is synthesized in oocytes. It lacks the first 118 residues that compose the DMAP1-binding domain. Dnmt1o is accumulated in oocyte cytoplasm and is transferred into nuclei of the 8-cell stage embryo where it is likely to be responsible for maintaining the methylation patterns of imprinted genes [82]. Dnmt1o is replaced by the regular Dnmt1 after implantation of the blastocyst. Homozygous mutant mice containing Dnmt1o in all somatic cells show a normal phenotype and have a normal level of genome methylation. This fact confirms the ability of Dnmt1o to perform all Dnmt1 functions. How‐ ever, the Dnmt1o amounts and the corresponding enzymatic activity are much higher than the ones of Dnmt1. In heterozygous embryonic stem cells, the expression levels of Dnmt1o and Dnmt1 are the same. Though, in adult mice the Dnmt1o amount is 5 times higher than the Dnmt1 amount. So Dnmt1o seems to be more stable than Dnmt1 [89]. The DMAP1-bind‐

ing to [9].

*5.1.1. DMAP1-binding domain*

complex formation [88].

Structures of two Dnmt1 complexes with DNA have been determined recently by X-ray crystallography: a fragment of murine Dnmt1 (residues 650–1602) with AdoHcy and a 19 bp DNA duplex containing two unmethylated 5′-CG-3′/3′-GC-5′ sites (PDB code: 3pt6) and a fragment of human Dnmt1 (residues 646–1600) with AdoHcy and the same duplex (PDB code: 3pta) [9]. The catalytic domain of Dnmt1 forms a core of the complex and makes con‐ tacts with the DNA on one side and with both BAH domains on the other side (Figure 9). AdoHcy molecule is located in the active center of the catalytic domain. The CXXC and BAH1 domains are located on different sides of the catalytic domain and are connected by a long CXXC–BAH1 linker. The BAH1 and BAH2 domains are located distantly from the bound DNA and are separated from each other by an α-helical linker. The KG linker is dis‐ ordered in the crystal. Different Dnmt1 domains are discussed further in this chapter.

It is worth to note that Dnmt1 targeting to replication foci is provided by three types of do‐ mains located in its N-terminal part: the PCNA binding domain, the RFTS domain, and the BAH domains [84-86]. Studying of Dnmt1 deletion mutants revealed 3 different DNA bind‐ ing regions: the residues 1–343, the CXXC domain (residues 613–748), and the catalytic do‐ main (residues 1124–1620) [13]. The catalytic domain binds preferentially monomethylated 5′-CG-3′/3′-GC-5′ sites while the fragment of residues 1–343 binds these sites independently of their methylation status [13]. The CXXC domain is shown to bind primarily dimethylated sites [13]. However, in the crystal structure the CXXC domain interacts with an unmethylat‐ ed substrate [9].

Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization http://dx.doi.org/10.5772/52046 47

**Figure 9.** Spatial structure of murine Dnmt1. (*a*) Dnmt1(650–1602) complexed with AdoHcy and a 19 bp DNA duplex containing two unmethylated 5′-CG-3′/3′-GC-5′ sites (PDB code: 3pt6). (*b*) Dnmt1(291–1620) in apo-form (PDB code: 3av4). The RFTS, CXXC, BAH1, BAH2, and MTase domains are colored in magenta, cyan, green, orange, and blue, re‐ spectively. Zn2+ ions are in red and AdoHcy is in yellow, both in space-filling representation. DNA is in grey, in sticks representation. The autoinhibitory linker CXXC–BAH1 is colored in red. The borders of the domains are shown accord‐ ing to [9].

#### *5.1.1. DMAP1-binding domain*

**Figure 8.** Domain architecture of mammalian С5-DNA MTases. The C-terminal domains are marked by a light blue fill‐ ing. The other domains are signed. Roman numerals indicate the conservative motifs characteristic for C5-DNA MTases

Structures of two Dnmt1 complexes with DNA have been determined recently by X-ray crystallography: a fragment of murine Dnmt1 (residues 650–1602) with AdoHcy and a 19 bp DNA duplex containing two unmethylated 5′-CG-3′/3′-GC-5′ sites (PDB code: 3pt6) and a fragment of human Dnmt1 (residues 646–1600) with AdoHcy and the same duplex (PDB code: 3pta) [9]. The catalytic domain of Dnmt1 forms a core of the complex and makes con‐ tacts with the DNA on one side and with both BAH domains on the other side (Figure 9). AdoHcy molecule is located in the active center of the catalytic domain. The CXXC and BAH1 domains are located on different sides of the catalytic domain and are connected by a long CXXC–BAH1 linker. The BAH1 and BAH2 domains are located distantly from the bound DNA and are separated from each other by an α-helical linker. The KG linker is dis‐

ordered in the crystal. Different Dnmt1 domains are discussed further in this chapter.

ed substrate [9].

It is worth to note that Dnmt1 targeting to replication foci is provided by three types of do‐ mains located in its N-terminal part: the PCNA binding domain, the RFTS domain, and the BAH domains [84-86]. Studying of Dnmt1 deletion mutants revealed 3 different DNA bind‐ ing regions: the residues 1–343, the CXXC domain (residues 613–748), and the catalytic do‐ main (residues 1124–1620) [13]. The catalytic domain binds preferentially monomethylated 5′-CG-3′/3′-GC-5′ sites while the fragment of residues 1–343 binds these sites independently of their methylation status [13]. The CXXC domain is shown to bind primarily dimethylated sites [13]. However, in the crystal structure the CXXC domain interacts with an unmethylat‐

in the C-terminal domains. The borders of the Dnmt1 domains are shown according to [9].

46 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

The first 120 residues of Dnmt1 can bind the transcription repressor DMAP1. This interac‐ tion provides DMAP1 presence in replication foci during the whole S phase of cell cycle. Moreover, Dnmt1 interacts directly with histone deacetylase 2 (HDAC2) during late S phase. Dnmt1, HDAC2, and DMAP1 form a complex *in vivo*. Since the direct interaction between HDAC2 and DMAP1 has not been demonstrated, Dnmt1 is likely to serve as a basis for this complex formation [88].

A special form of Dnmt1, Dnmt1o, is synthesized in oocytes. It lacks the first 118 residues that compose the DMAP1-binding domain. Dnmt1o is accumulated in oocyte cytoplasm and is transferred into nuclei of the 8-cell stage embryo where it is likely to be responsible for maintaining the methylation patterns of imprinted genes [82]. Dnmt1o is replaced by the regular Dnmt1 after implantation of the blastocyst. Homozygous mutant mice containing Dnmt1o in all somatic cells show a normal phenotype and have a normal level of genome methylation. This fact confirms the ability of Dnmt1o to perform all Dnmt1 functions. How‐ ever, the Dnmt1o amounts and the corresponding enzymatic activity are much higher than the ones of Dnmt1. In heterozygous embryonic stem cells, the expression levels of Dnmt1o and Dnmt1 are the same. Though, in adult mice the Dnmt1o amount is 5 times higher than the Dnmt1 amount. So Dnmt1o seems to be more stable than Dnmt1 [89]. The DMAP1-bind‐

ing domain is likely to decrease Dnmt1 stability *in vivo* and thus could be involved in Dnmt1 degradation.

#### *5.1.2. PCNA-binding domain*

The Dnmt1 residues 163–174 are responsible for binding of PCNA (proliferating cell nuclear antigen, also known as processivity factor for DNA polymerase δ). Dnmt1 relocates to DNA replication foci when the cell enters S phase [84, 90]. Its binding with PCNA is observed in the regions of newly synthesized DNA in intact cells. This binding does not influence the MTase activity of Dnmt1 [85].

In mammalian cells, newly replicated DNA is rapidly packaged into nucleosomes to which histone H1 is added further [91, 92]. Histone H1 has a high affinity to methylated DNA re‐ gardless its nucleotide sequence [93] and can suppress the Dnmt1 enzymatic activity [94]. Therefore, the maintenance methylation should be performed before DNA is packaged in nucleosomes. Dnmt1 binding with PCNA probably underlies a special mechanism required for coordination of these processes in a cell [85].

**Figure 10.** Spatial structure of the RFTS domain from human Dnmt1 (PDB code: 3epz). β-Barrel is colored in yellow,

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The cysteine-rich Zn2+-binding domain contains several cysteine residues organized in CXXC motifs which provide a name to the domain. The CXXC domain of Dnmt1 is similar to cysteine-rich domains of other chromatin-associated proteins such as the MeCP2 protein, the CG binding protein (CGBP), the histone MTase MLL, and the histone demethylases JHDM1A and JHDM1B. This domain is shown to bind unmethylated 5′-CG-3′/3′-GC-5′ sites *in vitro* in the case of MBD1, MLL, CGBP, JHDM1B, and Dnmt1 proteins [3]. The CXXC do‐ main of Dnmt1 is crescent-shaped and contains 8 conservative catalytically important Cys residues [98]. These residues are clustered in two groups and bind two Zn2+ ions. In the structure of murine Dnmt1(650–1602) complexed with AdoHcy and 19 bp DNA (PDB code: 3pt6), all the specific contacts with DNA are formed by the CXXC domain which interacts with both major and minor grooves [9]. A loop region of the CXXC domain (Arg684-Ser685- Lys686-Gln687) penetrates into the major groove and forms contacts with heterocyclic bases and phosphate groups. The guanine bases of the 5′-CG-3′ dinucleotide are recognized by the side chains of Lys686 and Gln687, whereas the cytosine bases are recognized by the backbone interactions of Ser685 and Lys686. Salt bridges are formed between the Arg side

The CXXC domain is known to bind specifically unmethylated 5′-CG-3′/3′-GC-5′ sites [98, 99]. The structural data confirm this type of specificity: a methyl group presence at the cyto‐ sine C5 atom would result in steric clashes between the DNA and the protein atoms [9]. The CXXC domain seems to bind newly synthesized unmethylated sites after DNA replication

As mentioned above, Dnmt1 is a maintenance MTase that modifies mainly monomethylated sites. A deletion of the CXXC domain and the part of the CXXC–BAH1 linker results in a 7 times decrease of the Dnmt1 affinity to monomethylated DNA relatively to unmethylated

Zn2+-binding motif – in brown, α-helical bundle – in green. Zn2+ ion is shown as a red sphere.

*5.1.4. CXXC domain*

chains and the DNA backbone.

which would protect them from *de novo* methylation.

#### *5.1.3. RFTS domain*

Besides the presence in DNA replication foci during S phase (provided by the PCNA-bind‐ ing domain) [84, 90], Dnmt1 is also associated with chromatin (mainly heterochromatin) from late S phase until early G1 phase [95]. This association is provided by the RFTS domain (replication foci targeting sequence) and does not depend on the methylation patterns spe‐ cific for heterochromatin or on the histone binding proteins. Moreover, the association with chromatin does not depend on DNA replication, since it takes place in G2 phase and M phase *de novo* [95].

The RFTS domain inhibits the Dnmt1 binding with both free DNA and nucleosomal DNA. It functions as an intrinsic competitive inhibitor of Dnmt1 and can decrease its enzymatic ac‐ tivity up to 600 times [96]. The RFTS domain also inhibits the CXXC domain binding with nucleosomal DNA. The inhibition is observed for the isolated RFTS and CXXC domains as well as for the two domains in a single polypeptide chain. However, a deletion mutant con‐ taining both the CXXC domain and the catalytic domain is able to bind polynucleosomes in the presence of the isolated RFTS domain. Thus, the simultaneous presence of the two DNA binding domains seems to make the complex relatively resistant to exclusion of DNA by the RFTS domain [96].

The RFTS domain contains a Zn2+-binding motif followed by a β-barrel and an α-helical bun‐ dle (Figure 10). Hydrophobic interactions of the RFTS domains provide Dnmt1 dimeriza‐ tion, although its functional importance is still unknown [97]. In murine Dnmt1(291–1620), the negatively charged RFTS domain penetrates deeply into the positively charged DNAbinding center of the catalytic domain and forms several hydrogen bonds inside it. Such a structural organization could explain the mechanism of DNA displacement from the catalyt‐ ic center by means of competition with the RFTS domain [87].

**Figure 10.** Spatial structure of the RFTS domain from human Dnmt1 (PDB code: 3epz). β-Barrel is colored in yellow, Zn2+-binding motif – in brown, α-helical bundle – in green. Zn2+ ion is shown as a red sphere.

#### *5.1.4. CXXC domain*

ing domain is likely to decrease Dnmt1 stability *in vivo* and thus could be involved in Dnmt1

The Dnmt1 residues 163–174 are responsible for binding of PCNA (proliferating cell nuclear antigen, also known as processivity factor for DNA polymerase δ). Dnmt1 relocates to DNA replication foci when the cell enters S phase [84, 90]. Its binding with PCNA is observed in the regions of newly synthesized DNA in intact cells. This binding does not influence the

In mammalian cells, newly replicated DNA is rapidly packaged into nucleosomes to which histone H1 is added further [91, 92]. Histone H1 has a high affinity to methylated DNA re‐ gardless its nucleotide sequence [93] and can suppress the Dnmt1 enzymatic activity [94]. Therefore, the maintenance methylation should be performed before DNA is packaged in nucleosomes. Dnmt1 binding with PCNA probably underlies a special mechanism required

Besides the presence in DNA replication foci during S phase (provided by the PCNA-bind‐ ing domain) [84, 90], Dnmt1 is also associated with chromatin (mainly heterochromatin) from late S phase until early G1 phase [95]. This association is provided by the RFTS domain (replication foci targeting sequence) and does not depend on the methylation patterns spe‐ cific for heterochromatin or on the histone binding proteins. Moreover, the association with chromatin does not depend on DNA replication, since it takes place in G2 phase and M

The RFTS domain inhibits the Dnmt1 binding with both free DNA and nucleosomal DNA. It functions as an intrinsic competitive inhibitor of Dnmt1 and can decrease its enzymatic ac‐ tivity up to 600 times [96]. The RFTS domain also inhibits the CXXC domain binding with nucleosomal DNA. The inhibition is observed for the isolated RFTS and CXXC domains as well as for the two domains in a single polypeptide chain. However, a deletion mutant con‐ taining both the CXXC domain and the catalytic domain is able to bind polynucleosomes in the presence of the isolated RFTS domain. Thus, the simultaneous presence of the two DNA binding domains seems to make the complex relatively resistant to exclusion of DNA by the

The RFTS domain contains a Zn2+-binding motif followed by a β-barrel and an α-helical bun‐ dle (Figure 10). Hydrophobic interactions of the RFTS domains provide Dnmt1 dimeriza‐ tion, although its functional importance is still unknown [97]. In murine Dnmt1(291–1620), the negatively charged RFTS domain penetrates deeply into the positively charged DNAbinding center of the catalytic domain and forms several hydrogen bonds inside it. Such a structural organization could explain the mechanism of DNA displacement from the catalyt‐

ic center by means of competition with the RFTS domain [87].

degradation.

*5.1.2. PCNA-binding domain*

MTase activity of Dnmt1 [85].

*5.1.3. RFTS domain*

phase *de novo* [95].

RFTS domain [96].

for coordination of these processes in a cell [85].

48 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

The cysteine-rich Zn2+-binding domain contains several cysteine residues organized in CXXC motifs which provide a name to the domain. The CXXC domain of Dnmt1 is similar to cysteine-rich domains of other chromatin-associated proteins such as the MeCP2 protein, the CG binding protein (CGBP), the histone MTase MLL, and the histone demethylases JHDM1A and JHDM1B. This domain is shown to bind unmethylated 5′-CG-3′/3′-GC-5′ sites *in vitro* in the case of MBD1, MLL, CGBP, JHDM1B, and Dnmt1 proteins [3]. The CXXC do‐ main of Dnmt1 is crescent-shaped and contains 8 conservative catalytically important Cys residues [98]. These residues are clustered in two groups and bind two Zn2+ ions. In the structure of murine Dnmt1(650–1602) complexed with AdoHcy and 19 bp DNA (PDB code: 3pt6), all the specific contacts with DNA are formed by the CXXC domain which interacts with both major and minor grooves [9]. A loop region of the CXXC domain (Arg684-Ser685- Lys686-Gln687) penetrates into the major groove and forms contacts with heterocyclic bases and phosphate groups. The guanine bases of the 5′-CG-3′ dinucleotide are recognized by the side chains of Lys686 and Gln687, whereas the cytosine bases are recognized by the backbone interactions of Ser685 and Lys686. Salt bridges are formed between the Arg side chains and the DNA backbone.

The CXXC domain is known to bind specifically unmethylated 5′-CG-3′/3′-GC-5′ sites [98, 99]. The structural data confirm this type of specificity: a methyl group presence at the cyto‐ sine C5 atom would result in steric clashes between the DNA and the protein atoms [9]. The CXXC domain seems to bind newly synthesized unmethylated sites after DNA replication which would protect them from *de novo* methylation.

As mentioned above, Dnmt1 is a maintenance MTase that modifies mainly monomethylated sites. A deletion of the CXXC domain and the part of the CXXC–BAH1 linker results in a 7 times decrease of the Dnmt1 affinity to monomethylated DNA relatively to unmethylated one. The same effect is observed after a substitution of two residues (K686A/Q687A) in the CXXC domain that form contacts with the guanine bases in the recognition site [9].

Addition of dimethylated DNA stimulates murine Dnmt1 to methylate unmodified sites. Such an allosteric activation of Dnmt1 results in lowering its specificity [13, 29, 34, 100]. This effect depends on the presence of Zn2+ ions and seems to be provided by the binding of the Dnmt1 residues 613–748 with dimethylated DNA [13]. This Dnmt1 region includes the CXXC domain. However, it remains unclear how the CXXC domain could bind dimethylat‐ ed DNA.

(a) (b)

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**Figure 11.** *a*) BAH1 and (*b*) BAH2 domains of murine Dnmt1 (PDB code: 3pt6) colored according to their secondary

The structure of Dnmt1(650–1602) complex with 19 bp DNA duplex and AdoHcy (PDB-код 3pt6) suggests an autoinhibition mechanism of Dnmt1: the CXXC domain binding with DNA results in DNA removal from the active center. The negatively charged CXXC–BAH1 linker is located between the DNA and the active center and thus prevents DNA entrance into the catalytic pocket (Figure 12). In addition, the BAH2–TRD loop fixes the TRD apart

**Figure 12.** A model of the Dnmt1 autoinhibition mechanism in maintenance DNA methylation. The domains are col‐ ored as in Figure 9. The CXXC domain and the autoinhibitory linker (in red) close the active center of the enzyme. Ad‐

The peptides containing the (LysX)nLys sequences (where Х = Gly, Ala or Lys) can effective‐ ly bind to Z-DNA and stabilize it even at low NaCl concentrations (10–150 mМ) and physio‐

structure. α-Helices are in red, β-strands are in yellow.

from DNA preventing its interaction with the major groove [9].

ditionally, the BAH2–TRD loop (in orange) keeps the TRD away from the DNA.

*5.1.6. KG linker*

#### *5.1.5. BAH domains*

The BAH1 and BAH2 domains (bromo-adjacent homology) are the parts of the so-called PBHD domain (polybromo homology domain). This domain is present in some transcrip‐ tion regulators and is supposed to participate in protein–protein interactions that lead to gene repression. The BAH1 and BAH2 domains in Dnmt1 molecule are connected by an αhelix and are arranged in a dumbbell shape (Figure 11). Three Cys and one His residues co‐ ordinate a Zn2+ ion that keeps the BAH1 domain near the linker α-helix. Despite low similarity of the primary structures, the both BAH domains have the same fold (Figure 11): the N-terminal subdomain consists of three antiparallel β-strands, the following subdomain consists of five antiparallel β-strands. Some smaller β-strands and loops which are located further are not homologous in different BAH domains.

Both BAH domains are physically associated with the MTase domain. Seven β-strands of the catalytic subdomain and two β-strands of the BAH1 domain form a common β-sheet. The BAH2 domain has a long loop (BAH2–TRD loop) which interacts with the TRD region of the MTase domain (Figure 11). Perhaps, this interaction prevents the TRD binding to DNA in the complex of murine Dnmt1(650–1602) with AdoHcy and 19 bp DNA duplex (PDB code: 3pt6) (Figure 9). The BAH1 and BAH2 domains have large solvent-accessible surfaces and thus could serve as platforms for interaction with other proteins.

Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization http://dx.doi.org/10.5772/52046 51

**Figure 11.** *a*) BAH1 and (*b*) BAH2 domains of murine Dnmt1 (PDB code: 3pt6) colored according to their secondary structure. α-Helices are in red, β-strands are in yellow.

The structure of Dnmt1(650–1602) complex with 19 bp DNA duplex and AdoHcy (PDB-код 3pt6) suggests an autoinhibition mechanism of Dnmt1: the CXXC domain binding with DNA results in DNA removal from the active center. The negatively charged CXXC–BAH1 linker is located between the DNA and the active center and thus prevents DNA entrance into the catalytic pocket (Figure 12). In addition, the BAH2–TRD loop fixes the TRD apart from DNA preventing its interaction with the major groove [9].

**Figure 12.** A model of the Dnmt1 autoinhibition mechanism in maintenance DNA methylation. The domains are col‐ ored as in Figure 9. The CXXC domain and the autoinhibitory linker (in red) close the active center of the enzyme. Ad‐ ditionally, the BAH2–TRD loop (in orange) keeps the TRD away from the DNA.

#### *5.1.6. KG linker*

one. The same effect is observed after a substitution of two residues (K686A/Q687A) in the

Addition of dimethylated DNA stimulates murine Dnmt1 to methylate unmodified sites.

Such an allosteric activation of Dnmt1 results in lowering its specificity [13, 29, 34, 100]. This

effect depends on the presence of Zn2+ ions and seems to be provided by the binding of the

Dnmt1 residues 613–748 with dimethylated DNA [13]. This Dnmt1 region includes the

CXXC domain. However, it remains unclear how the CXXC domain could bind dimethylat‐

The BAH1 and BAH2 domains (bromo-adjacent homology) are the parts of the so-called

PBHD domain (polybromo homology domain). This domain is present in some transcrip‐

tion regulators and is supposed to participate in protein–protein interactions that lead to

gene repression. The BAH1 and BAH2 domains in Dnmt1 molecule are connected by an α-

helix and are arranged in a dumbbell shape (Figure 11). Three Cys and one His residues co‐

ordinate a Zn2+ ion that keeps the BAH1 domain near the linker α-helix. Despite low

similarity of the primary structures, the both BAH domains have the same fold (Figure 11):

the N-terminal subdomain consists of three antiparallel β-strands, the following subdomain

consists of five antiparallel β-strands. Some smaller β-strands and loops which are located

Both BAH domains are physically associated with the MTase domain. Seven β-strands of

the catalytic subdomain and two β-strands of the BAH1 domain form a common β-sheet.

The BAH2 domain has a long loop (BAH2–TRD loop) which interacts with the TRD region

of the MTase domain (Figure 11). Perhaps, this interaction prevents the TRD binding to

DNA in the complex of murine Dnmt1(650–1602) with AdoHcy and 19 bp DNA duplex

(PDB code: 3pt6) (Figure 9). The BAH1 and BAH2 domains have large solvent-accessible

surfaces and thus could serve as platforms for interaction with other proteins.

further are not homologous in different BAH domains.

ed DNA.

*5.1.5. BAH domains*

CXXC domain that form contacts with the guanine bases in the recognition site [9].

50 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

The peptides containing the (LysX)nLys sequences (where Х = Gly, Ala or Lys) can effective‐ ly bind to Z-DNA and stabilize it even at low NaCl concentrations (10–150 mМ) and physio‐

logical pH. Such peptides can also induce B-DNA transition into Z-DNA. The efficacy of this process grows as the number of repeats (*n*) increases [101, 102]. The sequences (LysX)nLys are found in different proteins of plants, animals and unicellular eukaryotes. For example, they are present in the linkers which connect N- and C-terminal parts in eukaryotic MTases [102-104]. The number of repeats (*n*) varies from 5 to 7 in this case. The sequences of the linker and the preceding protein region are highly conservative. Thus, the linker is sup‐ posed to have an important but still unknown function.

*5.2.1. PWWP domain*

helices [120].

45 × 32 Å2

Dnmt3a and Dnmt3b contain a PWWP domain as well as some other chromatin-associated proteins. The PWWP domain is named after a conserved ProTrpTrpPro motif, though the first Pro is substituted with Ser in Dnmt3a and Dnmt3b. The second part of the motif is al‐ ways the same, TrpPro [117]. PWWP domain along with chromo domain, Tudor and MBT domains belongs to the Tudor domain "Royal family" [118, 119]. The members of this family are shown to bind modified lysine residues of histones [119]. The PWWP domain consists of 100–130 amino acid residues. Its structure includes 3 motifs: a β-barrel core, an insertion be‐ tween the second and the third β-strands, and a C-terminal α-helical bundle (Figure 13). Three aromatic residues form a cleft in the center of the β-barrel that is a distinctive feature of the "Royal family". The insertion motif varies in length and secondary structure among the different PWWP domains. The C-terminal α-helical bundle can contain from 1 to 5 α-

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The PWWP domain of Dnmt3b has a positively charged surface with an approximate area of

duplexes with unmethylated, mono-, and dimethylated 5′-CG-3′/3′-GC-5′ sites. However, it can also bind a nonspecific duplex of the same length with the same efficiency [117]. Dele‐ tion of the PWWP domain does not influence the Dnmt3b methylation efficiency *in vitro* [117]. On the contrary, the PWWP domain of Dnmt3a is almost unable to bind DNA [121].

**Figure 13.** Spatial structures of the PWWP domains from human Dnmt3a (PDB code: 3llr) and Dnmt3b colored ac‐

The PWWP domains of Dnmt3a and Dnmt3b are necessary for targeting these MTases to pericentromeric heterochromatin [121-123]. Deletions in the PWWP domain change the pro‐ tein distribution in a nucleus and therefore result in its disability to methylate satellite re‐ peats. However, such mutants are catalytically active since they are able to methylate DNA in other regions [121]. In humans, S282P point mutation in the PWWP domain of Dnmt3b

cording to their secondary structure (PDB code: 3qkj). α-Helices are in red, β-strands are in yellow.

and can bind DNA nonspecifically [117, 121]. The PWWP domain binds 30 bp

The most effective DNA transition into Z-form is observed for the sequences with alternat‐ ing purine and pyrimidine nucleosides, including 5′-(CG)n-3′ and especially when the cyto‐ sine residues are methylated at the C5 atoms. Additionally, the transition into Z-form is stimulated by high ionic strength in solution and by DNA negative supercoiling [105]. The KG linker probably promotes Dnmt1 binding to 5′-CG-3′ islands in Z-form. However, Dnmt1 is not able to methylate Z-DNA [106]. Perhaps, the KG linker participates in Dnmt1 targeting to the regions located behind a replication fork (due to their negative supercoiling when DNA polymerase has just passed) [3].

#### **5.2. Functional domains of mammalian Dnmt3 family**

Dnmt3 family includes C5-DNA MTases Dnmt3a and Dnmt3b, which are considered *de novo* MTases, and Dnmt3L – a catalytically inactive protein. There are also different isoforms of the proteins in the Dnmt3 family. Dnmt3a and Dnmt3b are expressed in embryonic cells and during gametogenesis [107]. Knockout of the corresponding genes suppresses *de novo* meth‐ ylation. Mouse embryos where the Dnmt3b gene is knocked out die *in utero* while mouse embryos lacking Dnmt3a gene die soon after birth [108]. Dnmt3b methylates microsatellite repeats. In humans, Dnmt3b point mutations decreasing its enzymatic activity lead to a se‐ vere disease – ICF syndrome (immunodeficiency, centromere instability, facial abnormalities syndrome) [108-110].

Like Dnmt1, the Dnmt3 family enzymes consist of the N-terminal regulatory part and the Cterminal part containing the conservative motifs typical for C5-DNA MTases (Figure 8). The catalytic domains of Dnmt1 and Dnmt3 enzymes are homologous while the N-terminal parts share no similarity. Thus, these families seem to have been evolved from different pro‐ karyotic predecessors [111, 112]. The intramolecular interactions between the N-terminal and the C-terminal parts are absent in Dnmt3a and Dnmt3b, in contrast to Dnmt1 [12].

The N-terminal parts of Dnmt3a and Dnmt3b contain two domains:


There is no PWWP domain in Dnmt3L. Moreover, Dnmt3L lacks some catalytic residues and the MTase motifs IX and X in the C-terminal domain. Dnmt3L functions as a stimulator of Dnmt3a and Dnmt3b enzymatic activity. Dnmt3L knockout mice are viable but the males are sterile while the females do not produce viable offspring [113-116].

#### *5.2.1. PWWP domain*

logical pH. Such peptides can also induce B-DNA transition into Z-DNA. The efficacy of this process grows as the number of repeats (*n*) increases [101, 102]. The sequences (LysX)nLys are found in different proteins of plants, animals and unicellular eukaryotes. For example, they are present in the linkers which connect N- and C-terminal parts in eukaryotic MTases [102-104]. The number of repeats (*n*) varies from 5 to 7 in this case. The sequences of the linker and the preceding protein region are highly conservative. Thus, the linker is sup‐

The most effective DNA transition into Z-form is observed for the sequences with alternat‐ ing purine and pyrimidine nucleosides, including 5′-(CG)n-3′ and especially when the cyto‐ sine residues are methylated at the C5 atoms. Additionally, the transition into Z-form is stimulated by high ionic strength in solution and by DNA negative supercoiling [105]. The KG linker probably promotes Dnmt1 binding to 5′-CG-3′ islands in Z-form. However, Dnmt1 is not able to methylate Z-DNA [106]. Perhaps, the KG linker participates in Dnmt1 targeting to the regions located behind a replication fork (due to their negative supercoiling

Dnmt3 family includes C5-DNA MTases Dnmt3a and Dnmt3b, which are considered *de novo* MTases, and Dnmt3L – a catalytically inactive protein. There are also different isoforms of the proteins in the Dnmt3 family. Dnmt3a and Dnmt3b are expressed in embryonic cells and during gametogenesis [107]. Knockout of the corresponding genes suppresses *de novo* meth‐ ylation. Mouse embryos where the Dnmt3b gene is knocked out die *in utero* while mouse embryos lacking Dnmt3a gene die soon after birth [108]. Dnmt3b methylates microsatellite repeats. In humans, Dnmt3b point mutations decreasing its enzymatic activity lead to a se‐ vere disease – ICF syndrome (immunodeficiency, centromere instability, facial abnormalities

Like Dnmt1, the Dnmt3 family enzymes consist of the N-terminal regulatory part and the Cterminal part containing the conservative motifs typical for C5-DNA MTases (Figure 8). The catalytic domains of Dnmt1 and Dnmt3 enzymes are homologous while the N-terminal parts share no similarity. Thus, these families seem to have been evolved from different pro‐ karyotic predecessors [111, 112]. The intramolecular interactions between the N-terminal and the C-terminal parts are absent in Dnmt3a and Dnmt3b, in contrast to Dnmt1 [12].

**1.** cysteine-rich domain ADD (ATRX–Dnmt3–Dnmt3L), also called PHD domain (plant

There is no PWWP domain in Dnmt3L. Moreover, Dnmt3L lacks some catalytic residues and the MTase motifs IX and X in the C-terminal domain. Dnmt3L functions as a stimulator of Dnmt3a and Dnmt3b enzymatic activity. Dnmt3L knockout mice are viable but the males

The N-terminal parts of Dnmt3a and Dnmt3b contain two domains:

are sterile while the females do not produce viable offspring [113-116].

posed to have an important but still unknown function.

52 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**5.2. Functional domains of mammalian Dnmt3 family**

when DNA polymerase has just passed) [3].

syndrome) [108-110].

homeodomain);

**2.** PWWP domain.

Dnmt3a and Dnmt3b contain a PWWP domain as well as some other chromatin-associated proteins. The PWWP domain is named after a conserved ProTrpTrpPro motif, though the first Pro is substituted with Ser in Dnmt3a and Dnmt3b. The second part of the motif is al‐ ways the same, TrpPro [117]. PWWP domain along with chromo domain, Tudor and MBT domains belongs to the Tudor domain "Royal family" [118, 119]. The members of this family are shown to bind modified lysine residues of histones [119]. The PWWP domain consists of 100–130 amino acid residues. Its structure includes 3 motifs: a β-barrel core, an insertion be‐ tween the second and the third β-strands, and a C-terminal α-helical bundle (Figure 13). Three aromatic residues form a cleft in the center of the β-barrel that is a distinctive feature of the "Royal family". The insertion motif varies in length and secondary structure among the different PWWP domains. The C-terminal α-helical bundle can contain from 1 to 5 αhelices [120].

The PWWP domain of Dnmt3b has a positively charged surface with an approximate area of 45 × 32 Å2 and can bind DNA nonspecifically [117, 121]. The PWWP domain binds 30 bp duplexes with unmethylated, mono-, and dimethylated 5′-CG-3′/3′-GC-5′ sites. However, it can also bind a nonspecific duplex of the same length with the same efficiency [117]. Dele‐ tion of the PWWP domain does not influence the Dnmt3b methylation efficiency *in vitro* [117]. On the contrary, the PWWP domain of Dnmt3a is almost unable to bind DNA [121].

**Figure 13.** Spatial structures of the PWWP domains from human Dnmt3a (PDB code: 3llr) and Dnmt3b colored ac‐ cording to their secondary structure (PDB code: 3qkj). α-Helices are in red, β-strands are in yellow.

The PWWP domains of Dnmt3a and Dnmt3b are necessary for targeting these MTases to pericentromeric heterochromatin [121-123]. Deletions in the PWWP domain change the pro‐ tein distribution in a nucleus and therefore result in its disability to methylate satellite re‐ peats. However, such mutants are catalytically active since they are able to methylate DNA in other regions [121]. In humans, S282P point mutation in the PWWP domain of Dnmt3b causes the ICF syndrome [124]. In this case, the disease is likely to be caused by the enzyme improper distribution in nuclei rather than by its insufficient catalytic activity.

domains of Dnmt3a, Dnmt3b, and Dnmt3L are shown to interact specifically with the N-ter‐ minal part of H3 histone when its Lys4 is not modified. This interaction stimulates Dnmt3a to methylate DNA [130-132]. Thus, the ADD domain of C5-DNA MTases can induce DNA

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Plant DNA MTases are very diverse but are studied much less than the mammalian DNA MTases up to now [133]. In particular, none of the plant MTases has been crystallized.

DNA MTase Met1 from *Arabidopsis thaliana* is quite similar to mammalian Dnmt1 (Figure 15). The C-terminal domains of these enzymes which are responsible for methylation share 50% identity. Both proteins possess an extended N-terminal part that is connected with the C-terminal domain *via* KG linker. The N-terminal parts of Met1 and Dnmt1 share 24% iden‐ tity [134]. There are four similar genes encoding Met1 in *A. thaliana* whereas only one gene encodes the mammalian Dnmt1*.* Genomes of *Daucus carota* and *Zea mays* contain two Met1 homologs. The N-terminal part of Met1 has two BAH domains. These domains seem to serve as a platform for protein–protein interactions which result in inhibition of gene expres‐ sion thus providing an interconnection of DNA methylation, replication, and transcription

**Figure 15.** Domain architecture of different С5-DNA MTases from *A. thaliana*. Met1 – MTase similar to mammalian Dnmt1, CMT3 – chromomethyltransferase 3, DRM2 – MTase with the circular permutation of conservative motifs. The catalytic domains are marked by a light blue filling. The other domains are signed in the bottom. Roman numerals

Dnmt3 family homologs are found in plants also. They form a DRM family (domains rear‐ ranged methyltransferase). The peculiarity of these enzymes is a circular permutation of their conservative motifs: the motifs VI–X are followed by the motifs I–V. The circular per‐ mutation of the conservative motifs is also found in some prokaryotic enzymes such as C5- DNA MTase BssHII [136]. Another characteristic of the DRM proteins is their ability to

methylation in response to specific histone modifications.

**5.3. Domains of plant DNA methyltransferases**

regulation [135].

indicate the conservative motifs of C5-DNA MTases.

The PWWP domain of Dnmt3a specifically binds trimethylated Lys36 of H3 histone (H3K36m3) that increases the Dnmt3a ability to methylate nucleosomal DNA [125]. Distri‐ bution of methylated sequences in a genome correlates with the presence of H3K36m3 [126, 127]. DNA methylation and H3K36m3 serve as marks for histone deacetylation and the fol‐ lowing gene suppression [128]. There is no crystal structure of Dnmt3a or Dnmt3b PWWP domain in complex with a histone protein or a peptide. The resolved spatial structures have a cleft that is supposed to bind methylated lysine residues. This cleft in the crystal contains a molecule of Bis-tris buffer (bis(2-hydroxyethyl)amino-tris(hydroxymethyl)methane) that is situated similar to the lysine in complexes of other PWWP domains with histone peptides containing di- or trimethylated lysine residues [120].

#### *5.2.2. ADD domain*

ADD domain is found only in the following proteins: ATRX (alpha thalassemia/mental re‐ tardation syndrome X-linked), Dnmt3a, Dnmt3b, and Dnmt3L. Thus, the domain is called ADD (ATRX–Dnmt3–Dnmt3L) [129]. To date, the crystal structures of ADD domain from Dnmt3a, Dnmt3L, and ATRX proteins are resolved. The structural organization of the do‐ main remains the same in all the cases: it contains two C4-type zinc fingers [129-131]. One of them is similar to GATA binding protein 1 (GATA1) and the other one – to plant homeodo‐ main (PHD).

**Figure 14.** Structure of a fusion protein containing the ADD domain and a fragment of histone H3 (residues 1–9, shown in sticks representation), PDB-code: 3a1b. GATA1-like finger is in yellow, PHD finger is in green. Zn2+ ions are shown as red spheres. The disordered linker peptide that connects the C-terminus of the histone H3 fragment and the N-terminus of ADD is not shown.

The ADD domain contains many cysteine residues which bind Zn2+ providing thus the Dnmt3a interaction with many other proteins such as transcription factors PU.1, Myc, RP58, histone deacetylase HDAC1, heterochromatin protein HP1, histone MTases SUV39H1, SETDB1 and EZH2, methyl-CG-binding protein Mbd3, and chromatin remodeling factor Brg1 [3]. The functions of most of these interactions remain unclear. Additionally, the ADD domains of Dnmt3a, Dnmt3b, and Dnmt3L are shown to interact specifically with the N-ter‐ minal part of H3 histone when its Lys4 is not modified. This interaction stimulates Dnmt3a to methylate DNA [130-132]. Thus, the ADD domain of C5-DNA MTases can induce DNA methylation in response to specific histone modifications.

#### **5.3. Domains of plant DNA methyltransferases**

causes the ICF syndrome [124]. In this case, the disease is likely to be caused by the enzyme

The PWWP domain of Dnmt3a specifically binds trimethylated Lys36 of H3 histone (H3K36m3) that increases the Dnmt3a ability to methylate nucleosomal DNA [125]. Distri‐ bution of methylated sequences in a genome correlates with the presence of H3K36m3 [126, 127]. DNA methylation and H3K36m3 serve as marks for histone deacetylation and the fol‐ lowing gene suppression [128]. There is no crystal structure of Dnmt3a or Dnmt3b PWWP domain in complex with a histone protein or a peptide. The resolved spatial structures have a cleft that is supposed to bind methylated lysine residues. This cleft in the crystal contains a molecule of Bis-tris buffer (bis(2-hydroxyethyl)amino-tris(hydroxymethyl)methane) that is situated similar to the lysine in complexes of other PWWP domains with histone peptides

ADD domain is found only in the following proteins: ATRX (alpha thalassemia/mental re‐ tardation syndrome X-linked), Dnmt3a, Dnmt3b, and Dnmt3L. Thus, the domain is called ADD (ATRX–Dnmt3–Dnmt3L) [129]. To date, the crystal structures of ADD domain from Dnmt3a, Dnmt3L, and ATRX proteins are resolved. The structural organization of the do‐ main remains the same in all the cases: it contains two C4-type zinc fingers [129-131]. One of them is similar to GATA binding protein 1 (GATA1) and the other one – to plant homeodo‐

**Figure 14.** Structure of a fusion protein containing the ADD domain and a fragment of histone H3 (residues 1–9, shown in sticks representation), PDB-code: 3a1b. GATA1-like finger is in yellow, PHD finger is in green. Zn2+ ions are shown as red spheres. The disordered linker peptide that connects the C-terminus of the histone H3 fragment and the

The ADD domain contains many cysteine residues which bind Zn2+ providing thus the Dnmt3a interaction with many other proteins such as transcription factors PU.1, Myc, RP58, histone deacetylase HDAC1, heterochromatin protein HP1, histone MTases SUV39H1, SETDB1 and EZH2, methyl-CG-binding protein Mbd3, and chromatin remodeling factor Brg1 [3]. The functions of most of these interactions remain unclear. Additionally, the ADD

improper distribution in nuclei rather than by its insufficient catalytic activity.

containing di- or trimethylated lysine residues [120].

54 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

*5.2.2. ADD domain*

main (PHD).

N-terminus of ADD is not shown.

Plant DNA MTases are very diverse but are studied much less than the mammalian DNA MTases up to now [133]. In particular, none of the plant MTases has been crystallized.

DNA MTase Met1 from *Arabidopsis thaliana* is quite similar to mammalian Dnmt1 (Figure 15). The C-terminal domains of these enzymes which are responsible for methylation share 50% identity. Both proteins possess an extended N-terminal part that is connected with the C-terminal domain *via* KG linker. The N-terminal parts of Met1 and Dnmt1 share 24% iden‐ tity [134]. There are four similar genes encoding Met1 in *A. thaliana* whereas only one gene encodes the mammalian Dnmt1*.* Genomes of *Daucus carota* and *Zea mays* contain two Met1 homologs. The N-terminal part of Met1 has two BAH domains. These domains seem to serve as a platform for protein–protein interactions which result in inhibition of gene expres‐ sion thus providing an interconnection of DNA methylation, replication, and transcription regulation [135].

**Figure 15.** Domain architecture of different С5-DNA MTases from *A. thaliana*. Met1 – MTase similar to mammalian Dnmt1, CMT3 – chromomethyltransferase 3, DRM2 – MTase with the circular permutation of conservative motifs. The catalytic domains are marked by a light blue filling. The other domains are signed in the bottom. Roman numerals indicate the conservative motifs of C5-DNA MTases.

Dnmt3 family homologs are found in plants also. They form a DRM family (domains rear‐ ranged methyltransferase). The peculiarity of these enzymes is a circular permutation of their conservative motifs: the motifs VI–X are followed by the motifs I–V. The circular per‐ mutation of the conservative motifs is also found in some prokaryotic enzymes such as C5- DNA MTase BssHII [136]. Another characteristic of the DRM proteins is their ability to methylate 5′-CHG-3′ and 5′-CHH-3′ sites (H = A, C or T) *de novo* in an RNA-dependent manner. Such DNA methylation is supposed to take place in the presence of short RNAs that guide methylation of homologous DNA [4]. The DRM1 MTase from *Nicotiana taba‐ cum* is shown to avoid cytosine methylation in 5′-CG-3′/3′-GC-5′ sites rather than specifi‐ cally recognize any sequence. A structural basis of such an unusual functioning is not yet clarified [137].

**6. Conclusion**

activity *etc*.

**Acknowledgements**

and 12-04-32103).

**Author details**

A. Yu. Ryazanova1

cow, Russia

Moscow, Russia

Precise time and space coordination of different molecular events underlies development of all living organisms, unicellular as well as multicellular. Synchronization of molecular processes can take place at a transcriptional level (when the same DNA-binding protein regulates expression of several genes) or at a post-translational level (when a multifunction‐ al protein participates in different processes). Multifunctionality of a protein can be based on the presence of several domains in a single polypeptide chain. For example, mammali‐ an Dnmt1, besides the catalytic domain which provides DNA methylation, contains sever‐ al other domains responsible for Dnmt1 cellular localization, its interaction with other proteins, and regulation of its catalytic domain activity. Among prokaryotes, multido‐ main proteins are less common. Nevertheless, some bacterial DNA methyltransferases con‐ tain additional domains which are responsible for transcription regulation, topoisomerase

Diverse Domains of (Cytosine-5)-DNA Methyltransferases: Structural and Functional Characterization

http://dx.doi.org/10.5772/52046

57

As shown in this review, the structural and functional features of the additional domains in C5-DNA MTases are studied yet insufficiently. On the basis of the existing data, it is impos‐ sible to draw a decisive conclusion on the effect of the additional domains onto the methyl‐ ating activity of C5-DNA MTases. Evidently, a complex research of multifunctional DNA

The authors would like to thank Mrs. Anna Nazarenko for her technical assistance. The work was supported by the Russian Foundation for Basic Research (grants no. 10-04-01578

, T. S. Oretskaya1,3 and E. A. Kubareva3\*

MTases with multidomain organization would be most promising.

, L. A. Abrosimova2

\*Address all correspondence to: kubareva@belozersky.msu.ru

1 Chemistry Department, Lomonosov Moscow State University, Moscow, Russia

2 Faculty of Bioengineering and Bioinformatics, Lomonosov Moscow State University, Mos‐

3 Belozersky Institute of Physico-Chemical Biology, Lomonosov Moscow State University,

Proteins of the DRM family are found only in flowering plants [4]. The C-terminal do‐ mains of DRM proteins share 28% identity with Dnmt3a or Dnmt3b and contain the same catalytic conservative motifs. Cysteine-rich regions of mammalian MTases (the RFTS, the СХХС, and the ADD domains) are found neither in proteins of DRM family nor in other plant homologs of C5-DNA MTases [112]. The N-terminal part of the DRM MTases con‐ tains several UBA domains (ubiquitin-associated). The presence of UBA domains seems to provide a link between DNA methylation and ubiquitin-mediated protein degradation. These domains could promote degradation of DRM molecules at specific points of the cell cycle [112]. The UBA domains are not found in any other families of DNA MTases. The UBA domains of DRM2 MTase in *A. thaliana* were experimentally shown to be required for normal RNA-directed DNA methylation. Perhaps, these domains are essential for proper localization of MTases in the cell [138].

C5-DNA MTases of chromomethylase (or chromomethyltransferase, CMT) family modify 5′-CNG-3′/3′-GNC-5′ sites (N = A, C, G or T) in plant genomes. These MTases contain chro‐ mo domains that were identified as conserved sequences between the II and the IV MTase motifs (Figure 15). The chromo domain (chromatin organization modifier) consists of 50 amino acid residues and contains three β-strands and a perpendicularly packed α-helix. This folding type belongs to OB class (oligonucleotide/oligosaccharide binding fold) and is considered to be evolutionary very old. The chromo domain is responsible for DNA binding [139]. This domain is originally found in polycomb-group proteins where it is important for the protein association with heterochromatin [140]. Therefore, CMT are thought to modify heterochromatin. The chromo domain is not found in other C5-DNA MTases. The *A. thaliana* genome contains 3 members of the CMT family: CMT1, CMT2, and CMT3. The members of this family are also found in genomes of *Oryza sativa* and *Brassica oleracea*. Function of CMT1 and CMT2 are still unknown while CMT3 seems to methylate 5′-CNG-3′/3′-GNC-5′ sites [4]. CMT3 deficiency in *A. thaliana* results in loss of DNA methylation in centromeric regions and also leads to retrotransposon activation [141, 142].

The enzymes from CMT family contain one BAH domain in its N-terminal region in con‐ trast to the MTases from Met1 family that possess two BAH domains.

#### **6. Conclusion**

methylate 5′-CHG-3′ and 5′-CHH-3′ sites (H = A, C or T) *de novo* in an RNA-dependent manner. Such DNA methylation is supposed to take place in the presence of short RNAs that guide methylation of homologous DNA [4]. The DRM1 MTase from *Nicotiana taba‐ cum* is shown to avoid cytosine methylation in 5′-CG-3′/3′-GC-5′ sites rather than specifi‐ cally recognize any sequence. A structural basis of such an unusual functioning is not yet

Proteins of the DRM family are found only in flowering plants [4]. The C-terminal do‐ mains of DRM proteins share 28% identity with Dnmt3a or Dnmt3b and contain the same catalytic conservative motifs. Cysteine-rich regions of mammalian MTases (the RFTS, the СХХС, and the ADD domains) are found neither in proteins of DRM family nor in other plant homologs of C5-DNA MTases [112]. The N-terminal part of the DRM MTases con‐ tains several UBA domains (ubiquitin-associated). The presence of UBA domains seems to provide a link between DNA methylation and ubiquitin-mediated protein degradation. These domains could promote degradation of DRM molecules at specific points of the cell cycle [112]. The UBA domains are not found in any other families of DNA MTases. The UBA domains of DRM2 MTase in *A. thaliana* were experimentally shown to be required for normal RNA-directed DNA methylation. Perhaps, these domains are essential for proper

C5-DNA MTases of chromomethylase (or chromomethyltransferase, CMT) family modify 5′-CNG-3′/3′-GNC-5′ sites (N = A, C, G or T) in plant genomes. These MTases contain chro‐ mo domains that were identified as conserved sequences between the II and the IV MTase motifs (Figure 15). The chromo domain (chromatin organization modifier) consists of 50 amino acid residues and contains three β-strands and a perpendicularly packed α-helix. This folding type belongs to OB class (oligonucleotide/oligosaccharide binding fold) and is considered to be evolutionary very old. The chromo domain is responsible for DNA binding [139]. This domain is originally found in polycomb-group proteins where it is important for the protein association with heterochromatin [140]. Therefore, CMT are thought to modify heterochromatin. The chromo domain is not found in other C5-DNA MTases. The *A. thaliana* genome contains 3 members of the CMT family: CMT1, CMT2, and CMT3. The members of this family are also found in genomes of *Oryza sativa* and *Brassica oleracea*. Function of CMT1 and CMT2 are still unknown while CMT3 seems to methylate 5′-CNG-3′/3′-GNC-5′ sites [4]. CMT3 deficiency in *A. thaliana* results in loss of DNA methylation in centromeric regions

The enzymes from CMT family contain one BAH domain in its N-terminal region in con‐

clarified [137].

localization of MTases in the cell [138].

56 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

and also leads to retrotransposon activation [141, 142].

trast to the MTases from Met1 family that possess two BAH domains.

Precise time and space coordination of different molecular events underlies development of all living organisms, unicellular as well as multicellular. Synchronization of molecular processes can take place at a transcriptional level (when the same DNA-binding protein regulates expression of several genes) or at a post-translational level (when a multifunction‐ al protein participates in different processes). Multifunctionality of a protein can be based on the presence of several domains in a single polypeptide chain. For example, mammali‐ an Dnmt1, besides the catalytic domain which provides DNA methylation, contains sever‐ al other domains responsible for Dnmt1 cellular localization, its interaction with other proteins, and regulation of its catalytic domain activity. Among prokaryotes, multido‐ main proteins are less common. Nevertheless, some bacterial DNA methyltransferases con‐ tain additional domains which are responsible for transcription regulation, topoisomerase activity *etc*.

As shown in this review, the structural and functional features of the additional domains in C5-DNA MTases are studied yet insufficiently. On the basis of the existing data, it is impos‐ sible to draw a decisive conclusion on the effect of the additional domains onto the methyl‐ ating activity of C5-DNA MTases. Evidently, a complex research of multifunctional DNA MTases with multidomain organization would be most promising.

#### **Acknowledgements**

The authors would like to thank Mrs. Anna Nazarenko for her technical assistance. The work was supported by the Russian Foundation for Basic Research (grants no. 10-04-01578 and 12-04-32103).

#### **Author details**

A. Yu. Ryazanova1 , L. A. Abrosimova2 , T. S. Oretskaya1,3 and E. A. Kubareva3\*

\*Address all correspondence to: kubareva@belozersky.msu.ru

1 Chemistry Department, Lomonosov Moscow State University, Moscow, Russia

2 Faculty of Bioengineering and Bioinformatics, Lomonosov Moscow State University, Mos‐ cow, Russia

3 Belozersky Institute of Physico-Chemical Biology, Lomonosov Moscow State University, Moscow, Russia

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**Chapter 3**

**Bifunctional Prokaryotic DNA-Methyltransferases**

Restriction-modification systems (RMS) are prokaryotic tools against invasion of foreign DNAs into cells [1]. They reduce horizontal gene transfer, thus stimulating microbial biodi‐ versity. Usually, they consist of a restriction endonuclease (REase) and a modification DNA methyltransferase (MTase) enzyme recognising the same short 4-8 nucleotide sequence. MTase is responsible for methyl group transfer to adenine or cytosine nucleotides within the target sequence, thus preventing its hydrolysis by cognate REase. Up to now, more than 20 000 different RMS have been collected in the REBASE, the database holding all known, and many putative, RMS [2]. Many of these RMS have head-to-tail gene orientation, thus provid‐ ing, by our hypothesis, the possibility of gene fusion through point mutations or genome re‐ arrangements such as deletions, insertions, inversions or translocations. These events could be responsible for the origin of bifunctional restriction enzymes of type IIC [3] such as AloI, BcgI, BseMII, BseRI, BsgI, BspLU11III, CjeI, Eco57I, HaeIV, MmeI, PpiI, TstI and TspWGI;

bifunctional MTases such as FokI and LlaI, and regulatory SsoII-related MTases [4].

In our previous work we proved the possibility of fully functional hybrid polypeptide origin through gene fusion, taking as an example Eco29kI RMS. In the given RMS, the REase gene precedes MTase gene and their Stop and Start codons overlap. By site-directed mutagenesis we joined these two ORFs into one and characterised the resulting protein, carrying both REase and MTase activities [5]. Its REase activity was decreased three times and the optima of the catalytic reaction changed, whereas MTase activity turned out to be intact [5-7]. The bifunctional enzyme could be changed as a result of evolution, leading to further divergence of its properties and functions in the cell. By our hypothesis, this example could serve as a molecular mechanism of new bifunctional RMS origin. In the current work, based on ge‐ nomic data and their bioinformatics analysis, we aimed to prove that gene fusion could play

> © 2013 Nikitin et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

Dmitry V. Nikitin, Attila Kertesz-Farkas,

http://dx.doi.org/10.5772/51025

**1. Introduction**

Alexander S. Solonin and Marina L. Mokrishcheva

Additional information is available at the end of the chapter

### **Bifunctional Prokaryotic DNA-Methyltransferases**

Dmitry V. Nikitin, Attila Kertesz-Farkas, Alexander S. Solonin and Marina L. Mokrishcheva

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51025

#### **1. Introduction**

Restriction-modification systems (RMS) are prokaryotic tools against invasion of foreign DNAs into cells [1]. They reduce horizontal gene transfer, thus stimulating microbial biodi‐ versity. Usually, they consist of a restriction endonuclease (REase) and a modification DNA methyltransferase (MTase) enzyme recognising the same short 4-8 nucleotide sequence. MTase is responsible for methyl group transfer to adenine or cytosine nucleotides within the target sequence, thus preventing its hydrolysis by cognate REase. Up to now, more than 20 000 different RMS have been collected in the REBASE, the database holding all known, and many putative, RMS [2]. Many of these RMS have head-to-tail gene orientation, thus provid‐ ing, by our hypothesis, the possibility of gene fusion through point mutations or genome re‐ arrangements such as deletions, insertions, inversions or translocations. These events could be responsible for the origin of bifunctional restriction enzymes of type IIC [3] such as AloI, BcgI, BseMII, BseRI, BsgI, BspLU11III, CjeI, Eco57I, HaeIV, MmeI, PpiI, TstI and TspWGI; bifunctional MTases such as FokI and LlaI, and regulatory SsoII-related MTases [4].

In our previous work we proved the possibility of fully functional hybrid polypeptide origin through gene fusion, taking as an example Eco29kI RMS. In the given RMS, the REase gene precedes MTase gene and their Stop and Start codons overlap. By site-directed mutagenesis we joined these two ORFs into one and characterised the resulting protein, carrying both REase and MTase activities [5]. Its REase activity was decreased three times and the optima of the catalytic reaction changed, whereas MTase activity turned out to be intact [5-7]. The bifunctional enzyme could be changed as a result of evolution, leading to further divergence of its properties and functions in the cell. By our hypothesis, this example could serve as a molecular mechanism of new bifunctional RMS origin. In the current work, based on ge‐ nomic data and their bioinformatics analysis, we aimed to prove that gene fusion could play

© 2013 Nikitin et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

an important role in evolution of metyltransferases and in origin of multidomain eukaryotic methyltransferases.

nit is the specificity subunit that determines which DNA sequence is recognised. The R sub‐ unit is essential for cleavage (restriction) and the M subunit catalyses the methylation reaction. Their protein products are marked as HsdS, HsdR and HsdM, respectively. Cova‐ lent linking of these subunits in one polypeptide is not thought to interfere with their cata‐ lytic activities, giving an opportunity for successful fusion. Currently, the REBASE contains more than 8000 entries corresponding to Type I RMS. Hypothetically, any of these RMS

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One RMS from *Bacteroides sp*. D22, probably originated from the fusion of type III enzymes. It has conserved motifs similar to Eco57I protein, which consists of Mod and Res subunits of type III enzymes [15]. Type III systems are composed of two genes (mod and res) encoding protein subunits that function in one protein complex either in DNA recognition and modi‐ fication, Mod, or restriction, Res [3]. As in the case of the type I enzymes, in-frame fusion might not affect their normal functioning, thus, being also favourable for new bifunctional

One of the found RMS from *Arthrospira maxima* CS-328 has a different domain organisation, belonging to R-M type of structure (type II REase and MTase fusion). It could be suggested that it originated from fusion of head-to-tail oriented Type II REase and MTase genes. The principal possibility of a new bifunctional RMS origin by this mechanism was proven by inframe joining of type IIEco29kI REase and MTase genes [5]. The resulting RMS was capable of defending host cells from phage invasion, although 100 times less effectively than the wild type. In a similar way, a new bifunctional RMS could appear from other head-to-tail oriented RMS of type II such as AccI, BanI, Bsp6I, BsuBI, Cfr9I, DdeI, EagI, EcoPI, EcoP15, EcoRI, FnuDI, HaeIII, HgiBI, HgiCI, HgiCII, HgiDI, HgiEI, HgiGI, HhaII, HincII, HindIII, HinfI, HpaI, MboII, MwoI, NcoI, NdeI, NgoMI, NgoPII, NlaIII, PaeR7I, RsrI, SalI, Sau3A, Sau96I, TaqI, TthHB8I, XbaI, and XmaI [16]. Type II restriction enzymes and modification enzymes work separately and their fusion could create steric difficulties for their functional‐ ity. In fact, in the case of the RM.Eco29kI enzyme, its REase activity decreased three times in comparison with initial R.Eco29kI nuclease. This is perhaps why it is the only found exam‐

could be joint, naturally or artificially, giving a new bifunctional RMS.

ple of natural bifunctional RMS originating from type II RMS.

ZP\_01728785.1, 1307 aa, type II restriction-modification enzyme, *Cyanothece sp*. CCY0110]

ZP\_00368052, 1343 aa, type I restriction modification enzyme, *Campylobacter coli* RM2228. ADC28633, 1364 aa, restriction modification enzyme, *Campylobacter jejuni subsp. Jejuni* IA3902. ZP\_01967949,1255 aa, hypothetical protein RUMTOR\_01515, *Ruminococcus torques* ATCC

YP\_892534, 1285 aa, restriction and modification enzyme, *Campylobacter fetus subsp. fetus* 82-40. YP\_001482568, 1190 aa, hypothetical protein C8J\_0992, *Campylobacter jejuni subsp. Jejuni* 81116.

protein origin.

**RMS**

27756.

For the current work we searched protein databases as described above and found 76 new potential bifunctional MTases. Their structural organisation is presented and discussed in the report. Beside this, we analysed structural organisation of 627 non-putative prokaryotic DNA-methyltransferases available out of 980 methyltransferases collected up to now in RE‐ BASE. The most frequently observed structural type, other than "canonical" MTases, repre‐ sents SsoII-related methyltransferases, capable to serve as transcriptional autoregulatory proteins. These data provide additional evidences that gene fusion might play an important role in evolution of methyltransferases, restriction-modification systems and other DNAmodifying proteins. We discuss the general consequences of a hypothetical protein fusion event with methyltransferases and RMS enzymes.

#### **1.1. Method used**

For this study 10619 methyltransferases and 3250 restriction enzyme sequences were down‐ loaded from the REBASE database [2] and searched against the non-redundant protein data‐ base which was downloaded from the NCBI's ftp site. The similarity search was carried out by BLAST version 2.2.23+ [8] on a Linux server using the BLAST default parameters. The lo‐ cal pairwise alignments hits were then filtered using the following criteria: a match to a methyltransferase enzyme was kept if its E-value was less than 1e-140, the sequence identity of the aligned region was greater than 80% and the subject sequence was at least twice lon‐ ger than the query. By this algorithm we found 272 candidate hits. A match to a restriction enzyme was kept if its E-value was less than 1e-140, the sequence identity within the aligned region was greater than 80% and the length of the subject sequence was at least 1.5 times longer than the query. By this algorithm we found 28 candidate matches. The candidate matches then were manually analysed.

#### **2. Research course**

#### *Methyltransferase fusions with a restriction endonuclease*

Newly found potential bifunctional restriction and modification enzymes are presented in Table 1. With our BLAST search we succeeded to filter from NCBI database 22 fusions of a DNA methyltransferase with a restriction endonuclease, carrying both endonuclease and methyltransferase domains in one polypeptide. As can be seen from Table 1, the enzymes were grouped according to their domain organisation as it was presented in Conserved Do‐ main Database [9]. As could be judged from their domain organisation, 20 new bifunctional REases are thought to represent the fusion of a REase with MTase and target recognition subunits of the type I restriction-modification systems (R-M-S structure), having similar or‐ ganisation with the known type IIC bifunctional enzymes such as AloI [10], CjeI [11], MmeI [12], PpiI [13], TstI [13] and TspGWI [14]. Type I RMS enzymes are multisubunit proteins that function as a single protein complex, consisting of R, M and S subunits [3]. The S subu‐ nit is the specificity subunit that determines which DNA sequence is recognised. The R sub‐ unit is essential for cleavage (restriction) and the M subunit catalyses the methylation reaction. Their protein products are marked as HsdS, HsdR and HsdM, respectively. Cova‐ lent linking of these subunits in one polypeptide is not thought to interfere with their cata‐ lytic activities, giving an opportunity for successful fusion. Currently, the REBASE contains more than 8000 entries corresponding to Type I RMS. Hypothetically, any of these RMS could be joint, naturally or artificially, giving a new bifunctional RMS.

an important role in evolution of metyltransferases and in origin of multidomain eukaryotic

For the current work we searched protein databases as described above and found 76 new potential bifunctional MTases. Their structural organisation is presented and discussed in the report. Beside this, we analysed structural organisation of 627 non-putative prokaryotic DNA-methyltransferases available out of 980 methyltransferases collected up to now in RE‐ BASE. The most frequently observed structural type, other than "canonical" MTases, repre‐ sents SsoII-related methyltransferases, capable to serve as transcriptional autoregulatory proteins. These data provide additional evidences that gene fusion might play an important role in evolution of methyltransferases, restriction-modification systems and other DNAmodifying proteins. We discuss the general consequences of a hypothetical protein fusion

For this study 10619 methyltransferases and 3250 restriction enzyme sequences were down‐ loaded from the REBASE database [2] and searched against the non-redundant protein data‐ base which was downloaded from the NCBI's ftp site. The similarity search was carried out by BLAST version 2.2.23+ [8] on a Linux server using the BLAST default parameters. The lo‐ cal pairwise alignments hits were then filtered using the following criteria: a match to a methyltransferase enzyme was kept if its E-value was less than 1e-140, the sequence identity of the aligned region was greater than 80% and the subject sequence was at least twice lon‐ ger than the query. By this algorithm we found 272 candidate hits. A match to a restriction enzyme was kept if its E-value was less than 1e-140, the sequence identity within the aligned region was greater than 80% and the length of the subject sequence was at least 1.5 times longer than the query. By this algorithm we found 28 candidate matches. The candidate

Newly found potential bifunctional restriction and modification enzymes are presented in Table 1. With our BLAST search we succeeded to filter from NCBI database 22 fusions of a DNA methyltransferase with a restriction endonuclease, carrying both endonuclease and methyltransferase domains in one polypeptide. As can be seen from Table 1, the enzymes were grouped according to their domain organisation as it was presented in Conserved Do‐ main Database [9]. As could be judged from their domain organisation, 20 new bifunctional REases are thought to represent the fusion of a REase with MTase and target recognition subunits of the type I restriction-modification systems (R-M-S structure), having similar or‐ ganisation with the known type IIC bifunctional enzymes such as AloI [10], CjeI [11], MmeI [12], PpiI [13], TstI [13] and TspGWI [14]. Type I RMS enzymes are multisubunit proteins that function as a single protein complex, consisting of R, M and S subunits [3]. The S subu‐

methyltransferases.

**1.1. Method used**

event with methyltransferases and RMS enzymes.

72 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

matches then were manually analysed.

*Methyltransferase fusions with a restriction endonuclease*

**2. Research course**

One RMS from *Bacteroides sp*. D22, probably originated from the fusion of type III enzymes. It has conserved motifs similar to Eco57I protein, which consists of Mod and Res subunits of type III enzymes [15]. Type III systems are composed of two genes (mod and res) encoding protein subunits that function in one protein complex either in DNA recognition and modi‐ fication, Mod, or restriction, Res [3]. As in the case of the type I enzymes, in-frame fusion might not affect their normal functioning, thus, being also favourable for new bifunctional protein origin.

One of the found RMS from *Arthrospira maxima* CS-328 has a different domain organisation, belonging to R-M type of structure (type II REase and MTase fusion). It could be suggested that it originated from fusion of head-to-tail oriented Type II REase and MTase genes. The principal possibility of a new bifunctional RMS origin by this mechanism was proven by inframe joining of type IIEco29kI REase and MTase genes [5]. The resulting RMS was capable of defending host cells from phage invasion, although 100 times less effectively than the wild type. In a similar way, a new bifunctional RMS could appear from other head-to-tail oriented RMS of type II such as AccI, BanI, Bsp6I, BsuBI, Cfr9I, DdeI, EagI, EcoPI, EcoP15, EcoRI, FnuDI, HaeIII, HgiBI, HgiCI, HgiCII, HgiDI, HgiEI, HgiGI, HhaII, HincII, HindIII, HinfI, HpaI, MboII, MwoI, NcoI, NdeI, NgoMI, NgoPII, NlaIII, PaeR7I, RsrI, SalI, Sau3A, Sau96I, TaqI, TthHB8I, XbaI, and XmaI [16]. Type II restriction enzymes and modification enzymes work separately and their fusion could create steric difficulties for their functional‐ ity. In fact, in the case of the RM.Eco29kI enzyme, its REase activity decreased three times in comparison with initial R.Eco29kI nuclease. This is perhaps why it is the only found exam‐ ple of natural bifunctional RMS originating from type II RMS.


NP\_860965, 1164 aa, type I restriction/modification enzyme, *Helicobacter hepaticus* ATCC 51449.

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ZP\_04582604, 894 aa, type II restriction-modification enzyme, *Helicobacter winghamensis* ATCC BAA-430.

ZP\_01253792, 1020 aa, type II restriction-modification enzyme, *Psychroflexus torquis* ATCC 700755.

ZP\_07401090, 737 aa, type II restriction-modification enzyme, *Campylobacter coli* JV20.

YP\_003127864, 1068 aa, N-6 DNA Methylase, *Methanocaldococcus fervens* AG86.

YP\_001783948, 1110 aa, N-6 DNA Methylase, *Haemophilus somnus* 2336.

ZP\_06253288, 1297 aa, putative type I restriction modification DNA specificity domain protein, *Prevotella copri* DSM

ZP\_06373928, 1080 aa, hypothetical protein, *Campylobacter jejuni subsp. jejuni* 1336.

74 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

YP\_002344446, 1339 aa, restriction modification enzyme, *Campylobacter jejuni subsp. Jejuni* NCTC 11168.

ZP\_06409683, 889 aa, putative type II restriction-modification enzyme, *Clostridium hathewayi* DSM 13479.

ZP\_06998902.1, 1053 aa, type IIS restriction endonuclease, *Bacteroides sp*. D22.

ZP\_03658113, 1171 aa, type II restriction-modification enzyme, *Helicobacter cinaedi* CCUG 18818.

ZP\_03656081, 1322 aa, restriction modification enzyme, *Helicobacter canadensis* MIT 98-5491.

YP\_003633850, 1134 aa, restriction modification system DNA specificity domain protein, *Brachyspira murdochii* DSM

18205.

12563.

the case of dam, and cytosine in the case of dcm. It could be suggested that originally they belonged to two different genes, and were joint in-frame occasionally. The post-segregation killing effect of restriction-modification enzymes prevents RMS from being lost [22], thus promoting maintenance of a fused ORF and its spreading in bacterial populations. The next chapter will be devoted to a more detailed analysis of this effect. The other 23 bifunctional MTases of M-M type probably originated from a similar joining of two dam methyltransfer‐ ase genes (Table 2). Further evolution of bifunctional MTases depends on their involvement in RMS functioning. If both activities are critical for the RMS work, for example, modifying two different bases of an asymmetric recognition sequence [23], they will be maintained as elements of this RMS. If the activity of at least one MTase domain of the bifunctional en‐ zyme is redundant, it could accumulate mutations and, after many generations, reduce or

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CBL28228.1, 825 aa, Type I restriction-modification system methyltransferase subunit, *Synergistetes bacterium* SGP1.

gain new substrate specificity and function in the cell history.

ZP\_04875770, 760 aa, N-6 DNA Methylase family, *Aciduliprofundum boonei* T469.

YP\_002004068, 785 aa, N-6 DNA methylase, *Candidatus phytoplasma mali*.

YP\_006537.1, 754 aa, DNA methyltransferase Dmt, *Enterobacteria* phage P1. ZP\_07502038, 862 aa, putative DNA methyltransferase, *Escherichia coli* M605.

YP\_002961053, 777 aa, hypothetical protein MCJ\_005510, *Mycoplasma conjunctivae* HRC/581.

ZP\_04643292, 907 aa, putative restriction-modification enzyme, *Lactobacillus gasseri* 202-4.

**MS**

**MM**

**Table 1.** Methyltransferase fusions with a restriction endonuclease. Domains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings and their classification is shown under the table. Short description for each found protein includes Gene Bank accession number, length in amino acids, current name in the database and host strain information.

The newly found potential type IIC proteins are good candidates to expand on the current list of 12 bifunctional enzymes: AloI[10], BcgI [17], BseMII [18], BseRI [19], BspLU11III [20], CjeI[11], Eco57I [15], HaeIV [21], MmeI [12], PpiI [13], TstI [13] and TspGWI [14]. Taking into consideration intensiveness with what new microbial genomes have been sequencing dur‐ ing the last decade, new bifunctional RMS could be discovered very soon.

#### **3. Fusions between two DNA methyltransferases**

As shown in Table 2, in contrast to the situation with bifunctional MTase - REase fusions, among 54 newly found fusions between two methyltransferases 49, apparently, are joining of two different methyltransferase ORFs (M-M structure) and 5 of methyltransferase (HsdM) and target recognition subunit (HsdS, M-S structure). 11 M-M type proteins represent inter‐ esting examples of dcm and dam methyltransferase fusion. In this case dam corresponds not only to one particular MTase, but to a conserved domain common for DNA adenine methyl‐ transferases, as adopted from the Conserved Domain Database web site [9]. In a similar way, dcm corresponds to a conserved domain common for DNA cytosine methyltransferas‐ es. These MTases catalyse methyl group transfers to different nucleotide bases, adenine in the case of dam, and cytosine in the case of dcm. It could be suggested that originally they belonged to two different genes, and were joint in-frame occasionally. The post-segregation killing effect of restriction-modification enzymes prevents RMS from being lost [22], thus promoting maintenance of a fused ORF and its spreading in bacterial populations. The next chapter will be devoted to a more detailed analysis of this effect. The other 23 bifunctional MTases of M-M type probably originated from a similar joining of two dam methyltransfer‐ ase genes (Table 2). Further evolution of bifunctional MTases depends on their involvement in RMS functioning. If both activities are critical for the RMS work, for example, modifying two different bases of an asymmetric recognition sequence [23], they will be maintained as elements of this RMS. If the activity of at least one MTase domain of the bifunctional en‐ zyme is redundant, it could accumulate mutations and, after many generations, reduce or gain new substrate specificity and function in the cell history.

ZP\_06009534, 727 aa, restriction and modification enzyme, *Campylobacter fetus subsp. venerealis* str. Azul-94.

**Table 1.** Methyltransferase fusions with a restriction endonuclease. Domains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings and their classification is shown under the table. Short description for each found protein includes Gene Bank accession number, length in amino

ing the last decade, new bifunctional RMS could be discovered very soon.

**3. Fusions between two DNA methyltransferases**

The newly found potential type IIC proteins are good candidates to expand on the current list of 12 bifunctional enzymes: AloI[10], BcgI [17], BseMII [18], BseRI [19], BspLU11III [20], CjeI[11], Eco57I [15], HaeIV [21], MmeI [12], PpiI [13], TstI [13] and TspGWI [14]. Taking into consideration intensiveness with what new microbial genomes have been sequencing dur‐

As shown in Table 2, in contrast to the situation with bifunctional MTase - REase fusions, among 54 newly found fusions between two methyltransferases 49, apparently, are joining of two different methyltransferase ORFs (M-M structure) and 5 of methyltransferase (HsdM) and target recognition subunit (HsdS, M-S structure). 11 M-M type proteins represent inter‐ esting examples of dcm and dam methyltransferase fusion. In this case dam corresponds not only to one particular MTase, but to a conserved domain common for DNA adenine methyl‐ transferases, as adopted from the Conserved Domain Database web site [9]. In a similar way, dcm corresponds to a conserved domain common for DNA cytosine methyltransferas‐ es. These MTases catalyse methyl group transfers to different nucleotide bases, adenine in

acids, current name in the database and host strain information.

76 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

ZP\_03272741.1, 1473 aa, protein of unknown function DUF559, *Arthrospira maxima* CS-328.

**RM**

ZP\_00231287, 608 aa, DNA methyltransferase, *Listeria monocytogenes* str. 4b H7858.

YP\_003532979, 617 aa, adenine methyltransferase, *Erwinia amylovora* CFBP1430.

ZP\_04573763, 670 aa, adenine methyltransferase, *Fusobacterium* sp. 7\_1.

ACZ68466, 710 aa, modification Methylase, *Staphylococcus aureus*. ADD81212, 700 aa, DNA methyltransferase, *Enterococcus faecalis*.

*gonorrhoeae* PID332.

*influenzae* PittEE.

*gonorrhoeae* PID332.

ZP\_07642163, 716 aa, modification Methylase, *Streptococcus mitis* SK597.

ZP\_06290558, 729 aa, modification methylase, *Peptoniphilus lacrimalis* 315-B.

ZP\_05861343, 722 aa, putative adenine methylase, *Jonquetella anthropi* E3\_33 E1. ZP\_04977178, 722 aa, adenine methyltransferase, *Mannheimia haemolytica* PHL213.

ZP\_01834649, 716 aa, adenine methyltransferase, *Streptococcus pneumonia* SP23-BS72.

YP\_002650601, 617 aa, adenine specific DNA methyltransferase, *Erwinia pyrifoliae* Ep1/96.

ZP\_06560574, 722 aa, DNA adenine methylase, *Megasphaera genomosp*. type\_1 str. 28L.

ZP\_03958097, 625 aa, possible site-specific DNA methyltransferase, *Lactobacillus ruminis* ATCC 25644.

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ZP\_06871587, 670 aa, adenine methyltransferase, *Fusobacterium nucleatum subsp. nucleatum* ATCC 23726.

ZP\_04452030, 711 aa, hypothetical protein GCWU000182\_01325, *Abiotrophia defectiva* ATCC 49176.

NP\_602723, 496 aa, adenine-specific methyltransferase, *Fusobacterium nucleatum subsp. nucleatum* ATCC 25586.

ZP\_05849684.1, 752 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* NT127.

ZP\_01788156.1, 687 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* 3655. YP\_001292975.1, 724 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* PittGG.

ZP\_01789924.1, 681 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* PittAA.

AAT40808, 518 aa, type III restriction-modification system methyltransferase-like protein, *Haemophilus influenzae*. ZP\_04736658.1, 724 aa, putative type III restriction/modification system modification Methylase, *Neisseria*

YP\_001291080.1, 713 aa, putative type III restriction/modification system modification methylase, *Haemophilus*

ZP\_04736658.1, 724 aa, putative type III restriction/modification system modification methylase, *Neisseria*

ZP\_00231287, 608 aa, DNA methyltransferase, *Listeria monocytogenes* str. 4b H7858.

ZP\_03958097, 625 aa, possible site-specific DNA methyltransferase, *Lactobacillus ruminis* ATCC 25644.

YP\_003532979, 617 aa, adenine methyltransferase, *Erwinia amylovora* CFBP1430.

YP\_002650601, 617 aa, adenine specific DNA methyltransferase, *Erwinia pyrifoliae* Ep1/96.

ZP\_06871587, 670 aa, adenine methyltransferase, *Fusobacterium nucleatum subsp. nucleatum* ATCC 23726.

ZP\_04573763, 670 aa, adenine methyltransferase, *Fusobacterium* sp. 7\_1.

ZP\_07381018, 808 aa, DNA adenine methylase, *Pantoea sp*. AB.

78 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

ZP\_06715601, 722 aa, DNA C5 cytosine methyltransferase, *Edwardsiella tarda* ATCC 23685.

YP\_003237972, 1013 aa putative DNA methyltransferase, *Escherichia coli* O111:H- str. 11128.

AF503408\_62, 672 aa, adenine DNA methyltransferase Dam, *Enterobacteria* phage P7.

AF458984\_2, 952 aa, N6 adenine and C5 cytosine DNA methyltransferase, *Acinetobacter lwoffii*.

AAS09913, 1068 aa, BsmBI M1-M2 methyltransferase fusion protein, *Geobacillus stearothermophilus.*

AAQ72364, 1007 aa, methylase fusion protein, *Geobacillus stearothermophilus*.

ACZ62643, 687 aa, BtsCIM Methylase, *Ureibacillus thermosphaericus*. ZP\_05132801, 653 aa, DNA methyltransferase, *Clostridium sp*. 7\_2\_43FAA.

AF458983\_2, 1061 aa, N6 adenine and C5 cytosine DNA methyltransferase, *Hafnia alvei*.

ZP\_02080986, 664 aa, hypothetical protein CLOLEP\_02452, *Clostridium leptum* DSM 753.

YP\_002744130, 627 aa, DNA adenine Methylase, *Streptococcus equi subsp. zooepidemicus*. YP\_002746827, 620 aa, DNA adenine Methylase, *Streptococcus equi subsp. equi* 4047. ZP\_04008314, 618 aa, adenine DNA methyltransferase, *Lactobacillus salivarius* ATCC 11741.

P29347, 653 aa, Adenine-specific methyltransferase StsI, *Streptococcus sanguinis*.

ZP\_02701644, 819 aa, DNA methyltransferase Dmt, *Salmonella enterica subsp. enterica* serovar Newport str. SL317.

ZP\_06560574, 722 aa, DNA adenine methylase, *Megasphaera genomosp*. type\_1 str. 28L.

ZP\_06290558, 729 aa, modification methylase, *Peptoniphilus lacrimalis* 315-B.

ZP\_04452030, 711 aa, hypothetical protein GCWU000182\_01325, *Abiotrophia defectiva* ATCC 49176.

ZP\_05861343, 722 aa, putative adenine methylase, *Jonquetella anthropi* E3\_33 E1.

ZP\_04977178, 722 aa, adenine methyltransferase, *Mannheimia haemolytica* PHL213.

ACZ68466, 710 aa, modification Methylase, *Staphylococcus aureus*.

ADD81212, 700 aa, DNA methyltransferase, *Enterococcus faecalis*.

ZP\_07642163, 716 aa, modification Methylase, *Streptococcus mitis* SK597.

ZP\_01834649, 716 aa, adenine methyltransferase, *Streptococcus pneumonia* SP23-BS72.

NP\_602723, 496 aa, adenine-specific methyltransferase, *Fusobacterium nucleatum subsp. nucleatum* ATCC 25586.

ZP\_05849684.1, 752 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* NT127.

AAT40808, 518 aa, type III restriction-modification system methyltransferase-like protein, *Haemophilus influenzae*.

ZP\_04736658.1, 724 aa, putative type III restriction/modification system modification Methylase, *Neisseria gonorrhoeae* PID332.

ZP\_01788156.1, 687 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* 3655.

YP\_001292975.1, 724 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* PittGG.

YP\_001291080.1, 713 aa, putative type III restriction/modification system modification methylase, *Haemophilus influenzae* PittEE.

ZP\_01789924.1, 681 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* PittAA.

ZP\_04736658.1, 724 aa, putative type III restriction/modification system modification methylase, *Neisseria gonorrhoeae* PID332.

ZP\_04719045.1, 736 aa, putative type III restriction/modification system modification methylase, *Neisseria gonorrhoeae* 35/02.

methylases. The pfam01555 conserved domain could be found both in type II MTases, such as M.KpnI, and type III Mod proteins, such as EcoP1I and EcoP15I. In contrast, a 60 aa con‐ served pfam12564 domain could be found only in several type III enzymes. Its addition could influence the biochemical properties of the corresponding proteins. To establish the character of its influence experimentally in the future, it would be necessary to compare the biochemical properties of MTases containing the pfam12564 60 aa sequence, with MTases

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**Figure 1.** Domain organisation of DNMT1, a DNA methyltransferase 1 from *H. sapiens* (a); M.AimAII, a methyltransfer‐ ase from *Ascobolus immersus* (b), M.NcrNI from *Neurospora crassa* (c) and M.AimAI from *Ascobolus immersus* (d)*.* Do‐ mains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different

Figure 2 shows the consequences of a RMS protein fusion with a hypothetical protein, X. The upper part of the figure represents normal RMS functioning, when host DNA is protect‐ ed by an MTase and foreign DNA is degraded by a cognate REase. In the case of RMS loss, MTase and REase enzymes will be diluted following cell divisions, host DNA will become

This effect is known as post-segregation killing [22] and is possibly due to REase activity lasting longer than MTase activity after the RMS loss. The lower part of the figure illustrates

**4. Fusion of RMS enzymes with a hypothetical protein**

unprotected and, finally, degraded by the residual REase activity.

not containing it.

fillings.

YP\_975326.1, 744 aa, putative type III restriction/modification system modification methylase, *Neisseria meningitidis* FAM18.

ZP\_05849684.1, 752 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* NT127.

ZP\_05986443.1, 701 aa, type III restriction/modification enzyme, methylase subunit, *Neisseria lactamica* ATCC 23970.

ABI85522.1, 717 aa, DNA methylase M.Hin1056ModP-2, *Haemophilus influenzae*.

YP\_207780.1, 756 aa, putative type III restriction/modification system modification methylase, *Neisseria gonorrhoeae* FA 1090.

ZP\_05986443.1, 701 aa, type III restriction/modification enzyme, *Neisseria lactamica* ATCC 23970.

**Table 2.** Fusions between two DNA methyltransferases. Domains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings and their classification is shown under the table. Short description for each found protein includes Gene Bank accession number, length in amino acids, current name in the database and host strain information.

For example, DNMT1, a DNA methyltransferase 1 from *H. sapiens* [24], contains a conserved domain of m5C MTases at its C-terminal domain. The N-terminal and central parts of the enzyme include several different domains such as DMAP binding domain, replication foci domain, zinc finger domain and two bromo adjacent homology domains (Figure 1 a). It could be suggested that several gene fusion events were involved in its evolutionary. This hypothesis is supported by existence of simpler homologs of DNMT1 such as, for example, M.AimAII (Figure 1 b) and M.AimAI (Figure 1 d) from *Ascobolus immersus*, and M.NcrNI from *Neurospora crassa* (Figure 1 c), looking like not completely assembled DNMT1 with one or several domains missing. 15 other M-M type bifunctional MTases from Table 2 represent the fusion of a conserved pfam12564 type III RMS 60 aa domain with pfam01555 N4-N6 MTase domain, characteristic both for N4 cytosine-specific and N6 adenine-specific DNA methylases. The pfam01555 conserved domain could be found both in type II MTases, such as M.KpnI, and type III Mod proteins, such as EcoP1I and EcoP15I. In contrast, a 60 aa con‐ served pfam12564 domain could be found only in several type III enzymes. Its addition could influence the biochemical properties of the corresponding proteins. To establish the character of its influence experimentally in the future, it would be necessary to compare the biochemical properties of MTases containing the pfam12564 60 aa sequence, with MTases not containing it.

ZP\_04719045.1, 736 aa, putative type III restriction/modification system modification methylase, *Neisseria*

YP\_975326.1, 744 aa, putative type III restriction/modification system modification methylase, *Neisseria meningitidis*

ZP\_05986443.1, 701 aa, type III restriction/modification enzyme, methylase subunit, *Neisseria lactamica* ATCC 23970.

YP\_207780.1, 756 aa, putative type III restriction/modification system modification methylase, *Neisseria gonorrhoeae*

**Table 2.** Fusions between two DNA methyltransferases. Domains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings and their classification is shown under the table. Short description for each found protein includes Gene Bank accession number, length in amino acids, current

For example, DNMT1, a DNA methyltransferase 1 from *H. sapiens* [24], contains a conserved domain of m5C MTases at its C-terminal domain. The N-terminal and central parts of the enzyme include several different domains such as DMAP binding domain, replication foci domain, zinc finger domain and two bromo adjacent homology domains (Figure 1 a). It could be suggested that several gene fusion events were involved in its evolutionary. This hypothesis is supported by existence of simpler homologs of DNMT1 such as, for example, M.AimAII (Figure 1 b) and M.AimAI (Figure 1 d) from *Ascobolus immersus*, and M.NcrNI from *Neurospora crassa* (Figure 1 c), looking like not completely assembled DNMT1 with one or several domains missing. 15 other M-M type bifunctional MTases from Table 2 represent the fusion of a conserved pfam12564 type III RMS 60 aa domain with pfam01555 N4-N6 MTase domain, characteristic both for N4 cytosine-specific and N6 adenine-specific DNA

name in the database and host strain information.

ZP\_05849684.1, 752 aa, twin-argninine leader-binding protein DmsD, *Haemophilus influenzae* NT127.

ZP\_05986443.1, 701 aa, type III restriction/modification enzyme, *Neisseria lactamica* ATCC 23970.

ABI85522.1, 717 aa, DNA methylase M.Hin1056ModP-2, *Haemophilus influenzae*.

80 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

*gonorrhoeae* 35/02.

FAM18.

FA 1090.

**Figure 1.** Domain organisation of DNMT1, a DNA methyltransferase 1 from *H. sapiens* (a); M.AimAII, a methyltransfer‐ ase from *Ascobolus immersus* (b), M.NcrNI from *Neurospora crassa* (c) and M.AimAI from *Ascobolus immersus* (d)*.* Do‐ mains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings.

#### **4. Fusion of RMS enzymes with a hypothetical protein**

Figure 2 shows the consequences of a RMS protein fusion with a hypothetical protein, X. The upper part of the figure represents normal RMS functioning, when host DNA is protect‐ ed by an MTase and foreign DNA is degraded by a cognate REase. In the case of RMS loss, MTase and REase enzymes will be diluted following cell divisions, host DNA will become unprotected and, finally, degraded by the residual REase activity.

This effect is known as post-segregation killing [22] and is possibly due to REase activity lasting longer than MTase activity after the RMS loss. The lower part of the figure illustrates the situation of the fusion of some protein X with one of the RMS enzymes. If this joining will not affect seriously the enzyme activities, the RMS will continue to protect cells from foreign DNA invasion. In the case of RMS loss, the same mechanism will lead to host DNA degradation. In this way, a fused protein with RMS enzymes will be supported and spread among bacterial populations. In fact, during a BLAST search we could see close homologues (>98%) of bifunctional enzymes such as, for example, CjeI, spread among numerous strains of the *Campylobacter* group of microorganisms. These observations could illustrate our hy‐ pothesis. Another example of the propagation of a RMS-fused protein is provided by M.SsoII-related enzymes. These MTases represent fusion of a regulatory protein with a C5 cytosine methyltransferase [4] and can be found in 63 different microbial taxa.

In another case, if fusion with a hypothetical protein is detrimental for the activity of one of the RMS enzymes, an outcome will depend on which of the enzymes is affected. If an MTase activity will be reduced, the corresponding RMS will be eliminated due to host DNA degra‐ dation by cognate REase. If a REase activity is disturbed, the corresponding RMS will be‐ come non-functional, will not be supported by post-segregation killing mechanism and, after many generations, could disappear or take on different functions in the cell. Another, less probable, scenario is possible if a joining with a hypothetical protein would improve the properties of RMS enzymes. In this situation, the corresponding RMS would protect host cells more effectively and that would increase their selective advantage over competitive mi‐ crobial populations, which, in turn, could lead to a wider distribution of the RMS carrying

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83

**5. Domain organisation of non-putative DNA methyltransferases from**

We analyse structural organisation of 627 non-putative prokaryotic DNA-methyltransferas‐ es collected up to now in REBASE, a major database of restriction-modification enzymes [2]. We succeeded to download sequences of 627 prokaryotic methyltransferases out of 980 nonputative MTases enlisted in REBASE on 01.12.2011 (for their detailed description see Supple‐ mentary materials). Out of 627, 190 sequences belong to dcm type of DNAmethyltransferases; 172 - to N6-N4 type; 99 - to HsdM type and 78 - to dam-related enzymes according to Conserved Domain Database [9]. We found that the most frequently observed structure, other than "canonical" methyltransferases with conserved motives responsible for binding with AdoMet and a methyl group transfer, represents C5-methyltransferase core domain fusion with a regulatory DNA-binding protein and up to now includes 18 potential enzymes (Table 3). These SsoII-related methyltransferases carry additional DNA-binding HTH-domains and they are capable to serve as transcriptional autorepressors [4]. Among these 18 SsoII-like MTases HTH-motif can be located in majority of cases on N-terminal part, and in three proteins - in the middle of their polypeptide chains (M.Esp1396I, M.PflMI and M.SfiI; Table 3). Ability for autoregulation was not confirmed for majority of these SsoII-like MTases and will require some experimental proofs, which could be considered as perspec‐ tive future directions of research. The fact of SsoII-like enzymes propagation among differ‐ ent bacterial taxa could illustrate well our analysis of a RMS protein fusion with a

hypothetical protein, described in the previous chapter.

M.AseI, Type II methyltransferase, recognition sequence ATTAAT, GenBank ADO24185, 552 aa

the fused protein.

**REBASE**

**Figure 2.** Schematic representation of post segregation killing effect responsible for maintaining RMS and proteins joint with one of its enzymes. The figure summarizes different outcomes of a hypothetical protein X joining to one of a RMS enzymes. If this joining is neutral or positive for the RMS functioning, it will be maintained and spread; if detri‐ mental, it will be eliminated. Black filled circles show methylated nucleotides, interrupted lines - degraded DNA.

In another case, if fusion with a hypothetical protein is detrimental for the activity of one of the RMS enzymes, an outcome will depend on which of the enzymes is affected. If an MTase activity will be reduced, the corresponding RMS will be eliminated due to host DNA degra‐ dation by cognate REase. If a REase activity is disturbed, the corresponding RMS will be‐ come non-functional, will not be supported by post-segregation killing mechanism and, after many generations, could disappear or take on different functions in the cell. Another, less probable, scenario is possible if a joining with a hypothetical protein would improve the properties of RMS enzymes. In this situation, the corresponding RMS would protect host cells more effectively and that would increase their selective advantage over competitive mi‐ crobial populations, which, in turn, could lead to a wider distribution of the RMS carrying the fused protein.

the situation of the fusion of some protein X with one of the RMS enzymes. If this joining will not affect seriously the enzyme activities, the RMS will continue to protect cells from foreign DNA invasion. In the case of RMS loss, the same mechanism will lead to host DNA degradation. In this way, a fused protein with RMS enzymes will be supported and spread among bacterial populations. In fact, during a BLAST search we could see close homologues (>98%) of bifunctional enzymes such as, for example, CjeI, spread among numerous strains of the *Campylobacter* group of microorganisms. These observations could illustrate our hy‐ pothesis. Another example of the propagation of a RMS-fused protein is provided by M.SsoII-related enzymes. These MTases represent fusion of a regulatory protein with a C5-

**Figure 2.** Schematic representation of post segregation killing effect responsible for maintaining RMS and proteins joint with one of its enzymes. The figure summarizes different outcomes of a hypothetical protein X joining to one of a RMS enzymes. If this joining is neutral or positive for the RMS functioning, it will be maintained and spread; if detri‐ mental, it will be eliminated. Black filled circles show methylated nucleotides, interrupted lines - degraded DNA.

cytosine methyltransferase [4] and can be found in 63 different microbial taxa.

82 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

#### **5. Domain organisation of non-putative DNA methyltransferases from REBASE**

We analyse structural organisation of 627 non-putative prokaryotic DNA-methyltransferas‐ es collected up to now in REBASE, a major database of restriction-modification enzymes [2]. We succeeded to download sequences of 627 prokaryotic methyltransferases out of 980 nonputative MTases enlisted in REBASE on 01.12.2011 (for their detailed description see Supple‐ mentary materials). Out of 627, 190 sequences belong to dcm type of DNAmethyltransferases; 172 - to N6-N4 type; 99 - to HsdM type and 78 - to dam-related enzymes according to Conserved Domain Database [9]. We found that the most frequently observed structure, other than "canonical" methyltransferases with conserved motives responsible for binding with AdoMet and a methyl group transfer, represents C5-methyltransferase core domain fusion with a regulatory DNA-binding protein and up to now includes 18 potential enzymes (Table 3). These SsoII-related methyltransferases carry additional DNA-binding HTH-domains and they are capable to serve as transcriptional autorepressors [4]. Among these 18 SsoII-like MTases HTH-motif can be located in majority of cases on N-terminal part, and in three proteins - in the middle of their polypeptide chains (M.Esp1396I, M.PflMI and M.SfiI; Table 3). Ability for autoregulation was not confirmed for majority of these SsoII-like MTases and will require some experimental proofs, which could be considered as perspec‐ tive future directions of research. The fact of SsoII-like enzymes propagation among differ‐ ent bacterial taxa could illustrate well our analysis of a RMS protein fusion with a hypothetical protein, described in the previous chapter.

**6. Conclusion**

*Abbreviations*

**Author details**

ces, Russia

**References**

Marina L. Mokrishcheva1

2 ICGEB, Area Science Park, Italy

*Research*, 31, 1805-1812.

*Acids Research*, 25, 2114-2120.

in restriction-modification enzyme evolution.

Dmitry V. Nikitin1,2\*, Attila Kertesz-Farkas2

\*Address all correspondence to: dvnikitin@rambler.ru

tions. *Molecular Biotechnology*, 23, 225-243.

Here we report finding 76 new bifunctional methyltransferases. The majority of the found joint proteins with a nuclease are thought to be fusions of a restriction nuclease with methyl‐ ase and target recognition subunits of type I restriction-modification systems (R-M-S struc‐ ture). The majority of the found joint proteins between two methylases appears to be damdcm and dam-dam enzyme fusions (M-M structure). Similar proteins could serve as structural intermediates for multidomain eukaryotic methyltransferase evolution. We sug‐ gest that a hypothetical protein fusion with a restriction-modification enzyme can promote its propagation in bacterial populations. Altogether, our data illustrate a role of gene fusion

ORF - open reading frame; REase - restriction endonuclease; MTase - methyltransferase; RMS - restriction-modification system; TRD - target recognition domain; aa - amino acids.

1 Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Scien‐

[1] Williams, R. (2003). Restriction endonucleases: classification, properties, and applica‐

[2] Vincze, T., Posfai, J., & Macelis, D. (2010). REBASE-a database for DNA restriction and modification: enzymes, genes and genomes. *Nucleic Acids Research*, 38, 234-236. [3] Roberts, R. J., Belfort, M., Bestor, T., et al. (2003). A nomenclature for restriction en‐ zymes, DNA methyltransferases, homing endonucleases and their genes. *Nucleic Acids*

[4] Karyagina, A., Shilov, I., Tashlitskii, V., Khodoun, M., Vasil'ev, S., Lau, P., & Nikol‐ skaya, I. (1997). Specific binding of SsoII DNA methyltransferase to its promoter re‐ gion provides the regulation of SsoII restriction-modification gene expression. *Nucleic*

, Alexander S. Solonin1

and

Bifunctional Prokaryotic DNA-Methyltransferases

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85

**Table 3.** Domain organisation of SsoII-like prokaryotic DNA methyltransferases, collected in REBASE. Domains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings.

#### **6. Conclusion**

M.AvaII, Type II methyltransferase, recognition sequence GGWCC, GenBank BA000019, 477 aa M.BadAII, Type II methyltransferase, recognition sequence CCNGG, GenBank AP009256, 383 aa M.Csp68KI, Type II methyltransferase, recognition sequence GGWCC, GenBank CP000806, 461 aa M.Ecl18kI, Type II methyltransferase, recognition sequence CCNGG, GenBank Y16897, 379 aa M.Eco47II, Type II methyltransferase, recognition sequence GGNCC, GenBank X82105, 417 aa M.Kpn2kI, Type II methyltransferase, recognition sequence CCNGG, GenBank AF300473, 379 aa M.MspI, Type II methyltransferase, recognition sequence CCGG, GenBank X14191, 418 aa M.Sau96I, Type II methyltransferase, recognition sequence GGNCC, GenBank X53096, 430 aa M1.ScrFI, Type II methyltransferase, recognition sequence CCNGG, GenBank U89998, 389 aa M.SenPI, Type II methyltransferase, recognition sequence CCNGG, GenBank U65460, 379 aa M.SinI, Type II methyltransferase, recognition sequence GGWCC, GenBank J03391, 461 aa M.SsoII, Type II methyltransferase, recognition sequence CCNGG, GenBank M86545, 379 aa

84 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

M.StyD4I, Type II methyltransferase, recognition CCNGG, GenBank D73442, 379 aa

M.Esp1396I, Type II methyltransferase, recognition sequence CCANNNNNTGG, GenBank AF527822, 332 aa M.PflMI, orphan methyltransferase, recognition sequence CCANNNNNTGG, GenBank ADZ31427, 363 aa M.SfiI, Type II methyltransferase, recognition sequence GGCCNNNNNGGCC, GenBank AF039750, 421 aa

M1.SfaNI, Type II methyltransferase, recognition sequence GCATC, GenBank GU565605, 700 aa

**Table 3.** Domain organisation of SsoII-like prokaryotic DNA methyltransferases, collected in REBASE. Domains are predicted and presented as in Conserved Domain Database [9]. Different domains are shown by different fillings.

Here we report finding 76 new bifunctional methyltransferases. The majority of the found joint proteins with a nuclease are thought to be fusions of a restriction nuclease with methyl‐ ase and target recognition subunits of type I restriction-modification systems (R-M-S struc‐ ture). The majority of the found joint proteins between two methylases appears to be damdcm and dam-dam enzyme fusions (M-M structure). Similar proteins could serve as structural intermediates for multidomain eukaryotic methyltransferase evolution. We sug‐ gest that a hypothetical protein fusion with a restriction-modification enzyme can promote its propagation in bacterial populations. Altogether, our data illustrate a role of gene fusion in restriction-modification enzyme evolution.

#### *Abbreviations*

ORF - open reading frame; REase - restriction endonuclease; MTase - methyltransferase; RMS - restriction-modification system; TRD - target recognition domain; aa - amino acids.

#### **Author details**

Dmitry V. Nikitin1,2\*, Attila Kertesz-Farkas2 , Alexander S. Solonin1 and Marina L. Mokrishcheva1

\*Address all correspondence to: dvnikitin@rambler.ru

1 Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Scien‐ ces, Russia

2 ICGEB, Area Science Park, Italy

#### **References**


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**Section 3**

**Protein Arginine Methylation in Mammals**

**Protein Arginine Methylation in Mammals**

**Chapter 4**

**Deciphering Protein Arginine**

Ruben Esse, Paula Leandro, Isabel Rivera, Isabel Tavares de Almeida, Henk J Blom and

Additional information is available at the end of the chapter

The myriad of different post-translation modifications (PTMs) of proteins augments the information encoded by the human genome, from about 25,000 genes to over 1 million proteins that compose the human proteome [1, 2]. PTMs involve the chemical modification of target amino acid residues and, among others, include the addition of a methyl group to arginine residues, or arginine methylation, which is one of the most extensive protein modifications occurring in mammalian cells. During the last years, arginine methylation has attracted growing attention due to its impact on cellular function. Attesting to its biological importance, protein arginine methylation appears to be an evolutionarily ancient modification. Arginine methylation is now recognized as a widespread PTM that occurs on multiple classes of proteins with distinct cellular localizations. The arginine residue is unique among amino acids, since its guanidine group contains five potential hydrogen bond donors (Figure 1) positioned for interactions with hydrogen bond acceptors as DNA, RNA and proteins. Each addition of a methyl group removes a hydrogen donor, so the methylation of arginine residues in proteins will readily modulate their binding interactions and thus their physiological functions. Here, we provide an introduction to protein arginine methylation and discuss the current state of knowledge regarding this modification in mammals. In addition, we provide insight into how protein arginine methylation relates with the homocysteine metabolism. Lastly, we briefly discuss how protein arginine methylation may be disturbed in the

context of hyperhomocysteinemia, sharing some of our recent results.

© 2013 Esse et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

**Methylation in Mammals**

Rita Castro

**1. Introduction**

http://dx.doi.org/10.5772/51984

**Chapter 4**

### **Deciphering Protein Arginine Methylation in Mammals**

Ruben Esse, Paula Leandro, Isabel Rivera, Isabel Tavares de Almeida, Henk J Blom and Rita Castro

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51984

#### **1. Introduction**

The myriad of different post-translation modifications (PTMs) of proteins augments the information encoded by the human genome, from about 25,000 genes to over 1 million proteins that compose the human proteome [1, 2]. PTMs involve the chemical modification of target amino acid residues and, among others, include the addition of a methyl group to arginine residues, or arginine methylation, which is one of the most extensive protein modifications occurring in mammalian cells. During the last years, arginine methylation has attracted growing attention due to its impact on cellular function. Attesting to its biological importance, protein arginine methylation appears to be an evolutionarily ancient modification. Arginine methylation is now recognized as a widespread PTM that occurs on multiple classes of proteins with distinct cellular localizations. The arginine residue is unique among amino acids, since its guanidine group contains five potential hydrogen bond donors (Figure 1) positioned for interactions with hydrogen bond acceptors as DNA, RNA and proteins. Each addition of a methyl group removes a hydrogen donor, so the methylation of arginine residues in proteins will readily modulate their binding interactions and thus their physiological functions. Here, we provide an introduction to protein arginine methylation and discuss the current state of knowledge regarding this modification in mammals. In addition, we provide insight into how protein arginine methylation relates with the homocysteine metabolism. Lastly, we briefly discuss how protein arginine methylation may be disturbed in the context of hyperhomocysteinemia, sharing some of our recent results.

© 2013 Esse et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

forming ω-*N*G,*N*<sup>G</sup>-dimethylarginine (ADMA), whereas type II members add it to the other N-terminal nitrogen of the arginine residue, yielding ω-*N*G,*N'*<sup>G</sup>-dimethylarginine (SDMA). Therefore, three distinct types of methylated arginine residues (MMA, ADMA and SDMA) are present on a horde of different proteins in the cytosol, nucleus and organelles of mam‐

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**Figure 2.** Schematic representation of the human protein arginine methyltransferase (PRMTs) family. For each mem‐ ber, the length of the longest isoform is indicated on the right. All members of the family possess at least one con‐ served MTase domain with signature motifs I, post-I, II, and III and a THW loop. Additional domains are marked in yellow boxes: SH3, ZnF (zinc finger), Myr (myristoylation), F-box, TPR (tetratricopeptide) and NosD (nitrous oxidase ac‐

To date, the human PRMT family includes eleven enzymes, termed from PRMT1 to PRMT11. Six are classified as type I (PRMT1, PRMT2, PRMT3, PRMT4, PRMT6 and PRMT8), and two as type II (PRMT5 and PRMT9). The classification of PRMT7 is controversial, as described below. The remaining two proteins, PRMT10 and PRMT11, were identified by their homology with PRMT7 and PRMT9, respectively, but their PRMT activity has not been sustained by experimental evidence [18, 19]. The genes encoding the different PRMTs are all located on different chromosomes, except the ones coding for PRMTs 1 and 4, which are both located in chromosome 19. Different splice variants coding for the corresponding isoforms are recognized for all PRMTs. Moreover, different

Although the human PRMT members vary in length from 316 to 956 amino acid residues, they all contain a highly conserved catalytic core region of around 300 residues (Figure 2). However, each PRMT presents a unique N-terminal region of variable length and distinct domain motifs, as for example PRMT3 and PRMT8 that bear a zinc finger (ZnF) and a myristoylation domain, respectively. Interestingly PRMT7 and PRMT10 exhibit a second catalytic domain. Until present, only the 3D structures of PRMT1, PRMT3 and PRMT5 were solved by X-ray crystallography [20-22]. These structures revealed that, also at the structural level, the catalytic core is highly conserved. Three structural regions

malian cells, ADMA being the most prevalent [3].

cessory protein). Adapted from [17].

cellular localizations are observed among them.

**Figure 1.** Methylation of the arginine side chain in proteins by protein arginine methyltransferases (PRMTs). The argi‐ nine residue holds 5 potential hydrogen bond donors. Mammalian PRMTs use the methyl group from a molecule of Sadenosyl-L-methionine (AdoMet) to form ω-NG-monomethylarginine (MMA). Subsequently, type I PRMTs add a methyl group to the same nitrogen atom forming ω-NG,NG-dimethylarginine (ADMA), whereas type II PRMTs generate ω-NG,N'G-dimethylarginine (SDMA).

#### **2. The mammalian PRMTs: A brief overview**

Methylation of arginine is catalyzed by protein arginine methyltransferases (PRMTs). PRMTs catalyze the transfer of methyl groups from the universal methyl donor, S-adenosyl-L-methionine (AdoMet), to the guanidine nitrogen atom of the target arginine residues with‐ in proteins [3].

The first documented protein with arginine methylation activity, then termed protein meth‐ ylase I, was purified from calf thymus [4, 5]. Additional family members were subsequently identified and purified from different tissues and cell lines [6-16]. In mammals, PRMTs are grouped in two major classes: type I and type II. Both PRMT types catalyze the transfer of a single methyl group from AdoMet to an arginine residue in the target protein, producing ω-*N*G-monomethylarginine (MMA) (Figure 1). However, differences exist when adding the second methyl group: type I members add it on the same previously methylated nitrogen, forming ω-*N*G,*N*<sup>G</sup>-dimethylarginine (ADMA), whereas type II members add it to the other N-terminal nitrogen of the arginine residue, yielding ω-*N*G,*N'*<sup>G</sup>-dimethylarginine (SDMA). Therefore, three distinct types of methylated arginine residues (MMA, ADMA and SDMA) are present on a horde of different proteins in the cytosol, nucleus and organelles of mam‐ malian cells, ADMA being the most prevalent [3].

**Figure 2.** Schematic representation of the human protein arginine methyltransferase (PRMTs) family. For each mem‐ ber, the length of the longest isoform is indicated on the right. All members of the family possess at least one con‐ served MTase domain with signature motifs I, post-I, II, and III and a THW loop. Additional domains are marked in yellow boxes: SH3, ZnF (zinc finger), Myr (myristoylation), F-box, TPR (tetratricopeptide) and NosD (nitrous oxidase ac‐ cessory protein). Adapted from [17].

**Figure 1.** Methylation of the arginine side chain in proteins by protein arginine methyltransferases (PRMTs). The argi‐ nine residue holds 5 potential hydrogen bond donors. Mammalian PRMTs use the methyl group from a molecule of Sadenosyl-L-methionine (AdoMet) to form ω-NG-monomethylarginine (MMA). Subsequently, type I PRMTs add a methyl group to the same nitrogen atom forming ω-NG,NG-dimethylarginine (ADMA), whereas type II PRMTs generate

Methylation of arginine is catalyzed by protein arginine methyltransferases (PRMTs). PRMTs catalyze the transfer of methyl groups from the universal methyl donor, S-adenosyl-L-methionine (AdoMet), to the guanidine nitrogen atom of the target arginine residues with‐

The first documented protein with arginine methylation activity, then termed protein meth‐ ylase I, was purified from calf thymus [4, 5]. Additional family members were subsequently identified and purified from different tissues and cell lines [6-16]. In mammals, PRMTs are grouped in two major classes: type I and type II. Both PRMT types catalyze the transfer of a single methyl group from AdoMet to an arginine residue in the target protein, producing ω-*N*G-monomethylarginine (MMA) (Figure 1). However, differences exist when adding the second methyl group: type I members add it on the same previously methylated nitrogen,

ω-NG,N'G-dimethylarginine (SDMA).

in proteins [3].

**2. The mammalian PRMTs: A brief overview**

92 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

To date, the human PRMT family includes eleven enzymes, termed from PRMT1 to PRMT11. Six are classified as type I (PRMT1, PRMT2, PRMT3, PRMT4, PRMT6 and PRMT8), and two as type II (PRMT5 and PRMT9). The classification of PRMT7 is controversial, as described below. The remaining two proteins, PRMT10 and PRMT11, were identified by their homology with PRMT7 and PRMT9, respectively, but their PRMT activity has not been sustained by experimental evidence [18, 19]. The genes encoding the different PRMTs are all located on different chromosomes, except the ones coding for PRMTs 1 and 4, which are both located in chromosome 19. Different splice variants coding for the corresponding isoforms are recognized for all PRMTs. Moreover, different cellular localizations are observed among them.

Although the human PRMT members vary in length from 316 to 956 amino acid residues, they all contain a highly conserved catalytic core region of around 300 residues (Figure 2). However, each PRMT presents a unique N-terminal region of variable length and distinct domain motifs, as for example PRMT3 and PRMT8 that bear a zinc finger (ZnF) and a myristoylation domain, respectively. Interestingly PRMT7 and PRMT10 exhibit a second catalytic domain. Until present, only the 3D structures of PRMT1, PRMT3 and PRMT5 were solved by X-ray crystallography [20-22]. These structures revealed that, also at the structural level, the catalytic core is highly conserved. Three structural regions were identified, namely, a methyltransferase (MTase) domain, a β-barrel domain unique to the PRMT family and a dimerization arm. The MTase domain is located in the Nterminal region of the catalytic core. It consists of a α/β Rossmann fold, typical of AdoMet-dependent methyltransferases, containing the conserved motifs I, post-I, II and III [23]. Motif I includes in its C-terminal part the signature sequences for most classes of MTases (GxGxG) involved in AdoMet binding. A relevant structural element localized in the MTase domain is the double-E loop (containing two invariant glutamate (E) residues). The C-terminal part of the catalytic core is a β-barrel domain containing 10 β-strands and a THW (threonine-histidine-tryptophan) loop (the most highly conserved sequence of this domain). Within the overall structure, the THW loop is located next to the double-E loop in the AdoMet binding domain, forming the active site. The dimerization arm is formed by a three-helix segment, which is inserted between strands 6 and 7 of the β-barrel domain and is responsible for dimerization, essential for enzyme activity [21].

PRMT4, or CARM1, was first identified as a steroid receptor coactivator [11], providing the first evidence that protein arginine methylation participates in the regulation of gene expres‐ sion (please, see sections *3.2.1 and 3.2.2.1*.). PRMT4 is ubiquitously expressed, exhibits a nu‐ clear localization, and presents restricted substrate specificity. Specifically, in addition to nuclear hormone receptors and histones, it targets splicing factors [39, 40] and ATP-remod‐ eling factors [41]. As opposed to most PRMTs, and as already referred, PRMT4 does not methylate GAR sequences. PRMT4 participates in many biological processes, including ear‐ ly T cell development [42], adipocyte differentiation [43], endochondral ossification [44], and proliferation and differentiation of pulmonary epithelial cells [45]. Embryos of *PRMT4 null* mice are small in size and die perinatally, thus showing the importance of PRMT4 activity to

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PRMT5 was the first PRMT type II to be identified. PRMT5 is widely expressed in mammals, and more extensively in heart, muscle and testis [12]. PRMT5 is located both in the nucleus and in the cytoplasm [47]. Interestingly, PRMT5 is localized in the Golgi apparatus (GA) and its loss disrupts the GA structure, thereby suggesting that this enzyme plays a role in the maintenance of GA architecture [47]. In addition, PRMT5 is involved in the maintenance of the spliceosome integrity [48, 49]. PRMT5 targets histones, transcriptional elongation factors, chromatin remodelers and co-repressors [50]. In mice, loss of PRMT5 results in early embry‐

PRMT6 is the smallest family member (316 amino acids) and is localized predominantly in the nucleus [13]. Little is known about its properties, besides the fact that it produces asymmetrically dimethylated arginine residues. Similarly, PRMT6 functions are not completely understood. However, a role for PRMT6 in the regulation of cell proliferation and senescence, through transcriptional repression of tumor suppressor genes, has just

PRMT7 is present mainly in thymus, dendritic cells and testis [15, 53]. It has both nuclear and cytosolic localizations [15]. PRMT7 is involved in the modulation of sensitivity to DNA damaging agents [54, 55], methylation of male germline imprinted genes [56], and embryon‐ ic stem cell pluripotency [57]. Although Lee and colleagues characterized PRMT7 as a type II enzyme capable of forming SDMA [15], this finding was recently contested. In fact, Zuri‐ ta-Lopez and coworkers have just reported that PRMT7 produces exclusively MMA resi‐ dues, constituting the sole mammalian PRMT member with this type of methyltransferase

PRMT8 is the type I PRMT with the highest degree of homology with PRMT1, but, unlike the latter, is mostly found in brain [14, 33]. PRMT8 possesses a myristoylation motif at its Nterminal end, which facilitates electrostatic interactions with membrane lipids and renders its distinct plasma membrane localization [14, 33]. Interestingly, its activity is regulated by the conformation of its N-terminal end, and removal of this domain results in enhancement of its enzymatic activity [33]. PRMT8 targets the pro-oncoprotein encoded by the Ewing sar‐

coma (EWS) gene, which contains a GAR motif in its C-terminal terminus [59].

life [46].

onic lethality [51].

been reported [52].

activity [58].

Most PRMTs methylate arginine residues localized within glycine- and arginine-rich (GAR) sequences, but there are exceptions to this rule [24]. For instance, PRMT4 cannot methylate GAR motifs, and PRMT5 methylates both GAR and non-GAR motifs. Other factors, such as the accessibility of the target arginine and the conformation of the involved sequence, also govern substrate methylation.

PRMT1 was the first mammalian PRMT to be cloned [25] and is the predominant type I en‐ zyme [26] found in all embryonic and adult tissues examined so far [27]. Due to alternative splicing, there are seven isoforms of the protein, all varying in their N-terminal domain, which are expressed in a tissue-specific manner and have distinct subcellular localization patterns [28]. PRMT1 has broad substrate specificity with over 40 targets, most being RNA processing proteins, and multiple interacting partners [29]. Targeted PRMT1 knockout in mice results in embryonic lethality, thus showing that loss of PRMT1 activity is incompati‐ ble with life [30].

PRMT2 transcripts have been found in most human tissues, with highest levels in heart, prostate, ovary and neuronal system [10, 31]. PRMT2 was found predominantly in the nu‐ cleus and to a lower degree in the cytoplasm of mammalian cells [17]. Only recently, based on the observation that it catalyzes the formation of MMA and ADMA residues on histone H4, PRMT2 was recognized as a type I enzyme [32]. PRMT2 may act in cooperation with PRMT8, since its SH3 domain binds the N-terminal domain of the latter enzyme [33]. Some of the known targets of PRMT2, besides the aforementioned histone H4 [32], are the STAT3 (signal transducer and activator of transcription 3) protein [34], the estrogen receptor alpha [35], the androgen receptor [31], the retinoblastoma gene product [36], and the heterogene‐ ous nuclear ribonucleoprotein (hnRNP) E1B-AP5 [37]. In contrast with PRMT1 knockouts, *PRMT2 null* mice are viable and grow normally [34].

PRMT3 was first identified as a PRMT1-binding partner. It is widely expressed in human tissues and has a predominantly cytosolic subcellular localization [9]. PRMT3 possesses a zinc finger domain that assists in its binding to ribosomal proteins, including the S2 protein of the small ribosomal subunit. Mouse embryos with a targeted disruption of PRMT3 are small in size but survive after birth and attain a normal size in adulthood [38].

PRMT4, or CARM1, was first identified as a steroid receptor coactivator [11], providing the first evidence that protein arginine methylation participates in the regulation of gene expres‐ sion (please, see sections *3.2.1 and 3.2.2.1*.). PRMT4 is ubiquitously expressed, exhibits a nu‐ clear localization, and presents restricted substrate specificity. Specifically, in addition to nuclear hormone receptors and histones, it targets splicing factors [39, 40] and ATP-remod‐ eling factors [41]. As opposed to most PRMTs, and as already referred, PRMT4 does not methylate GAR sequences. PRMT4 participates in many biological processes, including ear‐ ly T cell development [42], adipocyte differentiation [43], endochondral ossification [44], and proliferation and differentiation of pulmonary epithelial cells [45]. Embryos of *PRMT4 null* mice are small in size and die perinatally, thus showing the importance of PRMT4 activity to life [46].

were identified, namely, a methyltransferase (MTase) domain, a β-barrel domain unique to the PRMT family and a dimerization arm. The MTase domain is located in the Nterminal region of the catalytic core. It consists of a α/β Rossmann fold, typical of AdoMet-dependent methyltransferases, containing the conserved motifs I, post-I, II and III [23]. Motif I includes in its C-terminal part the signature sequences for most classes of MTases (GxGxG) involved in AdoMet binding. A relevant structural element localized in the MTase domain is the double-E loop (containing two invariant glutamate (E) residues). The C-terminal part of the catalytic core is a β-barrel domain containing 10 β-strands and a THW (threonine-histidine-tryptophan) loop (the most highly conserved sequence of this domain). Within the overall structure, the THW loop is located next to the double-E loop in the AdoMet binding domain, forming the active site. The dimerization arm is formed by a three-helix segment, which is inserted between strands 6 and 7 of the β-barrel

94 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

domain and is responsible for dimerization, essential for enzyme activity [21].

govern substrate methylation.

*PRMT2 null* mice are viable and grow normally [34].

ble with life [30].

Most PRMTs methylate arginine residues localized within glycine- and arginine-rich (GAR) sequences, but there are exceptions to this rule [24]. For instance, PRMT4 cannot methylate GAR motifs, and PRMT5 methylates both GAR and non-GAR motifs. Other factors, such as the accessibility of the target arginine and the conformation of the involved sequence, also

PRMT1 was the first mammalian PRMT to be cloned [25] and is the predominant type I en‐ zyme [26] found in all embryonic and adult tissues examined so far [27]. Due to alternative splicing, there are seven isoforms of the protein, all varying in their N-terminal domain, which are expressed in a tissue-specific manner and have distinct subcellular localization patterns [28]. PRMT1 has broad substrate specificity with over 40 targets, most being RNA processing proteins, and multiple interacting partners [29]. Targeted PRMT1 knockout in mice results in embryonic lethality, thus showing that loss of PRMT1 activity is incompati‐

PRMT2 transcripts have been found in most human tissues, with highest levels in heart, prostate, ovary and neuronal system [10, 31]. PRMT2 was found predominantly in the nu‐ cleus and to a lower degree in the cytoplasm of mammalian cells [17]. Only recently, based on the observation that it catalyzes the formation of MMA and ADMA residues on histone H4, PRMT2 was recognized as a type I enzyme [32]. PRMT2 may act in cooperation with PRMT8, since its SH3 domain binds the N-terminal domain of the latter enzyme [33]. Some of the known targets of PRMT2, besides the aforementioned histone H4 [32], are the STAT3 (signal transducer and activator of transcription 3) protein [34], the estrogen receptor alpha [35], the androgen receptor [31], the retinoblastoma gene product [36], and the heterogene‐ ous nuclear ribonucleoprotein (hnRNP) E1B-AP5 [37]. In contrast with PRMT1 knockouts,

PRMT3 was first identified as a PRMT1-binding partner. It is widely expressed in human tissues and has a predominantly cytosolic subcellular localization [9]. PRMT3 possesses a zinc finger domain that assists in its binding to ribosomal proteins, including the S2 protein of the small ribosomal subunit. Mouse embryos with a targeted disruption of PRMT3 are

small in size but survive after birth and attain a normal size in adulthood [38].

PRMT5 was the first PRMT type II to be identified. PRMT5 is widely expressed in mammals, and more extensively in heart, muscle and testis [12]. PRMT5 is located both in the nucleus and in the cytoplasm [47]. Interestingly, PRMT5 is localized in the Golgi apparatus (GA) and its loss disrupts the GA structure, thereby suggesting that this enzyme plays a role in the maintenance of GA architecture [47]. In addition, PRMT5 is involved in the maintenance of the spliceosome integrity [48, 49]. PRMT5 targets histones, transcriptional elongation factors, chromatin remodelers and co-repressors [50]. In mice, loss of PRMT5 results in early embry‐ onic lethality [51].

PRMT6 is the smallest family member (316 amino acids) and is localized predominantly in the nucleus [13]. Little is known about its properties, besides the fact that it produces asymmetrically dimethylated arginine residues. Similarly, PRMT6 functions are not completely understood. However, a role for PRMT6 in the regulation of cell proliferation and senescence, through transcriptional repression of tumor suppressor genes, has just been reported [52].

PRMT7 is present mainly in thymus, dendritic cells and testis [15, 53]. It has both nuclear and cytosolic localizations [15]. PRMT7 is involved in the modulation of sensitivity to DNA damaging agents [54, 55], methylation of male germline imprinted genes [56], and embryon‐ ic stem cell pluripotency [57]. Although Lee and colleagues characterized PRMT7 as a type II enzyme capable of forming SDMA [15], this finding was recently contested. In fact, Zuri‐ ta-Lopez and coworkers have just reported that PRMT7 produces exclusively MMA resi‐ dues, constituting the sole mammalian PRMT member with this type of methyltransferase activity [58].

PRMT8 is the type I PRMT with the highest degree of homology with PRMT1, but, unlike the latter, is mostly found in brain [14, 33]. PRMT8 possesses a myristoylation motif at its Nterminal end, which facilitates electrostatic interactions with membrane lipids and renders its distinct plasma membrane localization [14, 33]. Interestingly, its activity is regulated by the conformation of its N-terminal end, and removal of this domain results in enhancement of its enzymatic activity [33]. PRMT8 targets the pro-oncoprotein encoded by the Ewing sar‐ coma (EWS) gene, which contains a GAR motif in its C-terminal terminus [59].

PRMT9 was identified as a putative arginine methyltransferase and was classified as a type II PRMT due to its ability to catalyze the formation of MMA and ADMA [16]. However, there is limited sequence homology between PRMT9 and other human PRMTs [17]. Like most of the PRMTs, PRMT9 is widely expressed in mammalian tissues. PRMT9 has both nuclear and cytoplasmatic localization [17]. Several PRMT9 isoforms are recognized and alternate splicing may be an important mechanism to regulate methylation by PRMT9 [16]. So far, the only known substrates for PRMT9 are histones, the maltose binding protein and several peptides [16]. Disruption of the PRMT9 activity has been associated with inflammation of the middle ear, indicating the importance of methylation in disease [60, 61].

**3.2. Transcriptional regulation**

*3.2.1. Histone arginine methylation*

Protein arginine methylation was initially detected in histones and studied mainly in relation to transcriptional regulation. More recently, the presence of methylated arginines in a large num‐ ber of non-histonic proteins that also impact on transcriptional regulation was recognized.

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Methylation of histones at arginine residues has been very well documented and subject to periodic review [74-76]. Therefore, a comprehensive review of histone arginine methylation

A few arginine residues within the N-terminal tail of histones H3, H4 and H2A are subject to methylation. The N-terminal tail of histone H3 contains several arginine residues that are targeted by different PRMTs,1 while only one residue in histone H4 (H4R3) is subject to argi‐ nine methylation.2 Since the first residues of the N-terminal tails of histones H4 and H2A are identical, residue 3 of histone H2A may be methylated in the same manner as H4R3 [76]. Interestingly, histone asymmetrical dimethylation generally activates gene transcription, while generation of SDMA is normally a repressive mark, although exceptions to this rule exist [75]. Arginine methylation occurring in histone tails may either affect other PTMs or

In addition to histones, PRMTs also target other proteins that are involved in the transcrip‐ tional control of gene expression in mammals. These proteins include several types of tran‐ scription factors, transcription elongation factors, methyl DNA-binding proteins, and RNA polymerase II, as presented in the following sub-sections. In each, we choose to use some illustrative examples, rather than present an exhaustive list of all non-histonic targets for

Transcriptional coregulators are well recognized targets of PRMTs. Methylation of nuclear hormone receptors (NRs) illustrate well how arginine methylation of transcriptional coregu‐ lators may affect gene transcription. NRs are a large group of structurally related transcrip‐ tion factors that are responsible for mediating the biological effects of hormones. Transcription activation by NRs is a multistep process involving transcriptional coactivator proteins, as CREB-binding protein (CBP) and p300. These two co-activators have similar structures, sharing one N-terminal KIX domain that displays several arginine residues that are subject of methylation by PRMT4 [83]. CBP/p300 exhibits HAT (histone acetyl transfer‐

1 In histone H3, residues 17 (H3R17) and 26 (H3R26) are methylated by PRMT4 [77], H3R2 is methylated by PRMT6

2 In histone H4, one arginine residue (H4R3) is subject to methylation. Asymmetric dimethylation of this residue is catalyzed by PRMT1, PRMT6 and PRMT8 [80-82], while symmetric dimethylation is catalyzed by PRMT5 and PRMT7

would go beyond the scope of this paper and will be depicted in general terms only.

influence the docking of key transcriptional effector molecules [76].

*3.2.2. Arginine methylation of non-histonic proteins*

PRMTs that impact transcriptional regulation.

*3.2.2.1. Transcription factors*

[78], and H3R8 is methylated by PRMT5 [79].

[56, 79].

PRMT10 and PRMT11 were identified by their homology with PRMT7 and PRMT9, respec‐ tively, but, as referred to above, their PRMT activity has not been sustained by experimental evidence.

#### **3. Physiological roles of protein arginine methylation**

Following the advent of improved sensitivity conquered with mass spectrometry, current proteomic technologies have been uncovering an increasingly large fraction of the human proteome as substrates for PRMTs. Additionally, advances in molecular biology techniques and the generation of mice with targeted deletion of the various members of the PRMTs family have been disclosing the functional relevance of this PTM. As new substrates are be‐ ing characterized, a broadening spectrum of PRMT-regulated cellular processes is being re‐ alized. So far, targeted cellular processes can be loosely categorized in RNA processing, transcriptional regulation, DNA repair and signal transduction.

#### **3.1. RNA processing**

Most proteins targeted by PRMTs are RNA-binding proteins (RBPs) harboring GAR motifs, which are involved in all aspects of RNA metabolism [29]. During eukaryotic transcription, RBPs associate with the nascent pre-mRNA allowing a series of RNA processing events to take place, including 5'-end capping, splicing, 3'-end cleavage and polyadenylation [62].

The importance of protein arginine methylation for RNA processing was first revealed by the observation that in HeLa cells splicing reactions were completely inhibited by a pan SDMA-specific antibody [63]. Subsequent studies have shown that a number of spliceoso‐ mal proteins are subject to both asymmetrical and symmetrical dimethylation, uncovering the crucial role of protein arginine methylation in the assembly of the spliceosome [64-66]. Other RBPs (hnRNP1, fibrillarin, nucleolin, HuD and HuR) have also been shown to contain methylated arginine residues [67-70]. Importantly, several of these RBPs lack proper subcel‐ lular localization if hypomethylated [71, 72]. For instance, asymmetrical dimethylation of specific arginine residues comprising the nuclear poly(A) binding protein prevents its trans‐ port to the cytoplasm [73].

#### **3.2. Transcriptional regulation**

PRMT9 was identified as a putative arginine methyltransferase and was classified as a type II PRMT due to its ability to catalyze the formation of MMA and ADMA [16]. However, there is limited sequence homology between PRMT9 and other human PRMTs [17]. Like most of the PRMTs, PRMT9 is widely expressed in mammalian tissues. PRMT9 has both nuclear and cytoplasmatic localization [17]. Several PRMT9 isoforms are recognized and alternate splicing may be an important mechanism to regulate methylation by PRMT9 [16]. So far, the only known substrates for PRMT9 are histones, the maltose binding protein and several peptides [16]. Disruption of the PRMT9 activity has been associated with inflammation of the middle ear, indicating the importance of

PRMT10 and PRMT11 were identified by their homology with PRMT7 and PRMT9, respec‐ tively, but, as referred to above, their PRMT activity has not been sustained by experimental

Following the advent of improved sensitivity conquered with mass spectrometry, current proteomic technologies have been uncovering an increasingly large fraction of the human proteome as substrates for PRMTs. Additionally, advances in molecular biology techniques and the generation of mice with targeted deletion of the various members of the PRMTs family have been disclosing the functional relevance of this PTM. As new substrates are be‐ ing characterized, a broadening spectrum of PRMT-regulated cellular processes is being re‐ alized. So far, targeted cellular processes can be loosely categorized in RNA processing,

Most proteins targeted by PRMTs are RNA-binding proteins (RBPs) harboring GAR motifs, which are involved in all aspects of RNA metabolism [29]. During eukaryotic transcription, RBPs associate with the nascent pre-mRNA allowing a series of RNA processing events to take place, including 5'-end capping, splicing, 3'-end cleavage and polyadenylation [62].

The importance of protein arginine methylation for RNA processing was first revealed by the observation that in HeLa cells splicing reactions were completely inhibited by a pan SDMA-specific antibody [63]. Subsequent studies have shown that a number of spliceoso‐ mal proteins are subject to both asymmetrical and symmetrical dimethylation, uncovering the crucial role of protein arginine methylation in the assembly of the spliceosome [64-66]. Other RBPs (hnRNP1, fibrillarin, nucleolin, HuD and HuR) have also been shown to contain methylated arginine residues [67-70]. Importantly, several of these RBPs lack proper subcel‐ lular localization if hypomethylated [71, 72]. For instance, asymmetrical dimethylation of specific arginine residues comprising the nuclear poly(A) binding protein prevents its trans‐

**3. Physiological roles of protein arginine methylation**

transcriptional regulation, DNA repair and signal transduction.

methylation in disease [60, 61].

96 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

evidence.

**3.1. RNA processing**

port to the cytoplasm [73].

Protein arginine methylation was initially detected in histones and studied mainly in relation to transcriptional regulation. More recently, the presence of methylated arginines in a large num‐ ber of non-histonic proteins that also impact on transcriptional regulation was recognized.

#### *3.2.1. Histone arginine methylation*

Methylation of histones at arginine residues has been very well documented and subject to periodic review [74-76]. Therefore, a comprehensive review of histone arginine methylation would go beyond the scope of this paper and will be depicted in general terms only.

A few arginine residues within the N-terminal tail of histones H3, H4 and H2A are subject to methylation. The N-terminal tail of histone H3 contains several arginine residues that are targeted by different PRMTs,1 while only one residue in histone H4 (H4R3) is subject to argi‐ nine methylation.2 Since the first residues of the N-terminal tails of histones H4 and H2A are identical, residue 3 of histone H2A may be methylated in the same manner as H4R3 [76]. Interestingly, histone asymmetrical dimethylation generally activates gene transcription, while generation of SDMA is normally a repressive mark, although exceptions to this rule exist [75]. Arginine methylation occurring in histone tails may either affect other PTMs or influence the docking of key transcriptional effector molecules [76].

#### *3.2.2. Arginine methylation of non-histonic proteins*

In addition to histones, PRMTs also target other proteins that are involved in the transcrip‐ tional control of gene expression in mammals. These proteins include several types of tran‐ scription factors, transcription elongation factors, methyl DNA-binding proteins, and RNA polymerase II, as presented in the following sub-sections. In each, we choose to use some illustrative examples, rather than present an exhaustive list of all non-histonic targets for PRMTs that impact transcriptional regulation.

#### *3.2.2.1. Transcription factors*

Transcriptional coregulators are well recognized targets of PRMTs. Methylation of nuclear hormone receptors (NRs) illustrate well how arginine methylation of transcriptional coregu‐ lators may affect gene transcription. NRs are a large group of structurally related transcrip‐ tion factors that are responsible for mediating the biological effects of hormones. Transcription activation by NRs is a multistep process involving transcriptional coactivator proteins, as CREB-binding protein (CBP) and p300. These two co-activators have similar structures, sharing one N-terminal KIX domain that displays several arginine residues that are subject of methylation by PRMT4 [83]. CBP/p300 exhibits HAT (histone acetyl transfer‐

<sup>1</sup> In histone H3, residues 17 (H3R17) and 26 (H3R26) are methylated by PRMT4 [77], H3R2 is methylated by PRMT6 [78], and H3R8 is methylated by PRMT5 [79].

<sup>2</sup> In histone H4, one arginine residue (H4R3) is subject to methylation. Asymmetric dimethylation of this residue is catalyzed by PRMT1, PRMT6 and PRMT8 [80-82], while symmetric dimethylation is catalyzed by PRMT5 and PRMT7 [56, 79].

ase) and HMT (histone methyl transferase) activities. Both CBP/p300 and PRMT4 are recruit‐ ed to DNA by the NR bound to its ligand, where they stimulate transcription by remodeling chromatin through subsequent histone acetylation and methylation in the vicinity of hor‐ mone response element (HRE). Notably, it was shown that CBP/p300 activates NR-depend‐ ent gene transcription only if methylated in their KIX domain [83].

of a single residue in the CTD (R1810) was documented and implied in the regulation of

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99

Genome integrity is constantly subject to monitoring by a complicated and entangled network of mechanisms collectively termed as DNA damage response (DDR). Some pro‐ teins integrating this system are known targets of PRMTs. Mre11 is part of a complex that is critically implicated in homologous recombination repair of DNA double strand breaks. This protein harbors a GAR motif that is methylated by PRMT1, allowing its as‐ sociation with nuclear structures and recruitment to sites of DNA damage [90], as well as its exonuclease activity [91]. PRMT5-induced methylation of Rad9, a cell cycle check‐ point protein, enables activation of Chk1, an important downstream checkpoint effector, and loss of Rad9 methylation leads to S/M and G2/M checkpoint defects [92]. DNA poly‐ merase beta, which participates in DNA base excision repair, is methylated by PRMT6,

In addition to regulating numerous nuclear processes, arginine methylation is involved in conveying information from the cell surface, through the cytoplasm, to the nucleus. For in‐ stance, the implication of protein arginine methylation in interferon (IFN) signaling has been substantiated by the fact that PRMT1 interacts with the cytoplasmic domain of the type I IFN receptor [94], and that its depletion impairs the biological activity of type I IFN [95]. The involvement of arginine methylation within T cell receptor signaling cascades is also gaining prominence. In fact, stimulation of T helper cells induces arginine methylation of several cy‐ toplasmic proteins, which when inhibited causes signaling defects in CD4 T cells and severe immunosuppression [96]. PRMT1-mediated methylation also impacts insulin signaling and glucose uptake in skeletal muscle cells [97]. Estrogen rapid signaling is similarly mediated by PRMT1, which targets estrogen receptor alpha (ER-α) [98]. PRMT1 also methylates FoxO1, a member of the mammalian forkhead transcription factors of class O (FoxO) sub‐ family [99]. These transcription factors are involved in many vital processes (e.g. apoptosis, cell-cycle control, glucose metabolism, oxidative stress resistance and longevity) and, in re‐ sponse to insulin or several growth factors, are negatively regulated through phosphoryla‐ tion by serine-threonine kinase Akt/protein kinase B (Akt/PKB) (that results in their export from the nucleus to the cytoplasm) [100]. PRMT1 methylates FoxO1 at arginine residues within the Akt consensus motif, blocking the Akt-mediated phosphorylation and augment‐ ing FoxO1 target gene expression [99]. Another Akt-target protein is BCL-2, antagonist of cell death, which is also methylated by PRMT1 on its consensus phosphorylation motif [101]. Methylation-phosphorylation switches are presently regarded as paradigms operating in signal transduction pathways and other physiological processes, since numerous other examples of this crosstalk have been described. The implication of PRMTs other than PRMT1 in signaling routes has not been well studied. The association of PRMT8 with the plasma membrane provides a clue that this enzyme may facilitate the triggering of a signal‐

transcription driven by RNA polymerase II [89].

which enhances its binding to DNA and its processivity [93].

**3.3. DNA repair**

**3.4. Signal transduction**

Peroxisome proliferator-activated receptor-gamma coactivator (PGC) 1 alpha (PGC-1α) is other example of a coactivator of NRs whose activity was clearly shown to be dependent on PRMT activity. PGC-1α signaling is extremely important in the heart, where PGC-1α is highly expressed and controls cardiac energy pathways during development and in re‐ sponse to stressors. Interestingly, it was shown that PRMT1 activates PGC-1α through its methylation at several arginine residues [84]. Remarkably, experimentally based evidence has recently linked decreased levels of methyl-PGC-1α due to decreased PRMT1 activity to cardiomyopathy in mice [85].

Arginine methylation may also impact transcriptional activity by targeting transcription fac‐ tors that bind directly to DNA, like the STAT1 protein. STAT1 (signal transducer and activa‐ tor of transcription) is a member of the STAT family and mediates the cellular response to cytokines and growth factors thus regulating many aspects of growth, survival and differen‐ tiation of cells. After activation by cytoplasmic phosphorylation, STAT1 migrates to the nu‐ cleus, where it binds to the enhancer elements of response genes. The N-terminal region of STAT1 comprises a conserved arginine residue that was shown to be a PRMT1 target [86]. Inhibition of PRMT1 activity hinders the DNA binding activity of STAT1 and STAT1-medi‐ ated transcription.

#### *3.2.2.2. Transcription elongation factors*

Activity of PRMTs may also affect transcription by marking transcription elongation factors, like Spt5. The Spt5 protein displays several KOW domains, which contain consensus sites for arginine methylation and overlap the RNA polymerase II binding domain. Notably, Spt5 can be methylated in its RNA polymerase II binding domain by PRMT1 or PRMT5 and this methylation may cause transcriptional pausing [87].

#### *3.2.2.3. Methyl DNA-binding proteins*

PRMTs may also participate in the DNA methylation system of chromatin control. DNA methylation, an important regulator of gene transcription, occurs within a CpG context in differentiated mammalian cells. MBD2 is a methyl DNA-associated protein that docks me‐ thylated CpGs in DNA and negatively affects gene transcription by recruiting proteins that repress chromatin. Interestingly, both PRMT1 and PRMT5 methylate MBD2, thereby impair‐ ing the transcription repression function of MBD2 [88].

#### *3.2.2.4. RNA polymerase II*

In eukaryotic cells, the C-terminal domain (CTD) of RNA polymerase II provides a platform to recruit different regulators of the transcription apparatus. Recently, arginine methylation of a single residue in the CTD (R1810) was documented and implied in the regulation of transcription driven by RNA polymerase II [89].

#### **3.3. DNA repair**

ase) and HMT (histone methyl transferase) activities. Both CBP/p300 and PRMT4 are recruit‐ ed to DNA by the NR bound to its ligand, where they stimulate transcription by remodeling chromatin through subsequent histone acetylation and methylation in the vicinity of hor‐ mone response element (HRE). Notably, it was shown that CBP/p300 activates NR-depend‐

Peroxisome proliferator-activated receptor-gamma coactivator (PGC) 1 alpha (PGC-1α) is other example of a coactivator of NRs whose activity was clearly shown to be dependent on PRMT activity. PGC-1α signaling is extremely important in the heart, where PGC-1α is highly expressed and controls cardiac energy pathways during development and in re‐ sponse to stressors. Interestingly, it was shown that PRMT1 activates PGC-1α through its methylation at several arginine residues [84]. Remarkably, experimentally based evidence has recently linked decreased levels of methyl-PGC-1α due to decreased PRMT1 activity to

Arginine methylation may also impact transcriptional activity by targeting transcription fac‐ tors that bind directly to DNA, like the STAT1 protein. STAT1 (signal transducer and activa‐ tor of transcription) is a member of the STAT family and mediates the cellular response to cytokines and growth factors thus regulating many aspects of growth, survival and differen‐ tiation of cells. After activation by cytoplasmic phosphorylation, STAT1 migrates to the nu‐ cleus, where it binds to the enhancer elements of response genes. The N-terminal region of STAT1 comprises a conserved arginine residue that was shown to be a PRMT1 target [86]. Inhibition of PRMT1 activity hinders the DNA binding activity of STAT1 and STAT1-medi‐

Activity of PRMTs may also affect transcription by marking transcription elongation factors, like Spt5. The Spt5 protein displays several KOW domains, which contain consensus sites for arginine methylation and overlap the RNA polymerase II binding domain. Notably, Spt5 can be methylated in its RNA polymerase II binding domain by PRMT1 or PRMT5 and this

PRMTs may also participate in the DNA methylation system of chromatin control. DNA methylation, an important regulator of gene transcription, occurs within a CpG context in differentiated mammalian cells. MBD2 is a methyl DNA-associated protein that docks me‐ thylated CpGs in DNA and negatively affects gene transcription by recruiting proteins that repress chromatin. Interestingly, both PRMT1 and PRMT5 methylate MBD2, thereby impair‐

In eukaryotic cells, the C-terminal domain (CTD) of RNA polymerase II provides a platform to recruit different regulators of the transcription apparatus. Recently, arginine methylation

ent gene transcription only if methylated in their KIX domain [83].

98 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

cardiomyopathy in mice [85].

ated transcription.

*3.2.2.2. Transcription elongation factors*

*3.2.2.3. Methyl DNA-binding proteins*

*3.2.2.4. RNA polymerase II*

methylation may cause transcriptional pausing [87].

ing the transcription repression function of MBD2 [88].

Genome integrity is constantly subject to monitoring by a complicated and entangled network of mechanisms collectively termed as DNA damage response (DDR). Some pro‐ teins integrating this system are known targets of PRMTs. Mre11 is part of a complex that is critically implicated in homologous recombination repair of DNA double strand breaks. This protein harbors a GAR motif that is methylated by PRMT1, allowing its as‐ sociation with nuclear structures and recruitment to sites of DNA damage [90], as well as its exonuclease activity [91]. PRMT5-induced methylation of Rad9, a cell cycle check‐ point protein, enables activation of Chk1, an important downstream checkpoint effector, and loss of Rad9 methylation leads to S/M and G2/M checkpoint defects [92]. DNA poly‐ merase beta, which participates in DNA base excision repair, is methylated by PRMT6, which enhances its binding to DNA and its processivity [93].

#### **3.4. Signal transduction**

In addition to regulating numerous nuclear processes, arginine methylation is involved in conveying information from the cell surface, through the cytoplasm, to the nucleus. For in‐ stance, the implication of protein arginine methylation in interferon (IFN) signaling has been substantiated by the fact that PRMT1 interacts with the cytoplasmic domain of the type I IFN receptor [94], and that its depletion impairs the biological activity of type I IFN [95]. The involvement of arginine methylation within T cell receptor signaling cascades is also gaining prominence. In fact, stimulation of T helper cells induces arginine methylation of several cy‐ toplasmic proteins, which when inhibited causes signaling defects in CD4 T cells and severe immunosuppression [96]. PRMT1-mediated methylation also impacts insulin signaling and glucose uptake in skeletal muscle cells [97]. Estrogen rapid signaling is similarly mediated by PRMT1, which targets estrogen receptor alpha (ER-α) [98]. PRMT1 also methylates FoxO1, a member of the mammalian forkhead transcription factors of class O (FoxO) sub‐ family [99]. These transcription factors are involved in many vital processes (e.g. apoptosis, cell-cycle control, glucose metabolism, oxidative stress resistance and longevity) and, in re‐ sponse to insulin or several growth factors, are negatively regulated through phosphoryla‐ tion by serine-threonine kinase Akt/protein kinase B (Akt/PKB) (that results in their export from the nucleus to the cytoplasm) [100]. PRMT1 methylates FoxO1 at arginine residues within the Akt consensus motif, blocking the Akt-mediated phosphorylation and augment‐ ing FoxO1 target gene expression [99]. Another Akt-target protein is BCL-2, antagonist of cell death, which is also methylated by PRMT1 on its consensus phosphorylation motif [101]. Methylation-phosphorylation switches are presently regarded as paradigms operating in signal transduction pathways and other physiological processes, since numerous other examples of this crosstalk have been described. The implication of PRMTs other than PRMT1 in signaling routes has not been well studied. The association of PRMT8 with the plasma membrane provides a clue that this enzyme may facilitate the triggering of a signal‐ ing pathway [14]. A recent study has uncovered that PRMT5 regulates the amplitude of the RAS to extracellular signal-regulated kinase (ERK) signal transduction cascade, impacting on cell fate decision [102].

from the methylated substrate. Current view holds that JMJD6 may be a bifunctional en‐

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**Figure 3.** Reversing or annulling protein arginine methylation: demethylation and citrullination reactions. (A) Deme‐ thylation of a monomethylarginine (MMA) residue by JMJD6. (B) Citrullination of a MMA residue by PAD4. This reac‐

Monomethylated arginine residues may be converted to citrulline by deimination, a modification catalyzed by peptidylarginine deiminases (PADs). In this process, the guanidinium side chain of an arginine residue is hydrolyzed and methylamine released [112] (Figure 3B). To date, five human PAD homologs have been identified and designated as PAD1-4 and PAD6. Most mechanistic studies on PADs have been performed with PAD4. PADs may be activated by calcium, which seems to be essential for the catalysis to occur [113, 114]. Citrulline residues are found with great extent in

tion antagonizes arginine methylation, since it is not reversible. Adapted from [109].

zyme, catalyzing either demethylation or lysyl-hydroxylation reactions [111].

#### **4. Modulation of PRMTs activity**

PRMTs associate with proteins that may change their activity or substrate specificity. Exam‐ ples of proteins that enhance the activity of a specific PRMT are: BTG1 (B-cell translocation gene 1 protein), BTG2 (B-cell translocation gene 2 protein) and hCAF1 (chemokine receptor 4-associated factor 1), which associate with PRMT1 [25, 103]; the chromatin remodelers BRG and BRM, which enhance PRMT5 activity [79]; and BORIS (brother of the regulator of im‐ printed sites), which has been reported to elevate PRMT7 activity [56]. In contrast, tumor suppressor DAL-1 inhibits the activity of PRMT3 *in vitro* and *in vivo* [104]. PRMT4 associa‐ tion with other proteins within the nucleosomal methylation activator complex (NUMAC) enhances its ability to methylate nucleosomal histone H3, while the free enzyme preferen‐ tially methylates free core histone H3 [41]. Similarly, PRMT5 association with COPR5 shifts its preferential target from H3R8 to H4R3 [105].

PRMT activity may also be modulated by PTMs. For instance, several PRMTs are autome‐ thylated, but the relevancy of this phenomenon is not acquainted [33]. PRMT4 phosphoryla‐ tion during mitosis prevents its homodimerization, thereby decreasing its activity [106]. Additionally, PRMT4 phosphorylation abolishes the binding of AdoMet and thus its activi‐ ty, and promotes its cytoplasm localization [107].

### **5. The reversibility of protein arginine methylation: A fast-growing field of knowledge**

An important question under recent investigation was whether demethylation reactions oc‐ curred to reverse the effects of protein arginine methylation. Initial studies indicated that this PTM was a permanent mark on proteins. As such, the only way to reverse the effects of the presence of this methyl mark would be to degrade the protein to its amino acid compo‐ nents and then make a new unmethylated version by protein synthesis [108]. However, the long-standing notion that arginine methylation is a stable covalent mark has been chal‐ lenged in recent years due to the discovery of two types of enzymes that can remove methyl groups from arginine residues in histones by demethylation and citrullination (Figure 3).

The first evidence of demethylation of arginine residues came from a study by Chang and coworkers [110] that reported that JMJD6, a Jumonji domain-containing protein, demethy‐ lates H3R2 and H4R3 residues that are either asymmetrically or symmetrically dimethylated (Figure 3A). Mass spectrometry of a peptide containing dimethylated arginine residues, which was incubated with JMJD6 and then immunoprecipitated with a monomethylargi‐ nine specific antibody, revealed that the enzyme catalyzed the loss of one methyl group from the methylated substrate. Current view holds that JMJD6 may be a bifunctional en‐ zyme, catalyzing either demethylation or lysyl-hydroxylation reactions [111].

ing pathway [14]. A recent study has uncovered that PRMT5 regulates the amplitude of the RAS to extracellular signal-regulated kinase (ERK) signal transduction cascade, impacting

PRMTs associate with proteins that may change their activity or substrate specificity. Exam‐ ples of proteins that enhance the activity of a specific PRMT are: BTG1 (B-cell translocation gene 1 protein), BTG2 (B-cell translocation gene 2 protein) and hCAF1 (chemokine receptor 4-associated factor 1), which associate with PRMT1 [25, 103]; the chromatin remodelers BRG and BRM, which enhance PRMT5 activity [79]; and BORIS (brother of the regulator of im‐ printed sites), which has been reported to elevate PRMT7 activity [56]. In contrast, tumor suppressor DAL-1 inhibits the activity of PRMT3 *in vitro* and *in vivo* [104]. PRMT4 associa‐ tion with other proteins within the nucleosomal methylation activator complex (NUMAC) enhances its ability to methylate nucleosomal histone H3, while the free enzyme preferen‐ tially methylates free core histone H3 [41]. Similarly, PRMT5 association with COPR5 shifts

PRMT activity may also be modulated by PTMs. For instance, several PRMTs are autome‐ thylated, but the relevancy of this phenomenon is not acquainted [33]. PRMT4 phosphoryla‐ tion during mitosis prevents its homodimerization, thereby decreasing its activity [106]. Additionally, PRMT4 phosphorylation abolishes the binding of AdoMet and thus its activi‐

**5. The reversibility of protein arginine methylation: A fast-growing field**

An important question under recent investigation was whether demethylation reactions oc‐ curred to reverse the effects of protein arginine methylation. Initial studies indicated that this PTM was a permanent mark on proteins. As such, the only way to reverse the effects of the presence of this methyl mark would be to degrade the protein to its amino acid compo‐ nents and then make a new unmethylated version by protein synthesis [108]. However, the long-standing notion that arginine methylation is a stable covalent mark has been chal‐ lenged in recent years due to the discovery of two types of enzymes that can remove methyl groups from arginine residues in histones by demethylation and citrullination (Figure 3).

The first evidence of demethylation of arginine residues came from a study by Chang and coworkers [110] that reported that JMJD6, a Jumonji domain-containing protein, demethy‐ lates H3R2 and H4R3 residues that are either asymmetrically or symmetrically dimethylated (Figure 3A). Mass spectrometry of a peptide containing dimethylated arginine residues, which was incubated with JMJD6 and then immunoprecipitated with a monomethylargi‐ nine specific antibody, revealed that the enzyme catalyzed the loss of one methyl group

on cell fate decision [102].

**4. Modulation of PRMTs activity**

100 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

its preferential target from H3R8 to H4R3 [105].

ty, and promotes its cytoplasm localization [107].

**of knowledge**

**Figure 3.** Reversing or annulling protein arginine methylation: demethylation and citrullination reactions. (A) Deme‐ thylation of a monomethylarginine (MMA) residue by JMJD6. (B) Citrullination of a MMA residue by PAD4. This reac‐ tion antagonizes arginine methylation, since it is not reversible. Adapted from [109].

Monomethylated arginine residues may be converted to citrulline by deimination, a modification catalyzed by peptidylarginine deiminases (PADs). In this process, the guanidinium side chain of an arginine residue is hydrolyzed and methylamine released [112] (Figure 3B). To date, five human PAD homologs have been identified and designated as PAD1-4 and PAD6. Most mechanistic studies on PADs have been performed with PAD4. PADs may be activated by calcium, which seems to be essential for the catalysis to occur [113, 114]. Citrulline residues are found with great extent in histones H2A, H3, and H4 [115]. Importantly, the sites of deimination by PAD4 overlap those of arginine methylation on these histones [112], and an increase in histone citrulline levels correlates with a decrease in methylated arginine levels [116, 117]. However, PADs only catalyze the deimination of MMA residues, and no enzyme has been identified that can convert citrulline back to arginine. Thus, citrullination blocks remethylation, adding a new layer to the regulation of protein arginine methylation [118, 119].

tions, favoring the hydrolysis reaction [126]. Nevertheless, if homocysteine accumulates,

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**Figure 4.** Simplified homocysteine metabolism. Homocysteine (Hcy) results from methionine metabolism as a byproduct of cellular transmethylation reactions. Methionine (Met) is the "entry" point of this pathway and is provided by diet and protein breakdown. S-Adenosylmethionine (AdoMet) is first synthesized from Met by methionine adeno‐ syltransferase (MAT). AdoMet is the methyl group donor in a wide variety of transmethylation reactions, being con‐ verted to S-Adenosylhomocysteine (AdoHcy). AdoHcy is subsequently hydrolyzed to Hcy and adenosine by AdoHcy hydrolase. Hcy can then be converted to cysteine (Cys) through the transsulphuration pathway, remethylated back to

Despite recent controversy, elevated levels of homocysteine in plasma (or hyperhomocystei‐ nemia) are accepted as a risk factor for cardiovascular disease [127, 128]. Notably, despite several potential mechanisms underlying this association having been the focus of intense research, the subject remains to be fully understood [125]. Our group has been actively studying whether a hypomethylating environment associated with hyperhomocysteinemia contributes to the vascular toxicity of homocysteine [129-133]. AdoHcy is a competitive in‐ hibitor of most AdoMet-dependent methyltransferases, since it binds to their active sites with higher affinity than AdoMet [134]. As such, the ratio AdoMet/AdoHcy is taken as an index of cellular methylation capacity. As mentioned above, if homocysteine accumulates, AdoHcy will accumulate as well. Thus, increased homocysteine may be regarded as a cellu‐

lar hypomethylation effector, via AdoHcy accumulation.

AdoHcy will accumulate as well [125].

methionine or exported.

#### **6. The disposal of methylated arginine residues / ADMA and vascular disease**

Proteolysis of arginine-methylated proteins results in free MMA, ADMA and SDMA, which are secreted to the extracellular space [120]. These metabolites can then be taken up by other cells via carriers of the cationic amino acid family [121] or eliminated by renal excretion [122]. In addition, ADMA can be hydrolyzed to citrulline and dimethylamine, by dimethy‐ larginine dimethylaminohydrolase (DDAH), which is present in kidney, liver and vascula‐ ture [123]. Interestingly, MMA and ADMA are endogenous inhibitors of nitric oxide (NO) production in endothelial cells. Due to the paramount role of NO on the homeostasis of the vascular endothelium, ADMA, which is present in plasma in much larger quantities than MMA, has emerged as a novel risk factor in the setting of endothelial dysfunction and vas‐ cular disease [124]. As stated before, PRMTs are ubiquitously expressed and participate in crucial cellular processes. Thus, deregulation of these enzymes is likely implicated in the pathogenesis of a number of different diseases, including cardiovascular disease. In addition to the intricate set of potential interactions between protein arginine methylation and cardio‐ vascular disease, we must add ADMA, an additional player that may impact the cardiovas‐ cular system.

#### **7. Protein arginine hypomethylation in the context of hyperhomocysteinemia: "Another brick in the wall"?**

As illustrated in Figure 1, the methyl group responsible for the establishment of protein ar‐ ginine methylation patterns by PRMTs originates from AdoMet, an intermediate in the ho‐ mocysteine metabolism. Therefore, protein arginine methylation and homocysteine metabolism are biochemically linked (Figure 4). In addition to protein arginine methylation, AdoMet serves as the methyl donor for more than one hundred cellular methyl transfer re‐ actions, including the methylation of other residues in proteins (namely, histidine and ly‐ sine), DNA and RNA. Following the transfer of the methyl group, AdoMet is converted into S-adenosyl homocysteine (AdoHcy). AdoHcy is further converted into homocysteine and adenosine by AdoHcy hydrolase, which is widely distributed in mammalian tissues [125]. This reaction is reversible and strongly favors AdoHcy synthesis rather than its hydrolysis; however, both homocysteine and adenosine are rapidly removed under physiological condi‐ tions, favoring the hydrolysis reaction [126]. Nevertheless, if homocysteine accumulates, AdoHcy will accumulate as well [125].

histones H2A, H3, and H4 [115]. Importantly, the sites of deimination by PAD4 overlap those of arginine methylation on these histones [112], and an increase in histone citrulline levels correlates with a decrease in methylated arginine levels [116, 117]. However, PADs only catalyze the deimination of MMA residues, and no enzyme has been identified that can convert citrulline back to arginine. Thus, citrullination blocks remethylation, adding a

**6. The disposal of methylated arginine residues / ADMA and vascular**

Proteolysis of arginine-methylated proteins results in free MMA, ADMA and SDMA, which are secreted to the extracellular space [120]. These metabolites can then be taken up by other cells via carriers of the cationic amino acid family [121] or eliminated by renal excretion [122]. In addition, ADMA can be hydrolyzed to citrulline and dimethylamine, by dimethy‐ larginine dimethylaminohydrolase (DDAH), which is present in kidney, liver and vascula‐ ture [123]. Interestingly, MMA and ADMA are endogenous inhibitors of nitric oxide (NO) production in endothelial cells. Due to the paramount role of NO on the homeostasis of the vascular endothelium, ADMA, which is present in plasma in much larger quantities than MMA, has emerged as a novel risk factor in the setting of endothelial dysfunction and vas‐ cular disease [124]. As stated before, PRMTs are ubiquitously expressed and participate in crucial cellular processes. Thus, deregulation of these enzymes is likely implicated in the pathogenesis of a number of different diseases, including cardiovascular disease. In addition to the intricate set of potential interactions between protein arginine methylation and cardio‐ vascular disease, we must add ADMA, an additional player that may impact the cardiovas‐

As illustrated in Figure 1, the methyl group responsible for the establishment of protein ar‐ ginine methylation patterns by PRMTs originates from AdoMet, an intermediate in the ho‐ mocysteine metabolism. Therefore, protein arginine methylation and homocysteine metabolism are biochemically linked (Figure 4). In addition to protein arginine methylation, AdoMet serves as the methyl donor for more than one hundred cellular methyl transfer re‐ actions, including the methylation of other residues in proteins (namely, histidine and ly‐ sine), DNA and RNA. Following the transfer of the methyl group, AdoMet is converted into S-adenosyl homocysteine (AdoHcy). AdoHcy is further converted into homocysteine and adenosine by AdoHcy hydrolase, which is widely distributed in mammalian tissues [125]. This reaction is reversible and strongly favors AdoHcy synthesis rather than its hydrolysis; however, both homocysteine and adenosine are rapidly removed under physiological condi‐

new layer to the regulation of protein arginine methylation [118, 119].

102 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**7. Protein arginine hypomethylation in the context of hyperhomocysteinemia: "Another brick in the wall"?**

**disease**

cular system.

**Figure 4.** Simplified homocysteine metabolism. Homocysteine (Hcy) results from methionine metabolism as a byproduct of cellular transmethylation reactions. Methionine (Met) is the "entry" point of this pathway and is provided by diet and protein breakdown. S-Adenosylmethionine (AdoMet) is first synthesized from Met by methionine adeno‐ syltransferase (MAT). AdoMet is the methyl group donor in a wide variety of transmethylation reactions, being con‐ verted to S-Adenosylhomocysteine (AdoHcy). AdoHcy is subsequently hydrolyzed to Hcy and adenosine by AdoHcy hydrolase. Hcy can then be converted to cysteine (Cys) through the transsulphuration pathway, remethylated back to methionine or exported.

Despite recent controversy, elevated levels of homocysteine in plasma (or hyperhomocystei‐ nemia) are accepted as a risk factor for cardiovascular disease [127, 128]. Notably, despite several potential mechanisms underlying this association having been the focus of intense research, the subject remains to be fully understood [125]. Our group has been actively studying whether a hypomethylating environment associated with hyperhomocysteinemia contributes to the vascular toxicity of homocysteine [129-133]. AdoHcy is a competitive in‐ hibitor of most AdoMet-dependent methyltransferases, since it binds to their active sites with higher affinity than AdoMet [134]. As such, the ratio AdoMet/AdoHcy is taken as an index of cellular methylation capacity. As mentioned above, if homocysteine accumulates, AdoHcy will accumulate as well. Thus, increased homocysteine may be regarded as a cellu‐ lar hypomethylation effector, via AdoHcy accumulation.

We have demonstrated that patients with vascular disease present increased levels of both plasma homocysteine and intracellular AdoHcy, together with decreased DNA methylation [129]. Other studies also supported a role for hyperhomocysteinemia in modulating epige‐ netic mechanisms [135-138]. Additionally, we confirmed, *in vitro*, the AdoHcy ability to de‐ crease global DNA methylation status [131]. Currently, we are studying whether, in addition to DNA, protein arginine methylation extend is are also decreased by AdoHcy ac‐ cumulation. Supporting this possibility, results from kinetic studies regarding competitive inhibition of several methyltransferases by AdoHcy sustain that this metabolite is a stronger inhibitor of PRMT1 than of DNA methyltransferase 1, the enzyme responsible for the main‐ tenance of DNA methylation patterns [6, 139]. Interestingly, we observed that intracellular AdoHcy accumulation lowered ADMA production by human vascular endothelial cells ac‐ cumulation [133]. This observation led us to postulate that this effect was due to AdoHcyinduced protein arginine hypomethylation. Supporting this possibility, we subsequently found decreased levels of protein-incorporated ADMA in human vascular endothelial cells under AdoHcy intracellular accumulation ([140] and Esse R et *al*., manuscript 1 under sub‐ mission). Moreover, we observed that this effect was more pronounced than the parallel re‐ duction in global DNA methylation patterns. Therefore, we have concluded that AdoHcy build-up affects protein arginine methylation to a higher extent than DNA methylation. Im‐ portantly, this observation suggests that protein arginine methylation is more easily affected than DNA methylation in the context of hyperhomocysteinemia. Very recently, we have also assessed protein arginine methylation status in an animal model of diet-induced hyperho‐ mocysteinemia and found that increased homocysteine is associated with protein arginine hypomethylation, although in a tissue-dependent manner ([141] and Esse R *et al*., manu‐ script 2 under submission). Specifically, we found global protein arginine hypomethylation in heart and in brain from hyperhomocysteinemic rats, while methylation of hepatic pro‐ teins was not affected. Currently, we are undertaking both *in vitro* and *in vivo* studies to identify proteins sensitive to AdoHcy-induced hypomethylation, and to assess the impact of its different methylation states on vascular homeostasis. Actually, differential methylation of proteins may well play a role in pathologies related to elevated homocysteine, including vascular disease.

now recognized as a reversible and dynamic modification, increasing the complexity of the interplay between DNA, proteins and vital cellular process. Nevertheless, the characterization of enzymes that reverse and block protein arginine methylation has just commenced. Moreover, the pathological implications of protein arginine methylation disturbance have not been exhaustively explored. The methyl group marking of arginine residues originates from AdoMet, an intermediate in homocysteine metabolism. Thus, the metabolism of this amino acid and protein arginine methylation are biochemically linked. Homocysteine elevation relates with an increased risk of vascular disease and others pathologies. Notably, the potential detrimental effect of increased circulating homocysteine on protein arginine methylation status may constitute an additional molecular mechanism contributing to disease, and, as such, warrants further

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This work was supported by Fundação para a Ciência e a Tecnologia (FCT, Portugal) grants

, Paula Leandro1,2, Isabel Rivera1,2, Isabel Tavares de Almeida1,2,

1 Institute for Medicines and Pharmaceutical Sciences (iMed.UL), Faculty of Pharmacy, Uni‐

2 Department of Biochemistry and Human Biology, Faculty of Pharmacy, University of Lis‐

3 Metabolic Unit, Department of Clinical Chemistry, VU University Medical Center, Am‐

4 Institute for Cardiovascular Research ICaR-VU, VU University Medical Center, Amster‐

[1] Human Genome Sequencing Consortium. Finishing the euchromatic sequence of the

(SFRH/BD/48585/2008 to RE, and PTDC/SAU-ORG/112683/2009 to RC).

investigation.

**Acknowledgments**

**Author details**

bon, Lisbon, Portugal

sterdam,, The Netherlands

dam, The Netherlands

**References**

Henk J Blom3,4 and Rita Castro1,2\*

versity of Lisbon, Lisbon, Portugal

\*Address all correspondence to: rcastro@ff.ul.pt

human genome. Nature. 2004;431:931-945.

Ruben Esse1

#### **8. Conclusions**

The field of protein arginine methylation has been enjoying widespread interest in the scientific community, owing to the recent development of molecular biology techniques and mass spectrometry instrumentation. It is now clear that this common PTM impacts crucial biological functions, including RNA processing, transcriptional regulation, DNA repair and signal transduction. Yet, information is still scant regarding some important issues of protein arginine methylation, namely its regulation by other PTMs, how it affects interaction with the protein ligands, and what are the factors governing the specificity of the different PRMTs towards the target protein. Importantly, protein arginine methylation, which was considered a static modification during a long time, is now recognized as a reversible and dynamic modification, increasing the complexity of the interplay between DNA, proteins and vital cellular process. Nevertheless, the characterization of enzymes that reverse and block protein arginine methylation has just commenced. Moreover, the pathological implications of protein arginine methylation disturbance have not been exhaustively explored. The methyl group marking of arginine residues originates from AdoMet, an intermediate in homocysteine metabolism. Thus, the metabolism of this amino acid and protein arginine methylation are biochemically linked. Homocysteine elevation relates with an increased risk of vascular disease and others pathologies. Notably, the potential detrimental effect of increased circulating homocysteine on protein arginine methylation status may constitute an additional molecular mechanism contributing to disease, and, as such, warrants further investigation.

#### **Acknowledgments**

We have demonstrated that patients with vascular disease present increased levels of both plasma homocysteine and intracellular AdoHcy, together with decreased DNA methylation [129]. Other studies also supported a role for hyperhomocysteinemia in modulating epige‐ netic mechanisms [135-138]. Additionally, we confirmed, *in vitro*, the AdoHcy ability to de‐ crease global DNA methylation status [131]. Currently, we are studying whether, in addition to DNA, protein arginine methylation extend is are also decreased by AdoHcy ac‐ cumulation. Supporting this possibility, results from kinetic studies regarding competitive inhibition of several methyltransferases by AdoHcy sustain that this metabolite is a stronger inhibitor of PRMT1 than of DNA methyltransferase 1, the enzyme responsible for the main‐ tenance of DNA methylation patterns [6, 139]. Interestingly, we observed that intracellular AdoHcy accumulation lowered ADMA production by human vascular endothelial cells ac‐ cumulation [133]. This observation led us to postulate that this effect was due to AdoHcyinduced protein arginine hypomethylation. Supporting this possibility, we subsequently found decreased levels of protein-incorporated ADMA in human vascular endothelial cells under AdoHcy intracellular accumulation ([140] and Esse R et *al*., manuscript 1 under sub‐ mission). Moreover, we observed that this effect was more pronounced than the parallel re‐ duction in global DNA methylation patterns. Therefore, we have concluded that AdoHcy build-up affects protein arginine methylation to a higher extent than DNA methylation. Im‐ portantly, this observation suggests that protein arginine methylation is more easily affected than DNA methylation in the context of hyperhomocysteinemia. Very recently, we have also assessed protein arginine methylation status in an animal model of diet-induced hyperho‐ mocysteinemia and found that increased homocysteine is associated with protein arginine hypomethylation, although in a tissue-dependent manner ([141] and Esse R *et al*., manu‐ script 2 under submission). Specifically, we found global protein arginine hypomethylation in heart and in brain from hyperhomocysteinemic rats, while methylation of hepatic pro‐ teins was not affected. Currently, we are undertaking both *in vitro* and *in vivo* studies to identify proteins sensitive to AdoHcy-induced hypomethylation, and to assess the impact of its different methylation states on vascular homeostasis. Actually, differential methylation of proteins may well play a role in pathologies related to elevated homocysteine, including

104 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

The field of protein arginine methylation has been enjoying widespread interest in the scientific community, owing to the recent development of molecular biology techniques and mass spectrometry instrumentation. It is now clear that this common PTM impacts crucial biological functions, including RNA processing, transcriptional regulation, DNA repair and signal transduction. Yet, information is still scant regarding some important issues of protein arginine methylation, namely its regulation by other PTMs, how it affects interaction with the protein ligands, and what are the factors governing the specificity of the different PRMTs towards the target protein. Importantly, protein arginine methylation, which was considered a static modification during a long time, is

vascular disease.

**8. Conclusions**

This work was supported by Fundação para a Ciência e a Tecnologia (FCT, Portugal) grants (SFRH/BD/48585/2008 to RE, and PTDC/SAU-ORG/112683/2009 to RC).

#### **Author details**

Ruben Esse1 , Paula Leandro1,2, Isabel Rivera1,2, Isabel Tavares de Almeida1,2, Henk J Blom3,4 and Rita Castro1,2\*

\*Address all correspondence to: rcastro@ff.ul.pt

1 Institute for Medicines and Pharmaceutical Sciences (iMed.UL), Faculty of Pharmacy, Uni‐ versity of Lisbon, Lisbon, Portugal

2 Department of Biochemistry and Human Biology, Faculty of Pharmacy, University of Lis‐ bon, Lisbon, Portugal

3 Metabolic Unit, Department of Clinical Chemistry, VU University Medical Center, Am‐ sterdam,, The Netherlands

4 Institute for Cardiovascular Research ICaR-VU, VU University Medical Center, Amster‐ dam, The Netherlands

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Deciphering Protein Arginine Methylation in Mammals

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**Section 4**

**Cancer Research Through Study of Methylation**

**Cell Processes**


**Cancer Research Through Study of Methylation Cell Processes**

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116 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Chapter 5**

**Methylation in Tumorigenesis**

Melissa A. Edwards, Pashayar P. Lookian,

Additional information is available at the end of the chapter

tumorigenesis and subsequent clinical outcomes [17-23].

The development, maturation, and maintenance of tissues and organisms are anchored in distinct programs for protein expression which define the identities and roles of individual cell lines [1, 2]. These programs are maintained in a heritable state by epigenetic mecha‐ nisms that convey cellular memory [3, 4]. In this way, the global synchronization of patterns in gene expression broadly dictates developmental consequences [5, 6]. At the foundation of such gene regulation are coordinated cascades that affect the packaging of DNA into chro‐ matin, thereby establishing the degree of DNA accessibility to transcriptional complexes [7-10]. These pathways include histone methylation, methylation of transcriptional regula‐ tors, DNA methylation, histone replacement, chromatin remodeling, and other alterations to histone tails [11-16]. Abnormalities in these epigenetic events are commonly associated with

From tightly regulated transcription to mitosis, chromatin is an elastic repository of the ge‐ nome [24]. In this state, a chromosome is sequentially condensed through a succession of or‐ ganized compaction while limited regions of DNA are selectively made available to transcriptional complexes [25, 26]. Hence, chromatin exists in a dynamic state into which ap‐ proximately 2 m of DNA is packaged in the nucleus while maintaining an extraordinary lev‐ el of utility [25, 26]. At its core, chromatin is established in a series of nucleosomes, their basic structural unit [25, 27], comprised of 146 base pairs of DNA, wrapped 1.7 times around an octamer of histones and interspersed by regions of roughly 50 base pairs [28]. The key histones participating in the assembly of a nucleosome include histones H2A, H2B, H3 and H4. These histones form hetero-dimers resulting in each being twice represented in the nu‐ cleosomal unit [29-31]. Structurally, histones are are highly conserved, including a folded core followed by an unstructured tail [30, 31]. A globular domain forms the histone core as a

> © 2013 Edwards et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

Drew R. Neavin and Mark A. Brown

http://dx.doi.org/10.5772/52867

**1. Introduction**

#### **Chapter 5**

### **Methylation in Tumorigenesis**

Melissa A. Edwards, Pashayar P. Lookian, Drew R. Neavin and Mark A. Brown

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52867

#### **1. Introduction**

The development, maturation, and maintenance of tissues and organisms are anchored in distinct programs for protein expression which define the identities and roles of individual cell lines [1, 2]. These programs are maintained in a heritable state by epigenetic mecha‐ nisms that convey cellular memory [3, 4]. In this way, the global synchronization of patterns in gene expression broadly dictates developmental consequences [5, 6]. At the foundation of such gene regulation are coordinated cascades that affect the packaging of DNA into chro‐ matin, thereby establishing the degree of DNA accessibility to transcriptional complexes [7-10]. These pathways include histone methylation, methylation of transcriptional regula‐ tors, DNA methylation, histone replacement, chromatin remodeling, and other alterations to histone tails [11-16]. Abnormalities in these epigenetic events are commonly associated with tumorigenesis and subsequent clinical outcomes [17-23].

From tightly regulated transcription to mitosis, chromatin is an elastic repository of the ge‐ nome [24]. In this state, a chromosome is sequentially condensed through a succession of or‐ ganized compaction while limited regions of DNA are selectively made available to transcriptional complexes [25, 26]. Hence, chromatin exists in a dynamic state into which ap‐ proximately 2 m of DNA is packaged in the nucleus while maintaining an extraordinary lev‐ el of utility [25, 26]. At its core, chromatin is established in a series of nucleosomes, their basic structural unit [25, 27], comprised of 146 base pairs of DNA, wrapped 1.7 times around an octamer of histones and interspersed by regions of roughly 50 base pairs [28]. The key histones participating in the assembly of a nucleosome include histones H2A, H2B, H3 and H4. These histones form hetero-dimers resulting in each being twice represented in the nu‐ cleosomal unit [29-31]. Structurally, histones are are highly conserved, including a folded core followed by an unstructured tail [30, 31]. A globular domain forms the histone core as a

© 2013 Edwards et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

helix-turn-helix motif, which allows dimerization [31]. In contrast, the tails of histones do not exist in defined conformations except when attached to their cognate proteins [31]. Within the sequence of histone tail domains is a large representation of conserved amino acid residues including lysine, arginine, and serine [31, 32]. Under normal conditions, his‐ tone tails have a net basic charge facilitating their interaction with the poly-anionic back‐ bone of DNA, thereby contributing to the stability of nucleosomes [31]. Consequently, chromatin structure and transcriptional regulation are commonly mediated through posttranslational modifications that impact specific residues within the sequence of these tails [33, 34]. Modifications to tail residues can regulate the accessibility of nuclear factors to re‐ gions of DNA or induce the recruitment of such factors involved in chromatin structural and transcriptional regulatory pathways [33, 34].

ble [60]. Of particular importance in the role of chromatin modifications in human disease

Methylation in Tumorigenesis http://dx.doi.org/10.5772/52867 121

Methylation of DNA is a covalent modification that occurs at cytosines within CpG-rich re‐ gions of DNA and is catalyzed by DNA methyltransferases [61, 62]. The methylation of DNA affects the binding of proteins to their cognate DNA sequences [61, 63]. Such addition of methyl groups can prevent the binding of basal transcriptional machinery and ubiquitous transcription factors [61]. Thus, DNA methylation contributes to epigenetic inheritance, al‐ lele-specific expression, inactivation of the X chromosome, genomic stability and embryonic development. It is through these pathways that progressive DNA methylation is thought to be an agent for both normal aging as well as neoplasias [64, 65]. The majority of methylated CpG islands are located within repetitive elements including centromeric repeats, satellite sequences and gene repeats. These CpG regions are often found at the 5' end of genes where DNA methylation affects transcription by recruiting methyl-CpG binding domain (MBD) proteins that function as adaptors between methylated DNA and chromatin-modifying en‐ zymes [66]. There is a clear relationship between DNA methylation and other silencing mechanisms including histone modifications and chromatin remodeling [65, 66]. In fact, sev‐ eral studies suggest that DNA methylation affects genes that are already suppressed by oth‐

Changes in the pattern of DNA methylation have been correlated with altered histone posttranslational modifications and genetic lesions [67]. Either hypermethylation or hypomethy‐ lation have been identified in almost all types of cancer cells examined, to date [18, 21, 68]. Hypomethylation at centromeric repeat sequences has been linked to genomic instability [18] whereas local hypermethylation of individual genes has been associated with aberrant gene silencing [21]. In oncogenic cells, hypermethylation is often correlated with the repres‐ sion of tumor suppressor genes while hypomethylation is associated with the activation of genes required for invasion and metastasis [68, 69]. In neoplastic tissues, the incidence of hy‐ permethylation in genes with promoter associated CpG islands in markedly increased which, in turn, is associated with repression of tumor suppressors [70]. Although the com‐ plete mechanistic pathway for DNA methylation in cancers is still being determined, aber‐ rant methylation in tumors is already being examined as an instrument for diagnosis [21, 70]. For example, techniques, such as the polymerase chain reaction amplification of bisul‐ fite-modified DNA, have enabled the study of patterns of DNA methylation [71-73]. These methods are currently being improved and adapted for cancer cell identification, profiling of tumor-suppressor-gene expression, and prognostic factors that are linked to CpG island hypermethylation [74]. Likewise, reversal of hypermethylation by several indiscriminant de‐ methylating compounds has been approved for therapeutic intervention associated with

Detection of DNA methylation has recently been added to the armory of preventative/diag‐ nostic medicine for the prevention and treatment of colorectal cancer [75]. This represents

are methylation of DNA, histone targets, and other regulatory targets.

**2. DNA methylation in tumorigenesis**

er mechanisms [65].

blood-borne cancers [21].

The Histone-DNA interface is formed principally by inelastic hydrogen bonds between the phosphate oxygen of DNA and the main chain amide of the histone. Electrostatic interac‐ tions between basic side chains and negatively charged phosphate groups and other nonpo‐ lar interactions further strengthen the association between histones and DNA [35]. While this, in theory, should facilitate the establishment of nucleosomes upon any DNA sequence, there are likely specific sequence parameters for nucleosomal placement [36]. The composi‐ tion of the DNA sequences, by which the histone core is enveloped, is likely a major factor contributing to the positioning of core histones and the dynamic comportment of the nucleo‐ some under the influence of the SWI/SNF ATPase and sequence-specific transcription fac‐ tors [37]. The most broadly characterized nucleosomal assembly is the 30 nm fiber [38], which is anchored by linker histones [39-41] and the relative juxtaposition of each nucleo‐ some [42], establishing close physical proximity while generating only marginal internucleo‐ somal attraction energy [38, 43-45]. Hence, this architecture allows a great degree of variation in condensation without producing serious topological changes. Chromatin exists in a series of more densely compacted structures [46], which are commonly driven by inter‐ action with non-histone, structural proteins [47].

In recent decades, a number of process which impact the structure and/or function of chro‐ matin including post-translational modifications of histones, DNA methylation, incorpora‐ tion of histone variants, and ATP-dependent chromatin remodelling have been the subjects of intense study. The findings of these studies clearly show that chromatin modifications and the complexes involved with their facilitation are linked to the control of many biologi‐ cal processes which depend upon the level of chromatin accessibility [48-51]. Such processes include chromosome segregation during mitosis, X chromosome inactivation, gene expres‐ sion, DNA repair, and chromatin condensation during apoptosis [23, 52-56].

Chromatin modifications convey epigenetic regulation of protein expression without altera‐ tions in DNA sequence. Disturbing the equilibrium of epigenetic networks has been shown to be associated with numerous pathological events, including syndromes involving chro‐ mosomal instability, neurological disorders, and tumorigenesis [57-59]. Advances in knowl‐ edge related to epigenetic inheritance and chromatin structure/regulation have paved the way for promising novel therapeutics directed against the specific factors that are responsi‐ ble [60]. Of particular importance in the role of chromatin modifications in human disease are methylation of DNA, histone targets, and other regulatory targets.

#### **2. DNA methylation in tumorigenesis**

helix-turn-helix motif, which allows dimerization [31]. In contrast, the tails of histones do not exist in defined conformations except when attached to their cognate proteins [31]. Within the sequence of histone tail domains is a large representation of conserved amino acid residues including lysine, arginine, and serine [31, 32]. Under normal conditions, his‐ tone tails have a net basic charge facilitating their interaction with the poly-anionic back‐ bone of DNA, thereby contributing to the stability of nucleosomes [31]. Consequently, chromatin structure and transcriptional regulation are commonly mediated through posttranslational modifications that impact specific residues within the sequence of these tails [33, 34]. Modifications to tail residues can regulate the accessibility of nuclear factors to re‐ gions of DNA or induce the recruitment of such factors involved in chromatin structural

The Histone-DNA interface is formed principally by inelastic hydrogen bonds between the phosphate oxygen of DNA and the main chain amide of the histone. Electrostatic interac‐ tions between basic side chains and negatively charged phosphate groups and other nonpo‐ lar interactions further strengthen the association between histones and DNA [35]. While this, in theory, should facilitate the establishment of nucleosomes upon any DNA sequence, there are likely specific sequence parameters for nucleosomal placement [36]. The composi‐ tion of the DNA sequences, by which the histone core is enveloped, is likely a major factor contributing to the positioning of core histones and the dynamic comportment of the nucleo‐ some under the influence of the SWI/SNF ATPase and sequence-specific transcription fac‐ tors [37]. The most broadly characterized nucleosomal assembly is the 30 nm fiber [38], which is anchored by linker histones [39-41] and the relative juxtaposition of each nucleo‐ some [42], establishing close physical proximity while generating only marginal internucleo‐ somal attraction energy [38, 43-45]. Hence, this architecture allows a great degree of variation in condensation without producing serious topological changes. Chromatin exists in a series of more densely compacted structures [46], which are commonly driven by inter‐

In recent decades, a number of process which impact the structure and/or function of chro‐ matin including post-translational modifications of histones, DNA methylation, incorpora‐ tion of histone variants, and ATP-dependent chromatin remodelling have been the subjects of intense study. The findings of these studies clearly show that chromatin modifications and the complexes involved with their facilitation are linked to the control of many biologi‐ cal processes which depend upon the level of chromatin accessibility [48-51]. Such processes include chromosome segregation during mitosis, X chromosome inactivation, gene expres‐

Chromatin modifications convey epigenetic regulation of protein expression without altera‐ tions in DNA sequence. Disturbing the equilibrium of epigenetic networks has been shown to be associated with numerous pathological events, including syndromes involving chro‐ mosomal instability, neurological disorders, and tumorigenesis [57-59]. Advances in knowl‐ edge related to epigenetic inheritance and chromatin structure/regulation have paved the way for promising novel therapeutics directed against the specific factors that are responsi‐

sion, DNA repair, and chromatin condensation during apoptosis [23, 52-56].

and transcriptional regulatory pathways [33, 34].

120 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

action with non-histone, structural proteins [47].

Methylation of DNA is a covalent modification that occurs at cytosines within CpG-rich re‐ gions of DNA and is catalyzed by DNA methyltransferases [61, 62]. The methylation of DNA affects the binding of proteins to their cognate DNA sequences [61, 63]. Such addition of methyl groups can prevent the binding of basal transcriptional machinery and ubiquitous transcription factors [61]. Thus, DNA methylation contributes to epigenetic inheritance, al‐ lele-specific expression, inactivation of the X chromosome, genomic stability and embryonic development. It is through these pathways that progressive DNA methylation is thought to be an agent for both normal aging as well as neoplasias [64, 65]. The majority of methylated CpG islands are located within repetitive elements including centromeric repeats, satellite sequences and gene repeats. These CpG regions are often found at the 5' end of genes where DNA methylation affects transcription by recruiting methyl-CpG binding domain (MBD) proteins that function as adaptors between methylated DNA and chromatin-modifying en‐ zymes [66]. There is a clear relationship between DNA methylation and other silencing mechanisms including histone modifications and chromatin remodeling [65, 66]. In fact, sev‐ eral studies suggest that DNA methylation affects genes that are already suppressed by oth‐ er mechanisms [65].

Changes in the pattern of DNA methylation have been correlated with altered histone posttranslational modifications and genetic lesions [67]. Either hypermethylation or hypomethy‐ lation have been identified in almost all types of cancer cells examined, to date [18, 21, 68]. Hypomethylation at centromeric repeat sequences has been linked to genomic instability [18] whereas local hypermethylation of individual genes has been associated with aberrant gene silencing [21]. In oncogenic cells, hypermethylation is often correlated with the repres‐ sion of tumor suppressor genes while hypomethylation is associated with the activation of genes required for invasion and metastasis [68, 69]. In neoplastic tissues, the incidence of hy‐ permethylation in genes with promoter associated CpG islands in markedly increased which, in turn, is associated with repression of tumor suppressors [70]. Although the com‐ plete mechanistic pathway for DNA methylation in cancers is still being determined, aber‐ rant methylation in tumors is already being examined as an instrument for diagnosis [21, 70]. For example, techniques, such as the polymerase chain reaction amplification of bisul‐ fite-modified DNA, have enabled the study of patterns of DNA methylation [71-73]. These methods are currently being improved and adapted for cancer cell identification, profiling of tumor-suppressor-gene expression, and prognostic factors that are linked to CpG island hypermethylation [74]. Likewise, reversal of hypermethylation by several indiscriminant de‐ methylating compounds has been approved for therapeutic intervention associated with blood-borne cancers [21].

Detection of DNA methylation has recently been added to the armory of preventative/diag‐ nostic medicine for the prevention and treatment of colorectal cancer [75]. This represents the most common form of gastrointestinal cancer and is a leading killer among all malignan‐ cies. The colonoscopy is broadly employed for detection of lesions which often give rise to colorectal tumors. The invasiveness of this procedure and consequent lack of patient cooper‐ ation with recommended colonoscopies is a limiting factor in the efficacy of this procedure as a preventative. Fortunately, epigenetic screening has arisen as a new tier in preventative medicine targeting colorectal cancer. Specifically, DNA methylation is associated with genesilencing related to onset of colorectal cancer. Given that these changes are detectable prior to tumorigenesis, target-specific screening of DNA methylation represents a promising front in the war against colorectal cancer [75].

gree of achievable specificity is increased by the breadth of biological outcomes which are dependent upon whether a residue is mono-, di-, or tri-methylated [93-95]. Likewise, is has also been widely observed that histone lysine methylation works concomitantly with many transient histone modifications, thereby further enhancing the degree of information which

Methylation in Tumorigenesis http://dx.doi.org/10.5772/52867 123

Almost all histone lysine methyltransferases are dependent upon a SET domain for catalyz‐ ing the transfer of methyl groups. The SET domain is present in many proteins that control a range of biological processes, including several involved in development and proper cell cy‐ cle progression [7, 96]. Promoter associated, residue-specific histone methylation often cor‐ relates with distinct patterns of protein expression [96]. The bulk of known modifications occur on histone H3 which thereby serves as a central conduit of epigenetic regulation. Ly‐ sine methylation at histone H3, lysine 9 (H3K9), H3K27, and H4K20 is most often associated with gene silencing, whereas methylation of H3K4, H3K36, or H3K79 is commonly linked to the activation of transcription [7, 96]. Accumulating evidence points to histone methylation in the recruitment of chromatin remodeling complexes, such as CHD1, an ATP-dependent chromatin remodeling factor that binds specifically to methylated the forms of H3K4 [97]. While histone lysine methylation was previously believed to be a permanent mark, a num‐

can be communicated by this epigenetic modification [15].

**Figure 1.** Common sites of Histone Methylation

Similar to colorectal cancer, the role of DNA methylation in prostate cancers has been the subject of numerous studies [76-78]. Indeed, aberrant hyper/hypo methylation has been linked in numerous prostate-specific malignant processes ranging from early tumorigenesis to late stage, androgen independent tumors [79]. The identification of specific targets which are down-regulated as a result of hypermethylation at their promoters has lead to the devel‐ opment of methylation biomarkers for early detection [80]. These targets of inactivation by promoter hypermethylation include Ras-association domain family 1A (RASSF1A), GSTP1, and retinoic acid receptor beta2 (RARbeta2). Though the role of hypomethylation in prostate cancer is less understood, there are several recent studies which link it to alterations in the expression of genes associated with early and late stage prostate tumors [81-83].

Additional studies indicate that hyper/hypomethylation of specific promoter regions is asso‐ ciated with tumorigenesis in a broad range of tissues including lung, breast, thyroid, head and neck squamous cell carcinomas, and hepatocellular carcinomas [84-86]. There are ongo‐ ing studies in dozens of other tissue types indicating a role for hyper/hypomethylation in a broadening range of cancers. Thus, aberrant alterations in methylation promise to provide a broad-spectrum mechanism for early detection and prognostication of tumors.

#### **3. Histone methylation in tumorigenesis**

Modifications to histone tails comprise the broadest range of variation in epigenetic controls, encompassing more than four dozen known sites of alteration [87]. Histone proteins are the targets of many forms of post-translational modification such as citrullination, acetylation, phosphorylation, SUMOylation, ADP-ribosylation and methylation (Figure 1) [87, 88]. These alterations are translated into biological consequences by impacting the structure of the nu‐ cleosome as well as facilitating the recruitment of specific regulatory complexes. The combi‐ nation of different histone modifications in concert communicates a "histone code" that is interpreted in the form of distinct nuclear events [89].

Histone methylation is has been observed to be a mark that imparts long-term epigenetic memory [90]. Histone lysine methylation is a central factor in such processes as X chromo‐ some inactivation, DNA methylation, transcriptional regulation, and the formation of heter‐ ochromatin [91, 92]. This modification, catalyzed by histone methyltransferases, often facilitates the regulation of protein expression in a residue-dependent manner [90]. The de‐ gree of achievable specificity is increased by the breadth of biological outcomes which are dependent upon whether a residue is mono-, di-, or tri-methylated [93-95]. Likewise, is has also been widely observed that histone lysine methylation works concomitantly with many transient histone modifications, thereby further enhancing the degree of information which can be communicated by this epigenetic modification [15].

**Figure 1.** Common sites of Histone Methylation

the most common form of gastrointestinal cancer and is a leading killer among all malignan‐ cies. The colonoscopy is broadly employed for detection of lesions which often give rise to colorectal tumors. The invasiveness of this procedure and consequent lack of patient cooper‐ ation with recommended colonoscopies is a limiting factor in the efficacy of this procedure as a preventative. Fortunately, epigenetic screening has arisen as a new tier in preventative medicine targeting colorectal cancer. Specifically, DNA methylation is associated with genesilencing related to onset of colorectal cancer. Given that these changes are detectable prior to tumorigenesis, target-specific screening of DNA methylation represents a promising front

Similar to colorectal cancer, the role of DNA methylation in prostate cancers has been the subject of numerous studies [76-78]. Indeed, aberrant hyper/hypo methylation has been linked in numerous prostate-specific malignant processes ranging from early tumorigenesis to late stage, androgen independent tumors [79]. The identification of specific targets which are down-regulated as a result of hypermethylation at their promoters has lead to the devel‐ opment of methylation biomarkers for early detection [80]. These targets of inactivation by promoter hypermethylation include Ras-association domain family 1A (RASSF1A), GSTP1, and retinoic acid receptor beta2 (RARbeta2). Though the role of hypomethylation in prostate cancer is less understood, there are several recent studies which link it to alterations in the

Additional studies indicate that hyper/hypomethylation of specific promoter regions is asso‐ ciated with tumorigenesis in a broad range of tissues including lung, breast, thyroid, head and neck squamous cell carcinomas, and hepatocellular carcinomas [84-86]. There are ongo‐ ing studies in dozens of other tissue types indicating a role for hyper/hypomethylation in a broadening range of cancers. Thus, aberrant alterations in methylation promise to provide a

Modifications to histone tails comprise the broadest range of variation in epigenetic controls, encompassing more than four dozen known sites of alteration [87]. Histone proteins are the targets of many forms of post-translational modification such as citrullination, acetylation, phosphorylation, SUMOylation, ADP-ribosylation and methylation (Figure 1) [87, 88]. These alterations are translated into biological consequences by impacting the structure of the nu‐ cleosome as well as facilitating the recruitment of specific regulatory complexes. The combi‐ nation of different histone modifications in concert communicates a "histone code" that is

Histone methylation is has been observed to be a mark that imparts long-term epigenetic memory [90]. Histone lysine methylation is a central factor in such processes as X chromo‐ some inactivation, DNA methylation, transcriptional regulation, and the formation of heter‐ ochromatin [91, 92]. This modification, catalyzed by histone methyltransferases, often facilitates the regulation of protein expression in a residue-dependent manner [90]. The de‐

expression of genes associated with early and late stage prostate tumors [81-83].

broad-spectrum mechanism for early detection and prognostication of tumors.

**3. Histone methylation in tumorigenesis**

interpreted in the form of distinct nuclear events [89].

in the war against colorectal cancer [75].

122 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Almost all histone lysine methyltransferases are dependent upon a SET domain for catalyz‐ ing the transfer of methyl groups. The SET domain is present in many proteins that control a range of biological processes, including several involved in development and proper cell cy‐ cle progression [7, 96]. Promoter associated, residue-specific histone methylation often cor‐ relates with distinct patterns of protein expression [96]. The bulk of known modifications occur on histone H3 which thereby serves as a central conduit of epigenetic regulation. Ly‐ sine methylation at histone H3, lysine 9 (H3K9), H3K27, and H4K20 is most often associated with gene silencing, whereas methylation of H3K4, H3K36, or H3K79 is commonly linked to the activation of transcription [7, 96]. Accumulating evidence points to histone methylation in the recruitment of chromatin remodeling complexes, such as CHD1, an ATP-dependent chromatin remodeling factor that binds specifically to methylated the forms of H3K4 [97]. While histone lysine methylation was previously believed to be a permanent mark, a num‐ ber of enzymes have now been identified that are capable of reversing histone methylation in a site-specific manner [98-100].

H3K4 [114, 115], has also been linked to the regulation of p53 by its capacity to methylate that protein at lysine 372 [116, 117]. Methylation of that site stabilizes p53 and limits its lo‐

Methylation in Tumorigenesis http://dx.doi.org/10.5772/52867 125

The broad roles of aberrant lysine methylation in the induction of carcinogenesis have paved the way for a novel line of cancer therapeutics. [118]. Those therapeutics bank on the potential to manipulate of the demethylating activity of a host of demethylases. The fact that many demethylases target highly specific substrates heightens their potential utility as high‐

While it is true that chromatin architecture has the capacity to be epigenetically maintained and inherited via modified states of methylation, recent studies have highlighted the fact that methylation-induced control of gene expression may be altered by environmental stres‐ sors and toxicants. Such modifications may, in consequence, induce aberrations toward ge‐ nome integrity and stability. Distinct from genetic mutations, these epigenetic aberrations have been termed epimutations. In contrast to genetic mutations, which may be passively inherited, epimutations require active maintenance [119]. That epimutations rare appear‐ ance in normal tissues highlights the potential for epigenetic therapies based on high tumor specificity. Likewise, while therapeutics based on genetic deletions commonly induce an ir‐ reversible loss of gene function, epigenetic alterations are reversible, further enhancing their potential utility for therapeutic intervention [120]. Reversal of epimutations to restore nor‐ mal expression of tumor suppressors has become the holy grail in epigenetic cancer thera‐ peutics. Already, numerous studies have proven that aberrant gene silencing mediated by DNA methylation can be easily reversed by the incorporation of DNA methyltransferase in‐ hibitors [121]. Positive results have been observed after treating tumor cells with such phar‐

In the last two decades, our knowledge of chromatin methylation patterns and their role in the regulation of nuclear processes have been broadly elucidated. Understanding those pat‐ terns of histone changes, decoding the association between those alterations and DNA meth‐ ylation, and characterizing their relevance in tumorigenesis comprise the next hurdles in the

This work was supported by funding to Mark A. Brown from the National Science Founda‐

ly effective therapeutics with lower likelihood of instigating adverse effects.

calization to the nucleus [116].

**4. Conclusion**

macological agents [122, 123].

**Acknowledgements**

tion (1060548).

etiology of the role of methylation in cancer.

The presence of histone variants creates yet another tier to the potential of epigenetic mecha‐ nisms to communicate cellular information [53]. Variants affect the nucleosomal architecture as well as the proclivity of local chromatin to be remodeled. Thus, the inclusion of histone variants may modify nucleosome mobility, stability, and/or potential patterns of histone modifications. These, in turn, impact higher order structure and downstream events [101-104]. For example, a specialized H3-like variant CENP-A, replaces H3 in centromeric nucleosomes to establish a distinct architecture that is essential for proper segregation of the chromosomes [105]. In recent years, there have been an increasing number of experimental outcomes emphasizing the biological relevance of histone variants and their central role in epigenetic control [53].

Regulating the architecture of chromatin involves complex and dynamic mechanisms. The structure of chromatin is controlled on multiple levels by distinct processes such as nucleo‐ some remodeling, DNA methylation, histone post-translational modifications, incorporation of histone variants, and non-coding RNA. Aberrant activity within such epigenetic process‐ es is likely to broadly affect protein expression as well as other biological events such as apoptosis and condensation and segregation of chromosomes.

Tumorigenesis is a graded process through which a succession of genetic aberrations leads to the progressive transformation of healthy cells. While modifications in genetic sequence certainly account for many of these aberrations, an increasing number of modifications in gene expression observed in tumorigenesis have been found to be the result by epigenetic aberrations. These observations point to the relevance of epigenetic mechanisms in the maintenance of proper cell function. Aberrant events related to such mechanisms often act in concert with genetic mutations thereby contributing to the development and progression of cancer.

Over the last two decades, an increasing number of investigations have highlighted the aberrant gain or loss of histone methyltransferase activity in carcinogenesis. At one end of the spectrum, it has been shown that mice which fail to express the H3K9-specific histone methyltransferase, SUV39H1, are subject to increased incidence of chromosomal instability and subsequent tumorigenic potential [106]. At the opposite end of the spectrum, it is overexpression of Smyd3, another histone methyltransferase that is specific for H3K4, that has been shown to be responsible for unrestrained proliferation of many cancer cells [107]. A transcription factor binding element polymorphism in the upstream regulatory sequence for Smyd3 has been associated with an increased risk for cancer [108, 109].

In addition to histone targets, some SET domain-containing methyltransferases have been shown to methylate other proteins including tumor suppressors. Smyd2, which methylates H3K36 [6, 110], has also been directly linked to the regulation of p53 [111, 112]. Methylating lysine 370 of p53, Smyd2 has been shown to inhibit the transcriptional regulatory activity of p53. Smyd2 regulation of the retinoblastoma tumor suppressor (RB) has also been observed by its capacity to methylate of RB at K860 [113]. In a second example, Set9, which methylates

H3K4 [114, 115], has also been linked to the regulation of p53 by its capacity to methylate that protein at lysine 372 [116, 117]. Methylation of that site stabilizes p53 and limits its lo‐ calization to the nucleus [116].

The broad roles of aberrant lysine methylation in the induction of carcinogenesis have paved the way for a novel line of cancer therapeutics. [118]. Those therapeutics bank on the potential to manipulate of the demethylating activity of a host of demethylases. The fact that many demethylases target highly specific substrates heightens their potential utility as high‐ ly effective therapeutics with lower likelihood of instigating adverse effects.

#### **4. Conclusion**

ber of enzymes have now been identified that are capable of reversing histone methylation

The presence of histone variants creates yet another tier to the potential of epigenetic mecha‐ nisms to communicate cellular information [53]. Variants affect the nucleosomal architecture as well as the proclivity of local chromatin to be remodeled. Thus, the inclusion of histone variants may modify nucleosome mobility, stability, and/or potential patterns of histone modifications. These, in turn, impact higher order structure and downstream events [101-104]. For example, a specialized H3-like variant CENP-A, replaces H3 in centromeric nucleosomes to establish a distinct architecture that is essential for proper segregation of the chromosomes [105]. In recent years, there have been an increasing number of experimental outcomes emphasizing the biological relevance of histone variants and their central role in

Regulating the architecture of chromatin involves complex and dynamic mechanisms. The structure of chromatin is controlled on multiple levels by distinct processes such as nucleo‐ some remodeling, DNA methylation, histone post-translational modifications, incorporation of histone variants, and non-coding RNA. Aberrant activity within such epigenetic process‐ es is likely to broadly affect protein expression as well as other biological events such as

Tumorigenesis is a graded process through which a succession of genetic aberrations leads to the progressive transformation of healthy cells. While modifications in genetic sequence certainly account for many of these aberrations, an increasing number of modifications in gene expression observed in tumorigenesis have been found to be the result by epigenetic aberrations. These observations point to the relevance of epigenetic mechanisms in the maintenance of proper cell function. Aberrant events related to such mechanisms often act in concert with genetic mutations thereby contributing to the development and progression

Over the last two decades, an increasing number of investigations have highlighted the aberrant gain or loss of histone methyltransferase activity in carcinogenesis. At one end of the spectrum, it has been shown that mice which fail to express the H3K9-specific histone methyltransferase, SUV39H1, are subject to increased incidence of chromosomal instability and subsequent tumorigenic potential [106]. At the opposite end of the spectrum, it is overexpression of Smyd3, another histone methyltransferase that is specific for H3K4, that has been shown to be responsible for unrestrained proliferation of many cancer cells [107]. A transcription factor binding element polymorphism in the upstream regulatory sequence for

In addition to histone targets, some SET domain-containing methyltransferases have been shown to methylate other proteins including tumor suppressors. Smyd2, which methylates H3K36 [6, 110], has also been directly linked to the regulation of p53 [111, 112]. Methylating lysine 370 of p53, Smyd2 has been shown to inhibit the transcriptional regulatory activity of p53. Smyd2 regulation of the retinoblastoma tumor suppressor (RB) has also been observed by its capacity to methylate of RB at K860 [113]. In a second example, Set9, which methylates

apoptosis and condensation and segregation of chromosomes.

Smyd3 has been associated with an increased risk for cancer [108, 109].

in a site-specific manner [98-100].

124 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

epigenetic control [53].

of cancer.

While it is true that chromatin architecture has the capacity to be epigenetically maintained and inherited via modified states of methylation, recent studies have highlighted the fact that methylation-induced control of gene expression may be altered by environmental stres‐ sors and toxicants. Such modifications may, in consequence, induce aberrations toward ge‐ nome integrity and stability. Distinct from genetic mutations, these epigenetic aberrations have been termed epimutations. In contrast to genetic mutations, which may be passively inherited, epimutations require active maintenance [119]. That epimutations rare appear‐ ance in normal tissues highlights the potential for epigenetic therapies based on high tumor specificity. Likewise, while therapeutics based on genetic deletions commonly induce an ir‐ reversible loss of gene function, epigenetic alterations are reversible, further enhancing their potential utility for therapeutic intervention [120]. Reversal of epimutations to restore nor‐ mal expression of tumor suppressors has become the holy grail in epigenetic cancer thera‐ peutics. Already, numerous studies have proven that aberrant gene silencing mediated by DNA methylation can be easily reversed by the incorporation of DNA methyltransferase in‐ hibitors [121]. Positive results have been observed after treating tumor cells with such phar‐ macological agents [122, 123].

In the last two decades, our knowledge of chromatin methylation patterns and their role in the regulation of nuclear processes have been broadly elucidated. Understanding those pat‐ terns of histone changes, decoding the association between those alterations and DNA meth‐ ylation, and characterizing their relevance in tumorigenesis comprise the next hurdles in the etiology of the role of methylation in cancer.

#### **Acknowledgements**

This work was supported by funding to Mark A. Brown from the National Science Founda‐ tion (1060548).

#### **Author details**

Melissa A. Edwards1 , Pashayar P. Lookian2 , Drew R. Neavin3 and Mark A. Brown4\* [10] Imhof A: Epigenetic regulators and histone modification. Briefings in Functional Ge‐

Methylation in Tumorigenesis http://dx.doi.org/10.5772/52867 127

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1 Cell and Molecular Biology Program at Colorado State University, USA

2 Department of Biology at Colorado State University, USA

3 Department of Biology at Colorado State University, USA

4 Flint Cancer Center and Department of Clinical Sciences at Colorado State University, USA

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**Chapter 6**

**Circulating Methylated DNA as Biomarkers for Cancer**

In addition to genetic alterations including deletion or point mutations, epigenetic changes such as DNA methylation play an important role in silencing tumor suppressor genes dur‐ ing cancer development. By adding a methyl group from S-adenosyl-L-methionine to the cy‐ tosine pyrimidine or adenine purine ring, DNA methylation is important to maintain genome structure and regulate gene expression. In mammalian adult tissues, DNA methyla‐ tion occurs in CpG dinucleotides that often cluster in the genome as CpG islands in the 5' regulatory regions of the genes. Through recruiting transcriptional co-repressors including methyl-CpG-binding domain proteins (MBDs) and chromatin remodeling proteins like his‐ tone deacetylases (HDACs) or impeding the binding of transcriptional activators, DNA methylation could suppress the transcription of many tumor suppressor genes critical to

More and more results confirmed that cancer is a multi-stage process fuelled by many epige‐ netic changes in addition to genetic changes in DNA sequence [4]. Chemical molecules like Trichostatin A (TSA) and 5-aza-2'-deoxycytidine (5-Aza-CdR) targeting epigenetic regula‐ tors such as histone modifications and DNMTs (DNA methyltransferases) have been found to inhibit tumor growth both in vitro and in vivo. By reversing the epigenetic silencing of important tumor suppressor genes, an increasing number of epigenetic drugs such as 5-Aza-CdR, 5-Aza-CR and Vorinostat (SAHA) are currently investigated in the clinical trials for cancer treatment as a single drug or in combination with other epigenetic drugs or other ap‐ proaches such as chemotherapy and showed very promising activities by offering signifi‐

> © 2013 Jin et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

**Detection**

Xian Wang

**1. Introduction**

http://dx.doi.org/10.5772/51419

cancer initiation and progression [1-3].

cant clinical benefits to cancer patients [5-13].

Hongchuan Jin, Yanning Ma, Qi Shen and

Additional information is available at the end of the chapter

## **Circulating Methylated DNA as Biomarkers for Cancer Detection**

Hongchuan Jin, Yanning Ma, Qi Shen and Xian Wang

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51419

#### **1. Introduction**

In addition to genetic alterations including deletion or point mutations, epigenetic changes such as DNA methylation play an important role in silencing tumor suppressor genes dur‐ ing cancer development. By adding a methyl group from S-adenosyl-L-methionine to the cy‐ tosine pyrimidine or adenine purine ring, DNA methylation is important to maintain genome structure and regulate gene expression. In mammalian adult tissues, DNA methyla‐ tion occurs in CpG dinucleotides that often cluster in the genome as CpG islands in the 5' regulatory regions of the genes. Through recruiting transcriptional co-repressors including methyl-CpG-binding domain proteins (MBDs) and chromatin remodeling proteins like his‐ tone deacetylases (HDACs) or impeding the binding of transcriptional activators, DNA methylation could suppress the transcription of many tumor suppressor genes critical to cancer initiation and progression [1-3].

More and more results confirmed that cancer is a multi-stage process fuelled by many epige‐ netic changes in addition to genetic changes in DNA sequence [4]. Chemical molecules like Trichostatin A (TSA) and 5-aza-2'-deoxycytidine (5-Aza-CdR) targeting epigenetic regula‐ tors such as histone modifications and DNMTs (DNA methyltransferases) have been found to inhibit tumor growth both in vitro and in vivo. By reversing the epigenetic silencing of important tumor suppressor genes, an increasing number of epigenetic drugs such as 5-Aza-CdR, 5-Aza-CR and Vorinostat (SAHA) are currently investigated in the clinical trials for cancer treatment as a single drug or in combination with other epigenetic drugs or other ap‐ proaches such as chemotherapy and showed very promising activities by offering signifi‐ cant clinical benefits to cancer patients [5-13].

© 2013 Jin et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

As one of the major epigenetic changes to inactivate tumor suppressor genes critical to hu‐ man cancer development, DNA methylation was recognized as the biomarker for cancer de‐ tection or outcome prediction in addition to the identification of novel tumor suppressor genes. DNA mutations will occur randomly in any nucleotides of one particular gene and the comprehensive determination of DNA mutations is thus very difficult and time-consum‐ ing. In contrast, aberrant DNA hypermethylation usually takes place in defined CpG Islands within the regulatory region of the genes and it is much more convenient to detect DNA methylation in a quantitatively manner. In addition, DNA methylation can be amplified and is thus easily detectable using PCR-based approaches even when the DNA concentration af‐ ter sample extraction is relatively low. Due to such advantages over DNA mutation- or pro‐ tein-based biomarkers, DNA methylation-based biomarkers have been intensively investigated in the recent years. A large body of research reports has proved the value of DNA methylations in the prognosis prediction and detection of various cancers. DNAs used for such methylation analyses are usually extracted from tumor tissues harvested after sur‐ gical operation or biopsy, thus limiting its wide application as the biomarkers for the early detection or screening of human cancers. Recently, it has been reported that there are certain amount of circulating DNAs in the peripheral blood of cancer patients, providing an ideal source to identify novel biomarkers for non-invasive detection of cancers. Both genetic and epigenetic changes found in the genomic DNAs extracted from primary tumor cells could be detected in the circulating DNAs, indicating that the detection of methylated DNAs in the circulation represents a new direction to develop novel biomarkers for cancer detection or screening in a non-invasive manner.

DNA methylation within small amount of CTCs could be the next generation of CTC-based test for cancer detection. However, the cost of such tests will be greatly exacerbated, thus

Circulating Methylated DNA as Biomarkers for Cancer Detection

http://dx.doi.org/10.5772/51419

139

**Figure 1.** Circulating tumor cells and cell free DNA. Circulating Tumor cells (CTC) escape from primary sites and spread into the vessel to form metastases in the distal organs with. Cell free DNAs (cf-DNAs) are released into the circulation

Although its origin and biological relevance remains unknown, circulating cell free DNA (cf-DNA) is supposed to be valuable source to identify cancer markers with ideal sensitivity and specificity for non-invasive detection of cancer [23-24]. Early in 1948, two French scientists Mandel and Metais firstly reported the presence of cf-DNAs in human plasma [25]. Such an important discovery has been unnoticed for a long time until cell-free circulating nucleic acid was found to promote the spread and metastasis of crown gall tumor in plants [26]. Subse‐ quently, increased level of cf-DNAs was found in patients with various diseases such as lupus erythematosus and rheumatoid arthritis cancer [27-28]. In 1977, Leon et al. reported that higher level of circulating DNA in the plasma of cancer patients when compared to healthy con‐ trols. Moreover, greater amounts of cf-DNA were found in the peripheral blood of cancer patients with tumor metastases and cf-DNA levels decreased dramatically after radiothera‐ py while persistently high or increasing DNA concentrations were associated with a lack of response to treatment [29], clearly revealing the potential value of cf-DNA as biomarker for cancer detection. Following studies confirmed that cf-DNAs in the plasma contains genetic and epigenetic changes specific to DNAs within the tumor cells from primary tissues, indicat‐ ing that tumor specific cf-DNAs are originated from tumor cells rather than lymphocytes reacting towards the disease [30-31]. For example, K-Ras mutation was found in cf-DNA from 17 out of 21 patients with pancreatic adenocarcinoma and mutations were similar in corre‐ sponding plasma and tissues samples. Importantly, such DNA alterations were found in

from dead cancer cells or proliferating tumor cells. RBC: red blood cell; WBC: white blood cell.

limiting its wide application in the clinic [20-22].

#### **2. Cell free DNA in the circulation**

According to the origin of circulating tumor-related DNA, it could be grouped into circulat‐ ing cell free DNA or DNA from cells in the blood such as circulating tumor cells (CTC) in cancer patients (Figure 1).

In 1869, the Australian physician Thomas Ashworth observed CTCs in the blood of a cancer patient. Therefore, it was postulated that CTCs were responsible for the tumor metastases in distal sites and should have important prognostic and therapeutic implications [14-16]. However, the number of CTCs is very small compared with blood cells. Usually around 1-10 CTCs together with several million blood cells could be found in 1 ml of whole blood, mak‐ ing the specific and sensitive detection of CTCs very difficult [17-18]. Until recently, technol‐ ogies with the requisite sensitivity and reproducibility for CTC detection have been developed to precisely analyze its biological and clinical relevance. The US Food and Drug Administration (FDA) approved the test for determining CTC levels in patients with meta‐ static breast cancer in 2004. Currently, it has been expanded to other cancer types such as advanced colorectal cancer and prostate cancer. Although CTCs-counting based test have proven its value in predicting prognosis and monitoring therapeutic effects, the number of CTCs per ml of blood limited its sensitivity greatly [19]. With the development of high-sen‐ sitive PCR-based methods, the detection of gene mutations or epigenetic changes such as DNA methylation within small amount of CTCs could be the next generation of CTC-based test for cancer detection. However, the cost of such tests will be greatly exacerbated, thus limiting its wide application in the clinic [20-22].

As one of the major epigenetic changes to inactivate tumor suppressor genes critical to hu‐ man cancer development, DNA methylation was recognized as the biomarker for cancer de‐ tection or outcome prediction in addition to the identification of novel tumor suppressor genes. DNA mutations will occur randomly in any nucleotides of one particular gene and the comprehensive determination of DNA mutations is thus very difficult and time-consum‐ ing. In contrast, aberrant DNA hypermethylation usually takes place in defined CpG Islands within the regulatory region of the genes and it is much more convenient to detect DNA methylation in a quantitatively manner. In addition, DNA methylation can be amplified and is thus easily detectable using PCR-based approaches even when the DNA concentration af‐ ter sample extraction is relatively low. Due to such advantages over DNA mutation- or pro‐ tein-based biomarkers, DNA methylation-based biomarkers have been intensively investigated in the recent years. A large body of research reports has proved the value of DNA methylations in the prognosis prediction and detection of various cancers. DNAs used for such methylation analyses are usually extracted from tumor tissues harvested after sur‐ gical operation or biopsy, thus limiting its wide application as the biomarkers for the early detection or screening of human cancers. Recently, it has been reported that there are certain amount of circulating DNAs in the peripheral blood of cancer patients, providing an ideal source to identify novel biomarkers for non-invasive detection of cancers. Both genetic and epigenetic changes found in the genomic DNAs extracted from primary tumor cells could be detected in the circulating DNAs, indicating that the detection of methylated DNAs in the circulation represents a new direction to develop novel biomarkers for cancer detection or

According to the origin of circulating tumor-related DNA, it could be grouped into circulat‐ ing cell free DNA or DNA from cells in the blood such as circulating tumor cells (CTC) in

In 1869, the Australian physician Thomas Ashworth observed CTCs in the blood of a cancer patient. Therefore, it was postulated that CTCs were responsible for the tumor metastases in distal sites and should have important prognostic and therapeutic implications [14-16]. However, the number of CTCs is very small compared with blood cells. Usually around 1-10 CTCs together with several million blood cells could be found in 1 ml of whole blood, mak‐ ing the specific and sensitive detection of CTCs very difficult [17-18]. Until recently, technol‐ ogies with the requisite sensitivity and reproducibility for CTC detection have been developed to precisely analyze its biological and clinical relevance. The US Food and Drug Administration (FDA) approved the test for determining CTC levels in patients with meta‐ static breast cancer in 2004. Currently, it has been expanded to other cancer types such as advanced colorectal cancer and prostate cancer. Although CTCs-counting based test have proven its value in predicting prognosis and monitoring therapeutic effects, the number of CTCs per ml of blood limited its sensitivity greatly [19]. With the development of high-sen‐ sitive PCR-based methods, the detection of gene mutations or epigenetic changes such as

screening in a non-invasive manner.

cancer patients (Figure 1).

**2. Cell free DNA in the circulation**

138 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 1.** Circulating tumor cells and cell free DNA. Circulating Tumor cells (CTC) escape from primary sites and spread into the vessel to form metastases in the distal organs with. Cell free DNAs (cf-DNAs) are released into the circulation from dead cancer cells or proliferating tumor cells. RBC: red blood cell; WBC: white blood cell.

Although its origin and biological relevance remains unknown, circulating cell free DNA (cf-DNA) is supposed to be valuable source to identify cancer markers with ideal sensitivity and specificity for non-invasive detection of cancer [23-24]. Early in 1948, two French scientists Mandel and Metais firstly reported the presence of cf-DNAs in human plasma [25]. Such an important discovery has been unnoticed for a long time until cell-free circulating nucleic acid was found to promote the spread and metastasis of crown gall tumor in plants [26]. Subse‐ quently, increased level of cf-DNAs was found in patients with various diseases such as lupus erythematosus and rheumatoid arthritis cancer [27-28]. In 1977, Leon et al. reported that higher level of circulating DNA in the plasma of cancer patients when compared to healthy con‐ trols. Moreover, greater amounts of cf-DNA were found in the peripheral blood of cancer patients with tumor metastases and cf-DNA levels decreased dramatically after radiothera‐ py while persistently high or increasing DNA concentrations were associated with a lack of response to treatment [29], clearly revealing the potential value of cf-DNA as biomarker for cancer detection. Following studies confirmed that cf-DNAs in the plasma contains genetic and epigenetic changes specific to DNAs within the tumor cells from primary tissues, indicat‐ ing that tumor specific cf-DNAs are originated from tumor cells rather than lymphocytes reacting towards the disease [30-31]. For example, K-Ras mutation was found in cf-DNA from 17 out of 21 patients with pancreatic adenocarcinoma and mutations were similar in corre‐ sponding plasma and tissues samples. Importantly, such DNA alterations were found in patients with pancreatitis who were diagnosed as pancreatic cancer 5-14 months later, indi‐ cating that release of tumor-specific DNA into the circulation is an early event in cancer development and cf-DNA could be used as the biomarkers for early cancer detection [32]. Treatment resulted in disappearance of K-Ras mutations in plasma DNA in six of nine pa‐ tients. Three patients with a persistently positive K-Ras gene mutation in plasma samples from patients before and after treatment showed early recurrence or progression and pancreatic carcinoma patients with the mutant-type K-ras gene in plasma DNA exhibited a shorter survival time than patients with the wild-type gene, indicating the cf-DNA could be of value in monitoring disease progression or evaluating treatment response [31, 33].

well as the methodologies for cf-DNA extraction and use various internal controls to adjust

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141

**Figure 2.** Schematic introductions of various methods for methylation analyses. MSP, BGS and COBRA are based on bisulfite-mediated conversion of unmethylated cytosines into uracils. CpG methylation could block DNA digestion by some restriction enzymes, making it possible to determine methylation status independent of bisulfite treatment by analyzing digestion products. Alternatively, DNA fragments containing methylated CpG sites could be enriched by an‐ ti-methylcytosine antibody or methylation binding proteins. Advances in next generation genome sequencing tech‐ nology led to the development of noel techniques such as SMRT which can specially analyze 5-methylcytosines with

In general, the detection of DNA methylation could be bisulfite-dependent or -independent

The chemical reaction of sodium bisulfite with DNA could convert unmethylated cytosine of CpG into uracil or UpG but leave methylated cytosine of CpG unchanged. The following analyses such as methylation-and unmethylation specific polymerase chain reaction (M- and U-SP), bisulfite genome sequencing (BGS) or combined bisulfite restriction analysis (CO‐ BRA) could determine the conversion of CpG sites of interest, thus reflecting their methyla‐ tion status as methylated or unmethylated [37]. With varied resolution levels, different bisulfite-dependent DNA methylation analysis methods detect the conversion after bisulfite treatment of genomic DNA, which could have certain artificial effects such as incomplete conversion of unmethylated CpG into UpG, leading to high rate of false negative conclusion

Recently, some new modifications of cytosine in CpG dinucleotides have been discovered such as 5-hydoxymethylcytosine which was called the sixth base since 5-methylcytosine was named as the fifth base [38]. Generated from the oxidation of 5-methylcytosine by the Tet family of enzymes, 5-hydoxymethylcytosine was first found in bacteriophages and recently

possible inter-laboratory variations.

genome wide coverage.

of DNA methylation status.

(Figure 2).

Through quantitatively analyzing plasma DNAs from patients with organ transplantation, Lo et al found that the majority of plasma DNAs was released from the hematopoietic sys‐ tem. However, donor DNA could be detected in the plasma of recipients suffering from the graft rejection because of the large amount of cell death which promotes the release of donor DNAs into the peripheral blood of the recipients [34]. Therefore, it was postulated that cellfree tumor related DNA could originate from the apoptotic tumor cells since high-rate of apoptosis indeed occurs in primary and metastatic tumor tissues. However, cf-DNA quanti‐ ties are significantly reduced in cancer patients after radiotherapy when a great number of tumor cells were believed to undergo apoptotic cell death and cf-DNAs in supernatants of cultured cancer cells increases with cell proliferation rather than apoptosis or necrosis, indi‐ cating that proliferating tumor cells could actively release cf-DNA into the tumor microen‐ vironment and circulation.

In contrast to labile RNAs that were included into the actively secreted exosomes, the nature of cf-DNAs remains to be clarified. As negatively charged molecules, cf-DNA was bound by plasma proteins to escape from endonuclease-mediated degradation. Unfortunately, plasma proteins bound to cf-DNAs was not well characterized yet. Meanwhile, secreted exosomes could remodel microenviroments and promote tumor metastasis since RNAs within exo‐ somes especially microRNA with high stability may influence gene expression in neighbor cells. The biological relevance of cf-DNAs remains unknown. DNA was believed to be more structural rather than functional. However, it was supposed that cf-DNA could play a role as vaccine in tumor microenvironment.

#### **3. Methods for the detection of methylated DNA**

It is unclear so far whether serum or plasma is better for cf-DNA extraction. Although the DNA amount is significantly higher in the serum, the majority of the increase was due to the release of nuclear acids from destroyed blood cells during blood clotting [35]. In addition, the time gap between blooding drawing and DNA extraction as well as the methodologies used for DNA isolation contribute greatly to the amount of cf-DNA harvested. On an aver‐ age, around 30 ng cf-DNA could be extracted from one ml of blood sample [36]. Therefore, in order to determine the quantity of potential cf-DNA-based biomarkers precisely and pro‐ mote its wide application for cancer detection, it is very important to unify the source as well as the methodologies for cf-DNA extraction and use various internal controls to adjust possible inter-laboratory variations.

patients with pancreatitis who were diagnosed as pancreatic cancer 5-14 months later, indi‐ cating that release of tumor-specific DNA into the circulation is an early event in cancer development and cf-DNA could be used as the biomarkers for early cancer detection [32]. Treatment resulted in disappearance of K-Ras mutations in plasma DNA in six of nine pa‐ tients. Three patients with a persistently positive K-Ras gene mutation in plasma samples from patients before and after treatment showed early recurrence or progression and pancreatic carcinoma patients with the mutant-type K-ras gene in plasma DNA exhibited a shorter survival time than patients with the wild-type gene, indicating the cf-DNA could be of value in monitoring

Through quantitatively analyzing plasma DNAs from patients with organ transplantation, Lo et al found that the majority of plasma DNAs was released from the hematopoietic sys‐ tem. However, donor DNA could be detected in the plasma of recipients suffering from the graft rejection because of the large amount of cell death which promotes the release of donor DNAs into the peripheral blood of the recipients [34]. Therefore, it was postulated that cellfree tumor related DNA could originate from the apoptotic tumor cells since high-rate of apoptosis indeed occurs in primary and metastatic tumor tissues. However, cf-DNA quanti‐ ties are significantly reduced in cancer patients after radiotherapy when a great number of tumor cells were believed to undergo apoptotic cell death and cf-DNAs in supernatants of cultured cancer cells increases with cell proliferation rather than apoptosis or necrosis, indi‐ cating that proliferating tumor cells could actively release cf-DNA into the tumor microen‐

In contrast to labile RNAs that were included into the actively secreted exosomes, the nature of cf-DNAs remains to be clarified. As negatively charged molecules, cf-DNA was bound by plasma proteins to escape from endonuclease-mediated degradation. Unfortunately, plasma proteins bound to cf-DNAs was not well characterized yet. Meanwhile, secreted exosomes could remodel microenviroments and promote tumor metastasis since RNAs within exo‐ somes especially microRNA with high stability may influence gene expression in neighbor cells. The biological relevance of cf-DNAs remains unknown. DNA was believed to be more structural rather than functional. However, it was supposed that cf-DNA could play a role

It is unclear so far whether serum or plasma is better for cf-DNA extraction. Although the DNA amount is significantly higher in the serum, the majority of the increase was due to the release of nuclear acids from destroyed blood cells during blood clotting [35]. In addition, the time gap between blooding drawing and DNA extraction as well as the methodologies used for DNA isolation contribute greatly to the amount of cf-DNA harvested. On an aver‐ age, around 30 ng cf-DNA could be extracted from one ml of blood sample [36]. Therefore, in order to determine the quantity of potential cf-DNA-based biomarkers precisely and pro‐ mote its wide application for cancer detection, it is very important to unify the source as

disease progression or evaluating treatment response [31, 33].

140 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

vironment and circulation.

as vaccine in tumor microenvironment.

**3. Methods for the detection of methylated DNA**

**Figure 2.** Schematic introductions of various methods for methylation analyses. MSP, BGS and COBRA are based on bisulfite-mediated conversion of unmethylated cytosines into uracils. CpG methylation could block DNA digestion by some restriction enzymes, making it possible to determine methylation status independent of bisulfite treatment by analyzing digestion products. Alternatively, DNA fragments containing methylated CpG sites could be enriched by an‐ ti-methylcytosine antibody or methylation binding proteins. Advances in next generation genome sequencing tech‐ nology led to the development of noel techniques such as SMRT which can specially analyze 5-methylcytosines with genome wide coverage.

In general, the detection of DNA methylation could be bisulfite-dependent or -independent (Figure 2).

The chemical reaction of sodium bisulfite with DNA could convert unmethylated cytosine of CpG into uracil or UpG but leave methylated cytosine of CpG unchanged. The following analyses such as methylation-and unmethylation specific polymerase chain reaction (M- and U-SP), bisulfite genome sequencing (BGS) or combined bisulfite restriction analysis (CO‐ BRA) could determine the conversion of CpG sites of interest, thus reflecting their methyla‐ tion status as methylated or unmethylated [37]. With varied resolution levels, different bisulfite-dependent DNA methylation analysis methods detect the conversion after bisulfite treatment of genomic DNA, which could have certain artificial effects such as incomplete conversion of unmethylated CpG into UpG, leading to high rate of false negative conclusion of DNA methylation status.

Recently, some new modifications of cytosine in CpG dinucleotides have been discovered such as 5-hydoxymethylcytosine which was called the sixth base since 5-methylcytosine was named as the fifth base [38]. Generated from the oxidation of 5-methylcytosine by the Tet family of enzymes, 5-hydoxymethylcytosine was first found in bacteriophages and recently shown to be abundant in human and mouse brains as well as in embryonic stem cells [39-40]. Although the exact relevance of 5-hydoxymethylcytosine in the genome is still not fully clarified, it has been found to regulate gene expression or promote DNA demethyla‐ tion. The in vitro synthesized artificial oligonucleotides containing 5-hydoxymethylcyto‐ sines can be converted into unmodified cytosines when introduced into mammalian cells, indicating that 5-hydoxymethylcytosine might be one of intermediate products during ac‐ tive DNA demethylation [41]. Therefore, the increase of 5-hydoxymethylcytosine might re‐ flect the demethylation of CpG dinucleotides. Unfortunately, 5-hydoxymethylcytosines, similar to 5-methylcytosines, appear to be resistant to bisulfite-mediated conversion and PCR could amplify DNA fragments containing 5-hydoxymethylcytosines or 5-methylcyto‐ sines with similar efficiency [42-43]. Therefore, bisulfite-dependent methylation analyses could produce false positive results by counting 5-hydoxymethylcytosines into 5-methylcy‐ tosines. In addition to 5-hydroxymethylcytosines, some forms of DNA modifications such as the seventh base, 5-formylcytosine and the eighth base, 5-carboxylcytosine, have been found in mammalian cells recently [44-47]. As the products of 5-hydoxymethylcytosine oxidation through TET hydroxylases, both 5-formylcytosine and 5-carboxylcytosine will be read as the uracil after bisulfite conversion, thus making it impossible for bisulfite-dependent analyses to distinguish unmodified cytosines from 5-formylcytosines and 5-carboxylcytosines.

sine may precipitate methylated CHH and CHG trinucleotide containing DNA fragments in

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143

DNA methylation functions as the signal for DNA-interacting proteins to maintain genome structure or regulate gene expression. The proteins such as MBD1 (methyl-CpG binding do‐ main protein 1), MeCP2 (methyl CpG binding protein 2) and MBD4 (methyl-CpG binding domain protein 4) bind methylated CpG specifically to regulate gene expression [51-52]. Therefore, methyl-CpG binding domain could specifically enrich differentially methylated regions (DMRs) of physiological relevance [53]. Similar to MeDIP, MBD capture specifically enrich methylated CpG sites rather than hydroxymethlated CpG sites. The detailed analysis to compare MeDIP and MBD capture revealed that both enrichment techniques are sensitive enough to identify DMRs in human cancer cells. However, MeDIP enriched more methylat‐ ed regions with low CpG densities while MBD capture favors regions of high CpG densities

Recently, the advance of next generation sequencing led to the development of several novel techniques, making it possible to quantitatively analyze DNA methylation at single nucleo‐ tide resolution with genome wide coverage. Both the single molecule real time sequencing technology (SMRT) and the single-molecule nanopore DNA sequencing platform could dis‐ criminate 5-methylcytosines from other DNA bases including 5-hydroxymethylcytosines even methyladenine independent of bisulfite conversion [54-55]. With many advantages such as less bias during template preparation, lower cost and better accuracy, such new techniques could offer more methods to detect DNA methylation with high specificity and sensitivity in addition to more potential DNA methylation based biomarkers for cancer de‐

**4. Potential DNA methylation biomarkers for cancer detection**

It has been questioned whether the methylated DNA in the circulation is sensitive to detect cancers early enough for curative resection. However, the development of sensitive detection methods confirmed the potential value of DNA methylation in cancer detection (Table 1).

Most of DNA methylation biomarkers are well-known tumor suppressor genes silenced in primary tumor tissues. However, the biomarks do not have to be functional relevant. For ex‐ ample, currently well-used biomarkers such as AFP (Alpha-Fetal Protein), PSA (Prostatespecific antigen) and CEA (Carcinoembryonic antigen) are not tumor suppressor genes with important biological functions. Profiling of methylated DNA in the circulation instead of primary tumor tissues with MeDIP or MBD capture or other methylation specific analyses methods would identify more potential biomarks rather than functional important tumor

addition to DNA sequences with methylated CpG sites.

and identifies the greater proportion of CpG islands [49].

tection and screening.

suppressor genes.

Bisulfite independent analyses such as MedIP (methylated DNA immunoprecipitation) could more or less detect DNA methylation specifically. In bisulfite independent analyses, 5 methylcytosines are differentiated from unmethylated cytosine by either enzyme digestion or affinity enrichment. DNA methylation analysis using restriction enzyme digestion is based on the property of some methylation-sensitive and -resistant restriction enzymes such as HpaII and MspI that target CCGG for digestion. HpaII fails to cut it once the second cyto‐ sine was methylated while MspI-mediated digestion is not affected by DNA methylation, thus making it possible to determine the methylation status of CpG in the context of CCGG tetranucleotides by analyzing the products of DNAs digested by HpaII and MspI respective‐ ly. As a primary method to analyze DNA methylation, it can only determine the methyla‐ tion of CpG in the context of CCGG tetranucleotides and will overlook the majority of CpG dinucleotides in the genome.

The development of monoclonal antibody specific to 5-methylcytosines revolutionized the analyses of DNA methylation [48-49]. Immunoprecipitated DNA by this antibody could be subject to DNA microarray or even deep sequencing to reveal novel sequences or sites con‐ taining 5-methylcytosines [50]. This antibody specifically recognizes 5-methylcytosines but not 5-hydoxymethylcytosines. However, 5-methylcytosines could present not only in CpG dinucleotides but also in CHH or CHG trinucleotides, especially in plants, human embryon‐ ic stem cells and probably cancer cells as well. CHH methylation indicates a 5-methylcyto‐ sine followed by two nucleotides that may not be guanine and CHG methylation refers to a 5-methylcytosine preceding an adenine, thymine or cytosine base followed by guanine. Such non-CpG DNA methylations were enriched at transposons and repetitive regions, although the exact biological relevance remains unknown. However, antibody against 5-methylcyto‐ sine may precipitate methylated CHH and CHG trinucleotide containing DNA fragments in addition to DNA sequences with methylated CpG sites.

shown to be abundant in human and mouse brains as well as in embryonic stem cells [39-40]. Although the exact relevance of 5-hydoxymethylcytosine in the genome is still not fully clarified, it has been found to regulate gene expression or promote DNA demethyla‐ tion. The in vitro synthesized artificial oligonucleotides containing 5-hydoxymethylcyto‐ sines can be converted into unmodified cytosines when introduced into mammalian cells, indicating that 5-hydoxymethylcytosine might be one of intermediate products during ac‐ tive DNA demethylation [41]. Therefore, the increase of 5-hydoxymethylcytosine might re‐ flect the demethylation of CpG dinucleotides. Unfortunately, 5-hydoxymethylcytosines, similar to 5-methylcytosines, appear to be resistant to bisulfite-mediated conversion and PCR could amplify DNA fragments containing 5-hydoxymethylcytosines or 5-methylcyto‐ sines with similar efficiency [42-43]. Therefore, bisulfite-dependent methylation analyses could produce false positive results by counting 5-hydoxymethylcytosines into 5-methylcy‐ tosines. In addition to 5-hydroxymethylcytosines, some forms of DNA modifications such as the seventh base, 5-formylcytosine and the eighth base, 5-carboxylcytosine, have been found in mammalian cells recently [44-47]. As the products of 5-hydoxymethylcytosine oxidation through TET hydroxylases, both 5-formylcytosine and 5-carboxylcytosine will be read as the uracil after bisulfite conversion, thus making it impossible for bisulfite-dependent analyses

142 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

to distinguish unmodified cytosines from 5-formylcytosines and 5-carboxylcytosines.

dinucleotides in the genome.

Bisulfite independent analyses such as MedIP (methylated DNA immunoprecipitation) could more or less detect DNA methylation specifically. In bisulfite independent analyses, 5 methylcytosines are differentiated from unmethylated cytosine by either enzyme digestion or affinity enrichment. DNA methylation analysis using restriction enzyme digestion is based on the property of some methylation-sensitive and -resistant restriction enzymes such as HpaII and MspI that target CCGG for digestion. HpaII fails to cut it once the second cyto‐ sine was methylated while MspI-mediated digestion is not affected by DNA methylation, thus making it possible to determine the methylation status of CpG in the context of CCGG tetranucleotides by analyzing the products of DNAs digested by HpaII and MspI respective‐ ly. As a primary method to analyze DNA methylation, it can only determine the methyla‐ tion of CpG in the context of CCGG tetranucleotides and will overlook the majority of CpG

The development of monoclonal antibody specific to 5-methylcytosines revolutionized the analyses of DNA methylation [48-49]. Immunoprecipitated DNA by this antibody could be subject to DNA microarray or even deep sequencing to reveal novel sequences or sites con‐ taining 5-methylcytosines [50]. This antibody specifically recognizes 5-methylcytosines but not 5-hydoxymethylcytosines. However, 5-methylcytosines could present not only in CpG dinucleotides but also in CHH or CHG trinucleotides, especially in plants, human embryon‐ ic stem cells and probably cancer cells as well. CHH methylation indicates a 5-methylcyto‐ sine followed by two nucleotides that may not be guanine and CHG methylation refers to a 5-methylcytosine preceding an adenine, thymine or cytosine base followed by guanine. Such non-CpG DNA methylations were enriched at transposons and repetitive regions, although the exact biological relevance remains unknown. However, antibody against 5-methylcyto‐

DNA methylation functions as the signal for DNA-interacting proteins to maintain genome structure or regulate gene expression. The proteins such as MBD1 (methyl-CpG binding do‐ main protein 1), MeCP2 (methyl CpG binding protein 2) and MBD4 (methyl-CpG binding domain protein 4) bind methylated CpG specifically to regulate gene expression [51-52]. Therefore, methyl-CpG binding domain could specifically enrich differentially methylated regions (DMRs) of physiological relevance [53]. Similar to MeDIP, MBD capture specifically enrich methylated CpG sites rather than hydroxymethlated CpG sites. The detailed analysis to compare MeDIP and MBD capture revealed that both enrichment techniques are sensitive enough to identify DMRs in human cancer cells. However, MeDIP enriched more methylat‐ ed regions with low CpG densities while MBD capture favors regions of high CpG densities and identifies the greater proportion of CpG islands [49].

Recently, the advance of next generation sequencing led to the development of several novel techniques, making it possible to quantitatively analyze DNA methylation at single nucleo‐ tide resolution with genome wide coverage. Both the single molecule real time sequencing technology (SMRT) and the single-molecule nanopore DNA sequencing platform could dis‐ criminate 5-methylcytosines from other DNA bases including 5-hydroxymethylcytosines even methyladenine independent of bisulfite conversion [54-55]. With many advantages such as less bias during template preparation, lower cost and better accuracy, such new techniques could offer more methods to detect DNA methylation with high specificity and sensitivity in addition to more potential DNA methylation based biomarkers for cancer de‐ tection and screening.

#### **4. Potential DNA methylation biomarkers for cancer detection**

It has been questioned whether the methylated DNA in the circulation is sensitive to detect cancers early enough for curative resection. However, the development of sensitive detection methods confirmed the potential value of DNA methylation in cancer detection (Table 1).

Most of DNA methylation biomarkers are well-known tumor suppressor genes silenced in primary tumor tissues. However, the biomarks do not have to be functional relevant. For ex‐ ample, currently well-used biomarkers such as AFP (Alpha-Fetal Protein), PSA (Prostatespecific antigen) and CEA (Carcinoembryonic antigen) are not tumor suppressor genes with important biological functions. Profiling of methylated DNA in the circulation instead of primary tumor tissues with MeDIP or MBD capture or other methylation specific analyses methods would identify more potential biomarks rather than functional important tumor suppressor genes.


**Cancer Markers Sensitivity Specificity Methods Ref.**

77/105 (73%) 12/35 (34%)

23/33 (70%) 25/69 (36%)

Most of the methods used for methylation biomarkers analyses are still bisulfite dependent. Few reports used MS-AP-PCR (methylation-sensitive arbitrarily primed PCR) which takes the advantage of methylation sensitive restriction endonucleases to distinguish methylated CpG from unmethylated form, although the sensitivity seems to be lower than MSP [56-57]. Interestingly, combination of more than one methylated DNA as a methylation panel could great increase the sensitivity for cancer detection without significant reduction of specificity. Unfortunately, most of studies were performed in a retrospective manner. More prospective studies with large sample sizes will be warranted to compare different approaches especial‐ ly bisulfite-independent methods in addition to confirm the value of DNA methylation for

With the development of the next generation genome sequencing as well as single molecular PCR, it became possible to analyze trace amount of DNAs including circulating cell-free DNA. Circulating tumor cells have been proven its value in prognosis predication even ear‐ ly detection of various cancers. The analyses of methylated DNAs in the circulating will be the next promising epigenetic biomarkers for cancer detection. As one of the intermediate products of DNA demethylation, 5-hydroxymethlcytosines are resistant to bisulfite conver‐ sion. Therefore, it should be carefully to interpret the data of methylation analyses based on bisulfite treatment due to potentially high rate of false positive results. Although some me‐ thylated DNAs were found to valuable as a single biomarker for cancer detection, more po‐ tential DNA methylations will be found after the wide application of SMRT and other sequencing platforms with high speed, depth and accuracy. DNA methylation signatures in‐ cluding a panel of methylated DNAs will show the potential in the early diagnosis or screening and prognosis or therapy response prediction of many cancers. In addition, such DNA methylation biomarkers could be more sensitive and specific for cancer detection when combined with well-used biochemical biomarkers. However, unified methods with gold standards will be warranted to promote the development and clinical application of

N/A

15/15 (100%)

22/22 (100%) 31/31 (100%) MSP MSP

http://dx.doi.org/10.5772/51419

Circulating Methylated DNA as Biomarkers for Cancer Detection

MSP MSP [78] [79] 145

[80] [81]

CDKN2A (INK4A) CDKN2A (INK4A)

GSTP1

**Table 1.** Methylated DNA biomarkers in the literature.

**5. Conclusion and Perspectives**

DNA methylation biomarkers.

Prostate cancer GSTP1

cancer detection.


**Table 1.** Methylated DNA biomarkers in the literature.

**Cancer Markers Sensitivity Specificity Methods Ref.**

N/A N/A

N/A

N/A

N/A

N/A N/A 36/52 (70%) 17/17 (100%) 150/179 (84%) 10/10 (100%) 154/179 (86%) 123/179 (69%)

31/31 (100%)

44/44 (100%)

54/54 (100%) 54/54 (100%)

30/30 (100%) 30/30 (100%) 30/30 (100%) 30/30 (100%) 30/30 (100%) 30/30 (100%)

48/48 (100%) 35/35 (100%)

50/50 (100%)

N/A

N/A N/A N/A N/A N/A

N/A N/A N/A N/A N/A N/A MSP MSP MSP

MS-AP-PCR MS-AP-PCR

MSP MSP MSP MSP MSP MSP MSP MSP MSP MSP

MSP MSP MSP

MSP MSP MSP MSP MSP MSP

MSP MSP MSP MSP MSP

MSP MSP

MSP MSP MSP MSP MSP MSP MSP [58]

[59]

[56]

[57]

[60] [61] [62] [63] [64] [65] [66] [67] [66]

[68]

[69]

[70]

[71]

[72]

[73] [74]

[75]

[76] [77]

13/27 (48%) 2/27 (7%) 19/86 (22%)

5/35 (14%)

6/43 (14%)

3/18 (17%) 14/52 (27%) 13/94 (11%) 21/58 (36%) 25/30 (83%) 32/46 (70%) 68/133 (51%) 11/17 (65%) 92/133 (69%) 87/133 (65%)

13/52 (25%) 2/32 (6%) 7/38 (18%)

31/54 (57%) 28/54 (52%) 30/54 (56%) 26/54 (48%) 18/54 (15%) 45/54 (83%)

8/95 (8%) 3/95 (3%) 14/95 (15%) 21/95 (22%) N/A

13/22 (45%) 4/25 (16%)

3/22 (14%) 4/22 (18%) 1/22 (5%) 4/22 (18%) 11/22 (50%) N/A 42/89 (47%)

Bladder cancer CDKN2A (ARF) CDKN2A (INK4A)

Breast cancer CDKN2A (INK4A)

ALX4 CDH4 NGFR RUNX3 SEPT9 TMEFF2

APC APC

DAPK1 GSTP1 Panel of five

DAPK1 MGMT Panel of three DAPK1

Liver cancer CDKN2A (INK4A) CDKN2B (INK4B)

Lung cancer CDKN2A (INK4A) DAPK1 GSTP1 MGMT Panel of four CDKN2A (INK4A)

APC

CDKN2A (INK4A)

CDKN2A (INK4A) CDKN2B (INK4B)

CDKN2A (INK4A)

Colorectal cancerMLH1

Esophageal cancer

Gastric cancer CDH1

Head and neck cancer

CDKN2A (INK4A)

144 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

CDKN2A (INK4A)

CDKN2A (INK4A) CDKN2A (INK4A) CDKN2A (INK4A)

Most of the methods used for methylation biomarkers analyses are still bisulfite dependent. Few reports used MS-AP-PCR (methylation-sensitive arbitrarily primed PCR) which takes the advantage of methylation sensitive restriction endonucleases to distinguish methylated CpG from unmethylated form, although the sensitivity seems to be lower than MSP [56-57]. Interestingly, combination of more than one methylated DNA as a methylation panel could great increase the sensitivity for cancer detection without significant reduction of specificity. Unfortunately, most of studies were performed in a retrospective manner. More prospective studies with large sample sizes will be warranted to compare different approaches especial‐ ly bisulfite-independent methods in addition to confirm the value of DNA methylation for cancer detection.

#### **5. Conclusion and Perspectives**

With the development of the next generation genome sequencing as well as single molecular PCR, it became possible to analyze trace amount of DNAs including circulating cell-free DNA. Circulating tumor cells have been proven its value in prognosis predication even ear‐ ly detection of various cancers. The analyses of methylated DNAs in the circulating will be the next promising epigenetic biomarkers for cancer detection. As one of the intermediate products of DNA demethylation, 5-hydroxymethlcytosines are resistant to bisulfite conver‐ sion. Therefore, it should be carefully to interpret the data of methylation analyses based on bisulfite treatment due to potentially high rate of false positive results. Although some me‐ thylated DNAs were found to valuable as a single biomarker for cancer detection, more po‐ tential DNA methylations will be found after the wide application of SMRT and other sequencing platforms with high speed, depth and accuracy. DNA methylation signatures in‐ cluding a panel of methylated DNAs will show the potential in the early diagnosis or screening and prognosis or therapy response prediction of many cancers. In addition, such DNA methylation biomarkers could be more sensitive and specific for cancer detection when combined with well-used biochemical biomarkers. However, unified methods with gold standards will be warranted to promote the development and clinical application of DNA methylation biomarkers.

#### **Acknowledgements**

This work was supported by the National Natural Science Foundation of China (81071963; 81071652), Program for Innovative Research Team in Science and technology of Zhejiang Province (2010R50046) and Program for Qianjiang Scholarship in Zhejiang Province (2011R10061; 2011R10073).

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Circulating Methylated DNA as Biomarkers for Cancer Detection

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#### **Author details**

Hongchuan Jin, Yanning Ma, Qi Shen and Xian Wang\*

\*Address all correspondence to: wangx118@yahoo.com

Department of Medical Oncology, Laboratory of Cancer Epigenetics, Biomedical Research Center, Sir Runrun Shaw Hospital, Zhejiang University, China

#### **References**


[9] Font, P. (2011). Azacitidine for the treatment of patients with acute myeloid leukemia with 20%-30% blasts and multilineage dysplasia. *Adv Ther*, 3(28), 1-9.

**Acknowledgements**

(2011R10061; 2011R10073).

Hongchuan Jin, Yanning Ma, Qi Shen and Xian Wang\*

146 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

\*Address all correspondence to: wangx118@yahoo.com

cer. *Nat Rev Genet*, 3, 415-428.

*cent Clin Trials*, 1, 169-182.

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Center, Sir Runrun Shaw Hospital, Zhejiang University, China

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netics joins genetics. *Trends Genet*, 16, 168-174.

therapeutic targets. *Nat Biotechnol*, 28, 1069-1078.

**Author details**

**References**

This work was supported by the National Natural Science Foundation of China (81071963; 81071652), Program for Innovative Research Team in Science and technology of Zhejiang Province (2010R50046) and Program for Qianjiang Scholarship in Zhejiang Province

Department of Medical Oncology, Laboratory of Cancer Epigenetics, Biomedical Research

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**Chapter 7**

**DNA Methylation, Stem Cells and Cancer**

Cancer has been traditionally seen as a disease characterized by many genetic alterations, but recent studies have proven the implications of epigenetic abnormalities along carcino‐

The fundamental base of carcinogenesis is described by two major models: clonal evolution

In the past few years 'cancer stem cells' (CSCs) area has become an interesting field of cancer research. In 19th century, Durante and Conheim [6] and after one hundred year Sell and Pierce [6, 7] issued the hypothesis that stem cells could induce cancer in all type of tissues. Unlike normal tissue stem cells, cancer stem cells are characterized by an abnormal differen‐

**•** chemicals – John Hill, in 1761, was the first who showed that the chemicals agents pro‐

**•** infections – Francis Peyton Rous was the pathologist awarded the Nobel Prize in Medicine for his research that reported that the viral agents are involved in the origin of cancer [10]; **•** mutations – Theodor Heinrich Boveri and Von Hansemann argued the association be‐

and reproduction in any medium, provided the original work is properly cited.

© 2013 Dricu et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

The five principal factors, reported to be involved in carcinogenesis are:

tween development of cancer and abnormal mitoses [11];

Anica Dricu, Stefana Oana Purcaru,

Oana Daianu, Bogdan Stoleru,

Ligia Gabriela Tataranu

http://dx.doi.org/10.5772/53263

and cancer stem cell (CSC) model [3-5].

tiation rate, which can lead to tumor [8, 9].

duce cancer of the nasal cavity [6];

**1. Introduction**

genesis [1, 2].

Alice Sandra Buteica, Daniela Elise Tache,

Amelia Mihaela Dobrescu, Tiberiu Daianu and

Additional information is available at the end of the chapter


### **DNA Methylation, Stem Cells and Cancer**

Anica Dricu, Stefana Oana Purcaru, Alice Sandra Buteica, Daniela Elise Tache, Oana Daianu, Bogdan Stoleru, Amelia Mihaela Dobrescu, Tiberiu Daianu and Ligia Gabriela Tataranu

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53263

#### **1. Introduction**

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[79] Bearzatto, A., Conte, D., Frattini, M., Zaffaroni, N., Andriani, F., Balestra, D., Tavec‐ chio, L., Daidone, M. G., & Sozzi, G. (2002). p16INK4ahypermethylation detected by fluorescent methylation-specific PCR in plasmas from non-small cell lung cancer.

[80] Goessl, C., Krause, H., Müller, M., Heicappell, R., Schrader, M., Sachsinger, J., & Mill‐ er, K. (2000). Fluorescent methylation-specific polymerase chain reaction for DNAbased detection of prostate cancer in bodily fluids. *Cancer research*, 60, 5941-5945.

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tion in the plasma and serum of liver cancer patients. *Cancer research*, 59, 71.

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lung cancer. *International journal of oncology*, 19, 277.

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*Clinical Cancer Research*, 8, 3782-3787.

prostate cancer. *Urology*, 60, 1131-1135.

tumors and serum of head and neck cancer patients. *Cancer research*, 60, 892.

433-437.

Cancer has been traditionally seen as a disease characterized by many genetic alterations, but recent studies have proven the implications of epigenetic abnormalities along carcino‐ genesis [1, 2].

The fundamental base of carcinogenesis is described by two major models: clonal evolution and cancer stem cell (CSC) model [3-5].

In the past few years 'cancer stem cells' (CSCs) area has become an interesting field of cancer research. In 19th century, Durante and Conheim [6] and after one hundred year Sell and Pierce [6, 7] issued the hypothesis that stem cells could induce cancer in all type of tissues. Unlike normal tissue stem cells, cancer stem cells are characterized by an abnormal differen‐ tiation rate, which can lead to tumor [8, 9].

The five principal factors, reported to be involved in carcinogenesis are:


© 2013 Dricu et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

**•** teratocarcinomas – the field theory, which explains that in pathology of cancer are impli‐ cated a mixture of mature and differentiated cells and also embryonic tissue [12];

in tumor cells, DNA methylation patterns become altered. A family of germline-specific genes that use DNA methylation as a primary silencing mechanism, has been indicated as a stem cells signature. These germline-specific genes expression in tumors, has also been hy‐

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 155

Post-translational modification of histone proteins is an important area of regulation epige‐ netic. The post-translational modifications of the N-terminal tail domains include: methyla‐ tion, acetylation, phosphorylation, citrullination, ADP-ribosylation, sumoylation and ubiquitination [18-20]. The most studied of these modifications are methylation and acetyla‐ tion. Modifications of histone N terminal ends by methylation and acetylation processes are closely related to cancer and are done by the competition of two families of enzymes: his‐ tone acetyltransferases (HAT) and histone acetylases (HDAC). Lysine residues acetylation in the histone H3 and H4 by (HAT and lysine 4 methylation in the histone H3 (H3K4me) by histone methyltransferase (HMT) are generally correlated with active transcription of chro‐ matin. In contrast, methylation of lysine 9 and lysine 27 (H3K9, H3K27me) in the histone 3,

pothesised to reflect the expansion of constitutively expressing cancer stem cells [8].

have been considered as markers in transcriptionally silenced-chromatin [21, 22].

histone modifications in stem cells influence carcinogenesis.

formation of stem cells in cancer stem cells [14].

changes of microRNA expression [23].

progression and therapy.

**2. DNA methylation**

cysteine (SAH) [23].

Dysregulation of epigenetic mechanisms in stem cells may induce alteration in stem cells function, (i.e. self-renewal and differentiation potential), leading to cancer initiation and pro‐ gression. During the last years, a major challenge in cancer biology is to elucidate how the

Thus, epigenetic control of gene expression patterns in embryogenesis, stem cells and cancer stem cells is a very important aspect for our understanding of human cancer development,

According to the cancer stem cell theory, aberrant epigenetic changes may allow the trans‐

Epigenetic regulation is realized by modifications that consist in four important mecha‐ nisms: DNA methylation, covalent modification of histone, nucleosome positioning and

The biological process of DNA methylation is found in both eukaryotic and prokaryotic cells and it can be involved in pathogenesis of several diseases, especially in cancer. This is the most studied mechanism of epigenetic regulation, consisting in addition of a methyl group to the carbon-5 position of the pyrimidine base cytosine (C) from the nucleotide structure cytidine-5'-monophosphate (CMP). S-adenosyl-L-methionine (SAM), the active form of amino acid methionine, is the donor of methyl group resulting S-adenosylhomo‐

In the mammalian genome, cytosine is coupled to guanine (G) to form a base pair, common‐ ly called cytosine-phosphate-guanine (CpG) dinucleotides. These dinucleotides are general‐

**•** epigenetic alterations – coerce to development of abnormal phenotypes, without any structural changes of DNA [13];

The term epigenetic was introduced by Conrad Waddington in 1942, to explain the relation‐ ship between environment and genome. Model of "cancer stem cells" indicates that epige‐ netic changes occurred in stem or precursor cell are the earliest events that take place in cancer [14].

There are two primary mechanisms involved in the epigenetic process: methylation of DNA and covalent modification of histones [15].

DNA methylation is an inheritance mechanism, fundamentally important in normal devel‐ opment and cellular differentiation in mammalian organisms. This is a post-replication DNA modification, by the addition of a methyl group to carbon 5 (C5] of the pyrimidine ring of cytosines, predominantly in cytosine-phospho-guanine (CpG) dinucleotides.

In the eukaryotic cells, the pattern of methylation is the result of complex interactions be‐ tween three types of normal methylation processes: *de novo* methylation, the maintenance of existing methylation and demethylation. DNA-methylation is catalyzed by several DNA cytosine-5-methyltransferases (DNMTs), which can catalyse cytosine methylation in differ‐ ent sequence context. DNMT family include: DNMT1, which is responsible for methylation maintenance and DNMT3a and DNMT3b, which are responsible for the *de novo* DNAmethylation [16].

The decisive developmental effect of DNA-methylation on gene expression is the long-term silencing of gene expression. In human, the process of DNA-methylation is associated with transcriptional silencing imprinted genes and X-chromosome inactivation. Both genomic im‐ printing and X-chromosome inactivation are suggested to regulate gene expression in em‐ bryonic and fetal growth.

Dysregulated normal imprinting is supposed to induce embryonic death and to impair fetal growth. Defects in DNA-methylation process may also have major consequences for embry‐ onic development and are associated with congenital defects, autoimmunity, aging and ma‐ lignant transformation.

In recent years, the human methylation profile of the whole genome has been investigated by DNA methylomic studies and altered DNA methylation has been found in cancer DNA [2, 17] The transformation of normal cells into dysplastic and cancerous cells is due to a broad range of genetic and epigenetic changes. Some of the epigenetic mechanisms of initia‐ tion and progression of cancer are strongly related to post translational modifications of his‐ tones, among which the methylation process is highly involved. The resistance of various types of cancer to therapy led to the hypothesis that cancers present some cells, able to self renew and differentiate into all types of cells that compose a tumor, named cancer stem cells [8]. During embryonic development, characteristic patterns of CpG methylation are pro‐ duced in the different cell lineages that are then well conserved in normal adult cells, while in tumor cells, DNA methylation patterns become altered. A family of germline-specific genes that use DNA methylation as a primary silencing mechanism, has been indicated as a stem cells signature. These germline-specific genes expression in tumors, has also been hy‐ pothesised to reflect the expansion of constitutively expressing cancer stem cells [8].

Post-translational modification of histone proteins is an important area of regulation epige‐ netic. The post-translational modifications of the N-terminal tail domains include: methyla‐ tion, acetylation, phosphorylation, citrullination, ADP-ribosylation, sumoylation and ubiquitination [18-20]. The most studied of these modifications are methylation and acetyla‐ tion. Modifications of histone N terminal ends by methylation and acetylation processes are closely related to cancer and are done by the competition of two families of enzymes: his‐ tone acetyltransferases (HAT) and histone acetylases (HDAC). Lysine residues acetylation in the histone H3 and H4 by (HAT and lysine 4 methylation in the histone H3 (H3K4me) by histone methyltransferase (HMT) are generally correlated with active transcription of chro‐ matin. In contrast, methylation of lysine 9 and lysine 27 (H3K9, H3K27me) in the histone 3, have been considered as markers in transcriptionally silenced-chromatin [21, 22].

Dysregulation of epigenetic mechanisms in stem cells may induce alteration in stem cells function, (i.e. self-renewal and differentiation potential), leading to cancer initiation and pro‐ gression. During the last years, a major challenge in cancer biology is to elucidate how the histone modifications in stem cells influence carcinogenesis.

Thus, epigenetic control of gene expression patterns in embryogenesis, stem cells and cancer stem cells is a very important aspect for our understanding of human cancer development, progression and therapy.

#### **2. DNA methylation**

**•** teratocarcinomas – the field theory, which explains that in pathology of cancer are impli‐ cated a mixture of mature and differentiated cells and also embryonic tissue [12];

**•** epigenetic alterations – coerce to development of abnormal phenotypes, without any

The term epigenetic was introduced by Conrad Waddington in 1942, to explain the relation‐ ship between environment and genome. Model of "cancer stem cells" indicates that epige‐ netic changes occurred in stem or precursor cell are the earliest events that take place in

There are two primary mechanisms involved in the epigenetic process: methylation of DNA

DNA methylation is an inheritance mechanism, fundamentally important in normal devel‐ opment and cellular differentiation in mammalian organisms. This is a post-replication DNA modification, by the addition of a methyl group to carbon 5 (C5] of the pyrimidine

In the eukaryotic cells, the pattern of methylation is the result of complex interactions be‐ tween three types of normal methylation processes: *de novo* methylation, the maintenance of existing methylation and demethylation. DNA-methylation is catalyzed by several DNA cytosine-5-methyltransferases (DNMTs), which can catalyse cytosine methylation in differ‐ ent sequence context. DNMT family include: DNMT1, which is responsible for methylation maintenance and DNMT3a and DNMT3b, which are responsible for the *de novo* DNA-

The decisive developmental effect of DNA-methylation on gene expression is the long-term silencing of gene expression. In human, the process of DNA-methylation is associated with transcriptional silencing imprinted genes and X-chromosome inactivation. Both genomic im‐ printing and X-chromosome inactivation are suggested to regulate gene expression in em‐

Dysregulated normal imprinting is supposed to induce embryonic death and to impair fetal growth. Defects in DNA-methylation process may also have major consequences for embry‐ onic development and are associated with congenital defects, autoimmunity, aging and ma‐

In recent years, the human methylation profile of the whole genome has been investigated by DNA methylomic studies and altered DNA methylation has been found in cancer DNA [2, 17] The transformation of normal cells into dysplastic and cancerous cells is due to a broad range of genetic and epigenetic changes. Some of the epigenetic mechanisms of initia‐ tion and progression of cancer are strongly related to post translational modifications of his‐ tones, among which the methylation process is highly involved. The resistance of various types of cancer to therapy led to the hypothesis that cancers present some cells, able to self renew and differentiate into all types of cells that compose a tumor, named cancer stem cells [8]. During embryonic development, characteristic patterns of CpG methylation are pro‐ duced in the different cell lineages that are then well conserved in normal adult cells, while

ring of cytosines, predominantly in cytosine-phospho-guanine (CpG) dinucleotides.

structural changes of DNA [13];

154 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

and covalent modification of histones [15].

cancer [14].

methylation [16].

bryonic and fetal growth.

lignant transformation.

According to the cancer stem cell theory, aberrant epigenetic changes may allow the trans‐ formation of stem cells in cancer stem cells [14].

Epigenetic regulation is realized by modifications that consist in four important mecha‐ nisms: DNA methylation, covalent modification of histone, nucleosome positioning and changes of microRNA expression [23].

The biological process of DNA methylation is found in both eukaryotic and prokaryotic cells and it can be involved in pathogenesis of several diseases, especially in cancer. This is the most studied mechanism of epigenetic regulation, consisting in addition of a methyl group to the carbon-5 position of the pyrimidine base cytosine (C) from the nucleotide structure cytidine-5'-monophosphate (CMP). S-adenosyl-L-methionine (SAM), the active form of amino acid methionine, is the donor of methyl group resulting S-adenosylhomo‐ cysteine (SAH) [23].

In the mammalian genome, cytosine is coupled to guanine (G) to form a base pair, common‐ ly called cytosine-phosphate-guanine (CpG) dinucleotides. These dinucleotides are general‐ ly methylated (CpG-poor regions) with the exception of GC-rich regions known as the CpG islands [24, 25]. In human normal cells, it was observed that CpG islands are often hypome‐ thylated. In oncogenesis CpG islands suffer a hypermethylation process whereas the entire pool CpG-poor regions are hypomethylated. DNA hypermethylation and hypomethylation coexist in cancer cells, both processes demonstrating the importance of DNA methylation in sustaining a normal gene expression pattern, genomic imprinting and silencing of genes in‐ volved in X-chromosome inactivation [1, 23, 26-29].

tumour types, inducing genes transcriptional activation. In their study, Costa et al. hypothe‐

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 157

In addition to DNA hypermethylation and hypomethylation, a DNA demethylation process was also described. While the active DNA demethylation process takes place in presence of enzymes that catalyzed specific reactions, passive DNA demethylation can occur during replication cycles and operates on DNA methyltransferases [42, 43]. Although the demethy‐ lation process is not fully elucidated, there are many studies showing transient involvement of this process in various types of tumors, especially in advanced stages of their develop‐ ment [44]. The mechanism of DNA demethylation in cancer has been relatively less studied*.* Until just a few years ago, scientists believed that the hypomethylation affects the whole ge‐ nome, randomly [45]. Dysregulations in embryonic normal development and also in normal stem cells development, are generated by many signaling pathways, which can be associat‐ ed with cancer. Pathways signaling implicated in regulation of normal stem cell evolution are also involved in stem cells self-renewal and carcinogenesis. The most common signaling pathways in all processes mentioned are: Wnt, Notch and Sonic hedgehog (Shh) [46, 47]. Other signaling pathways reported to be involved in stem cell maintenance and pluripoten‐

It is well-known that Wingless gene encodes the Wnt protein family that control the self– renewal and tumorigenesis processes. Wnt protein is well preserved from Drosophila mela‐ nogaster and has an essential role during normal embryonic development. Wnt protein is a part of a particularly signalling pathway, common to humans and Wnt protein dysregula‐ tion was reported to be involved in the development of tumors. It has been demonstrated that some genes implicated in the Wnt signaling pathway, are inactivated by promoter hy‐

The Notch signaling pathway was also suggested to have an important role in stem cell dif‐ ferentiation, proliferation and oncogenesis as well. Scientists have shown that in humans, exist four Notch paralogs (i.e. Notch 1, 2, 3, and 4) and five ligands (i.e. Delta-like 1, 3, 4 and Jagged 1 and 2). Activation of these Notch paralogs are found in stem cell self-renewal but

Some epigenetic changes like histone methylation and downregulation of gene expression, which collaborate with the Notch developmental pathway during oncogenesis, were also

Chromatin is represented by the mandatory association between nuclear DNA and proteins. Chromatin presents different compression degrees during to cell cycles. It exists in two dif‐

Euchromatin has more non repetitive DNA with prevailing of guanine and cytosine bases and nonhistonic proteins; it is also less condensed and represents the active and transcrip‐

sized that expression of CG genes may be indispensable for stem cell biology [40, 41].

cy are: TGF-beta, MET, MYC, EGF, p53, BMI, etc [5].

also in many types of cancers [49].

described [50].

permethylation, generating lung metastasis from primary tumors [48].

**3. Chromatin dynamic and histones modifications**

ferent types: euchromatin and heterochromatin.

The DNA methylation process is catalyzed by specific DNA methyltransferase (DNMTs). In human cells, five types of DNMTs enzymes have been reported [23, 30-33]. DNMT1 - DNA (cytosine-5-)-methyltransferase 1, with role in regulation of normal tissue-specific methyla‐ tion; unusual methylation is related to the appearance of human cancer. DNMT2 has an un‐ certain role in human health and illness [34]. Grant A Challen et al. have shown that DNMT3A and DNMT3B are implicated in embryonic stem cells differentiation [35] and DNMT3L was reported to stimulate the activity of both DNMT3A and DNMT3B [36].

**Figure 1.** Methylation of cytosine. Cytosine methylation is one of the most extensive studied epigenetic processes. The donor of the methyl group is the active form of methionine, S-adenosyl-L-methionine (SAM) and its addition to cyto‐ sine is realised at the carbon-5 position.

Over the past two decades, it was shown that DNA methylation plays a major role in the regulation of the specific gene expression, during mitotic cell division, in the normal mam‐ malian cell, as well as in the stem cell [13, 37, 38].

A new hypothesize about tumorigenesis consists in dysregulation of the stem cell self-re‐ newal process. Dissemination of cancer stem cells is suggested to be induced by gene muta‐ tions and epigenetic modifications that may lead to metastasis [39].

DNA hypomethylation was found to activate cancer-germline (CG) genes or cancer-testis (CT) gene family in tumours. The promoter region of CG genes is demethylated in a several tumour types, inducing genes transcriptional activation. In their study, Costa et al. hypothe‐ sized that expression of CG genes may be indispensable for stem cell biology [40, 41].

ly methylated (CpG-poor regions) with the exception of GC-rich regions known as the CpG islands [24, 25]. In human normal cells, it was observed that CpG islands are often hypome‐ thylated. In oncogenesis CpG islands suffer a hypermethylation process whereas the entire pool CpG-poor regions are hypomethylated. DNA hypermethylation and hypomethylation coexist in cancer cells, both processes demonstrating the importance of DNA methylation in sustaining a normal gene expression pattern, genomic imprinting and silencing of genes in‐

The DNA methylation process is catalyzed by specific DNA methyltransferase (DNMTs). In human cells, five types of DNMTs enzymes have been reported [23, 30-33]. DNMT1 - DNA (cytosine-5-)-methyltransferase 1, with role in regulation of normal tissue-specific methyla‐ tion; unusual methylation is related to the appearance of human cancer. DNMT2 has an un‐ certain role in human health and illness [34]. Grant A Challen et al. have shown that DNMT3A and DNMT3B are implicated in embryonic stem cells differentiation [35] and DNMT3L was reported to stimulate the activity of both DNMT3A and DNMT3B [36].

**Figure 1.** Methylation of cytosine. Cytosine methylation is one of the most extensive studied epigenetic processes. The donor of the methyl group is the active form of methionine, S-adenosyl-L-methionine (SAM) and its addition to cyto‐

Over the past two decades, it was shown that DNA methylation plays a major role in the regulation of the specific gene expression, during mitotic cell division, in the normal mam‐

A new hypothesize about tumorigenesis consists in dysregulation of the stem cell self-re‐ newal process. Dissemination of cancer stem cells is suggested to be induced by gene muta‐

DNA hypomethylation was found to activate cancer-germline (CG) genes or cancer-testis (CT) gene family in tumours. The promoter region of CG genes is demethylated in a several

volved in X-chromosome inactivation [1, 23, 26-29].

156 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

sine is realised at the carbon-5 position.

malian cell, as well as in the stem cell [13, 37, 38].

tions and epigenetic modifications that may lead to metastasis [39].

In addition to DNA hypermethylation and hypomethylation, a DNA demethylation process was also described. While the active DNA demethylation process takes place in presence of enzymes that catalyzed specific reactions, passive DNA demethylation can occur during replication cycles and operates on DNA methyltransferases [42, 43]. Although the demethy‐ lation process is not fully elucidated, there are many studies showing transient involvement of this process in various types of tumors, especially in advanced stages of their develop‐ ment [44]. The mechanism of DNA demethylation in cancer has been relatively less studied*.* Until just a few years ago, scientists believed that the hypomethylation affects the whole ge‐ nome, randomly [45]. Dysregulations in embryonic normal development and also in normal stem cells development, are generated by many signaling pathways, which can be associat‐ ed with cancer. Pathways signaling implicated in regulation of normal stem cell evolution are also involved in stem cells self-renewal and carcinogenesis. The most common signaling pathways in all processes mentioned are: Wnt, Notch and Sonic hedgehog (Shh) [46, 47]. Other signaling pathways reported to be involved in stem cell maintenance and pluripoten‐ cy are: TGF-beta, MET, MYC, EGF, p53, BMI, etc [5].

It is well-known that Wingless gene encodes the Wnt protein family that control the self– renewal and tumorigenesis processes. Wnt protein is well preserved from Drosophila mela‐ nogaster and has an essential role during normal embryonic development. Wnt protein is a part of a particularly signalling pathway, common to humans and Wnt protein dysregula‐ tion was reported to be involved in the development of tumors. It has been demonstrated that some genes implicated in the Wnt signaling pathway, are inactivated by promoter hy‐ permethylation, generating lung metastasis from primary tumors [48].

The Notch signaling pathway was also suggested to have an important role in stem cell dif‐ ferentiation, proliferation and oncogenesis as well. Scientists have shown that in humans, exist four Notch paralogs (i.e. Notch 1, 2, 3, and 4) and five ligands (i.e. Delta-like 1, 3, 4 and Jagged 1 and 2). Activation of these Notch paralogs are found in stem cell self-renewal but also in many types of cancers [49].

Some epigenetic changes like histone methylation and downregulation of gene expression, which collaborate with the Notch developmental pathway during oncogenesis, were also described [50].

#### **3. Chromatin dynamic and histones modifications**

Chromatin is represented by the mandatory association between nuclear DNA and proteins. Chromatin presents different compression degrees during to cell cycles. It exists in two dif‐ ferent types: euchromatin and heterochromatin.

Euchromatin has more non repetitive DNA with prevailing of guanine and cytosine bases and nonhistonic proteins; it is also less condensed and represents the active and transcrip‐ tional part of the chromatin; it replicates early at the beginning of S faze, being R positive bands from bands marked chromosomes [51].

bution of genetic loci inside it [59]. Polarization of some chromosomes, with their genes lo‐ cated in the interior of the nucleus and their centromeres located at the nuclear periphery, along with all the other data already presented, strongly support the existing relation be‐ tween the chromatin pattern and tumor development, but still many aspects remain to be

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 159

The debate concerning chromatin remodeling as a cause or a consequence of tumorigenesis

Many works indicate that DNA methylation and chromatin remodeling are in reciprocal causal relationship: DNA methylation may cause chromatin modifications and specific chro‐ matin modification may induce DNA methylation [1]. Recent data suggest that chromatin remodeling is a combination between a CIS effect determined by the action of a proximal genetic sequence and a TRANS effect induced by sequence independent complexes, most

Alu sequences are a class of repetitive DNA characterized by a pattern of CG dinucleotides (CpG) repeating every 31-32 bases. They may modulate the nucleosome strength when the CG elements are methylated. Thus, epigenetic nucleosomes within Alu sequences may have

Emerging data are suggesting that since genome regulation might be influenced by nucleo‐ some positioning and their compositional modifications, nucleosomes are regulating the ini‐ tiation of transcription, therefore nucleosomes positioning is leading to cancer or developmental effects [17]. Nucleosomes adopt preferential positions near promoter regions and random positions inside genes [63]. Transcription needs exposed binding sites consist‐ ing in nucleosome free regions at the 5′ and 3′ ends of the genes, so any change in nucleo‐

Nucleosome positioning is also influenced by another protein complex that activates or re‐ presses transcription through biochemical processes, such as octamer transfer, nucleosome remodeling or nucleosome sliding: switch/sucrose nonfermentable (Swi/Snf) complex. It consists in approximately 10 subunits of 2 MDa, with many variants of combinations, first discovered in Saccharomyces cerevisiae [16, 66]. The multiple varieties of Swi/Snf complexes exist in many cell types [67]. Swi/ Snf performs a crucial function in gene regulation and chromosome organization by directly altering the contacts between nucleosomes and DNA [68], using the energy of ATP hydrolysis [69]. The *in vitro* studies revealed that two subunits of the complex, Brg1 or Brm, are able to remodel nucleosomes, with maximal results when subunits BAF155, BAF170 and Ini1 presents a 2:1 stoichiometry relative to Brg1 [70]. The ac‐ tivity of chromatin remodelers appears to be gene specific [71]. The subunits of Swi/ Snf complex seem to have a broad range of functions: BAF155 and BAF170 regulate the protein levels and ensure framing functions for other SWI/SNF subunits [72, 73]; BAF53 is an actin and β-actin related protein signaling, through phosphatidylinositol 4,5- bisphosphate, which binds to Brg1, stimulating the binding to the actin filaments [74-76]; Ini1 is involved in rare and aggressive pediatric cancers [77, 78] as well as in HIV 1 infection [79-81]. The role of SWI/SNF components in cancer stem cells and tumor suppression is still vaguely under‐

likely by ATP dependent nucleosomes remodeling complexes [60, 61].

some positioning at this level might determine gene activation [64, 65].

methylation-dependent regulatory functions [62].

proved.

is still on.

Heterochromatin has more repetitive DNA with predominating adenine and thymine bases and histones; it is very compact, genetically inactive, with late replication. Heterochromatin functions are to stabilize the centromere and the telomeres of the chromosome, playing an important role in meiosis and in cellular differentiation [52]. Heterochromatin is expressed as constitutive or facultative chromatin. Constitutive heterochromatin is constantly found in a condensed form. It doesn't have functional genes and it is made of highly repetitive DNA (satellite DNA).

Facultative heterochromatin is a chromosome region, densely packed and inactive in a par‐ ticular cells, having lost gene expression [53]. Both constitutive and facultative heterochro‐ matin, are regulated by the DNA silencing in the mammalian cells. Constitutive heterochromatin is mandatory transcriptional silenced while facultative heterochromatin is conditionally silenced [53].

Electronic microscope analysis shows a hierarchical system of chromatin fibers with differ‐ ent dimensions, made of DNA, histones and nonhistone proteins. The supramolecular or‐ ganization of DNA has four different levels. The first level is represented by four core histone proteins (H2A, H2B, H3 and H4) which form an octamer wrapped around 1.75 times by 146 base of DNA, making together nucleosomes [54]. Nucleosomes are linked by a short fragment of free DNA (approximately 60 pairs of bases), closed tight by histone H1. The compactness of DNA at this level is 10:1 [54]. Nucleosomes seem to be dynamic structures, since they have to suffer structure modifications during transcription, replication or recom‐ bination of DNA. The second level of chromatin economy is represented by the chromatin fiber of 30 nm, creating a solenoid aspect. A fundamental unit in the interphase, this chro‐ matin plays an important role by putting together regions of linear DNA, stimulating genic interaction. The solenoid has a heterogeneous structure, characterized by an alternation of spiral and not spiraled areas, creating a proper configuration for RNA polymerase action, during transcription. The third level of chromatin organization results from creation of later‐ al loops of 300 nm diameter, attached to a protein nonhistonic matrix. At the beginning of prophase, a matrix will be formed by a 20 times compaction a chromatid, the highest level of chromatin organization [55]. So, the basic DNA will suffer an overall 10000 times compac‐ tion, being able to fit a small place into a nucleus. This conformation offers sterically occlu‐ sion for nucleosomes, which will be there for protected against nucleases cleavage, while the linker DNA doesn't have this kind of protection [56].

Many cancers are associated with translocations which can be explained by mutual rear‐ rangements due to misfit of two unrepaired double stranded breaks, determined by the close proximity of some genetic regions, thus suggesting the dynamic properties of chroma‐ tin [57, 58]. Translocation that characterizes tumorigenesis may depend on the physical dis‐ tance between individual genetic elements. Chromatin is a dynamic structure, with its own mobility that influences either gene regulation (local diffusion of chromatin) or genomic sta‐ bility (global chromatin immobility) [57]. In normal cells, as well as in tumor cells, there are similar nuclear layers, defined as center of nucleus-to-locus distance, with a random distri‐ bution of genetic loci inside it [59]. Polarization of some chromosomes, with their genes lo‐ cated in the interior of the nucleus and their centromeres located at the nuclear periphery, along with all the other data already presented, strongly support the existing relation be‐ tween the chromatin pattern and tumor development, but still many aspects remain to be proved.

tional part of the chromatin; it replicates early at the beginning of S faze, being R positive

Heterochromatin has more repetitive DNA with predominating adenine and thymine bases and histones; it is very compact, genetically inactive, with late replication. Heterochromatin functions are to stabilize the centromere and the telomeres of the chromosome, playing an important role in meiosis and in cellular differentiation [52]. Heterochromatin is expressed as constitutive or facultative chromatin. Constitutive heterochromatin is constantly found in a condensed form. It doesn't have functional genes and it is made of highly repetitive DNA

Facultative heterochromatin is a chromosome region, densely packed and inactive in a par‐ ticular cells, having lost gene expression [53]. Both constitutive and facultative heterochro‐ matin, are regulated by the DNA silencing in the mammalian cells. Constitutive heterochromatin is mandatory transcriptional silenced while facultative heterochromatin is

Electronic microscope analysis shows a hierarchical system of chromatin fibers with differ‐ ent dimensions, made of DNA, histones and nonhistone proteins. The supramolecular or‐ ganization of DNA has four different levels. The first level is represented by four core histone proteins (H2A, H2B, H3 and H4) which form an octamer wrapped around 1.75 times by 146 base of DNA, making together nucleosomes [54]. Nucleosomes are linked by a short fragment of free DNA (approximately 60 pairs of bases), closed tight by histone H1. The compactness of DNA at this level is 10:1 [54]. Nucleosomes seem to be dynamic structures, since they have to suffer structure modifications during transcription, replication or recom‐ bination of DNA. The second level of chromatin economy is represented by the chromatin fiber of 30 nm, creating a solenoid aspect. A fundamental unit in the interphase, this chro‐ matin plays an important role by putting together regions of linear DNA, stimulating genic interaction. The solenoid has a heterogeneous structure, characterized by an alternation of spiral and not spiraled areas, creating a proper configuration for RNA polymerase action, during transcription. The third level of chromatin organization results from creation of later‐ al loops of 300 nm diameter, attached to a protein nonhistonic matrix. At the beginning of prophase, a matrix will be formed by a 20 times compaction a chromatid, the highest level of chromatin organization [55]. So, the basic DNA will suffer an overall 10000 times compac‐ tion, being able to fit a small place into a nucleus. This conformation offers sterically occlu‐ sion for nucleosomes, which will be there for protected against nucleases cleavage, while the

Many cancers are associated with translocations which can be explained by mutual rear‐ rangements due to misfit of two unrepaired double stranded breaks, determined by the close proximity of some genetic regions, thus suggesting the dynamic properties of chroma‐ tin [57, 58]. Translocation that characterizes tumorigenesis may depend on the physical dis‐ tance between individual genetic elements. Chromatin is a dynamic structure, with its own mobility that influences either gene regulation (local diffusion of chromatin) or genomic sta‐ bility (global chromatin immobility) [57]. In normal cells, as well as in tumor cells, there are similar nuclear layers, defined as center of nucleus-to-locus distance, with a random distri‐

bands from bands marked chromosomes [51].

158 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

linker DNA doesn't have this kind of protection [56].

(satellite DNA).

conditionally silenced [53].

The debate concerning chromatin remodeling as a cause or a consequence of tumorigenesis is still on.

Many works indicate that DNA methylation and chromatin remodeling are in reciprocal causal relationship: DNA methylation may cause chromatin modifications and specific chro‐ matin modification may induce DNA methylation [1]. Recent data suggest that chromatin remodeling is a combination between a CIS effect determined by the action of a proximal genetic sequence and a TRANS effect induced by sequence independent complexes, most likely by ATP dependent nucleosomes remodeling complexes [60, 61].

Alu sequences are a class of repetitive DNA characterized by a pattern of CG dinucleotides (CpG) repeating every 31-32 bases. They may modulate the nucleosome strength when the CG elements are methylated. Thus, epigenetic nucleosomes within Alu sequences may have methylation-dependent regulatory functions [62].

Emerging data are suggesting that since genome regulation might be influenced by nucleo‐ some positioning and their compositional modifications, nucleosomes are regulating the ini‐ tiation of transcription, therefore nucleosomes positioning is leading to cancer or developmental effects [17]. Nucleosomes adopt preferential positions near promoter regions and random positions inside genes [63]. Transcription needs exposed binding sites consist‐ ing in nucleosome free regions at the 5′ and 3′ ends of the genes, so any change in nucleo‐ some positioning at this level might determine gene activation [64, 65].

Nucleosome positioning is also influenced by another protein complex that activates or re‐ presses transcription through biochemical processes, such as octamer transfer, nucleosome remodeling or nucleosome sliding: switch/sucrose nonfermentable (Swi/Snf) complex. It consists in approximately 10 subunits of 2 MDa, with many variants of combinations, first discovered in Saccharomyces cerevisiae [16, 66]. The multiple varieties of Swi/Snf complexes exist in many cell types [67]. Swi/ Snf performs a crucial function in gene regulation and chromosome organization by directly altering the contacts between nucleosomes and DNA [68], using the energy of ATP hydrolysis [69]. The *in vitro* studies revealed that two subunits of the complex, Brg1 or Brm, are able to remodel nucleosomes, with maximal results when subunits BAF155, BAF170 and Ini1 presents a 2:1 stoichiometry relative to Brg1 [70]. The ac‐ tivity of chromatin remodelers appears to be gene specific [71]. The subunits of Swi/ Snf complex seem to have a broad range of functions: BAF155 and BAF170 regulate the protein levels and ensure framing functions for other SWI/SNF subunits [72, 73]; BAF53 is an actin and β-actin related protein signaling, through phosphatidylinositol 4,5- bisphosphate, which binds to Brg1, stimulating the binding to the actin filaments [74-76]; Ini1 is involved in rare and aggressive pediatric cancers [77, 78] as well as in HIV 1 infection [79-81]. The role of SWI/SNF components in cancer stem cells and tumor suppression is still vaguely under‐ stood, but their transcriptional pathways are already described, including the cell cycle and p53 signaling [82], insulin signaling [83], and TGFb signaling [84], or signaling through sev‐ eral different nuclear hormone receptors [85]. The biological roles of Swi/Snf components and their involvement in human disease remain to be completed. It is also known that loss of Snf2h impairs embryonic development and differentiation [86] and contributes to tumor development [87]. These tumors are also characterized by polyploidy and chromosomal in‐ stability [88]. Since Swi/Snf complex plays also an important role in DNA double strand brake repair, alteration of its function may lead to genomic instability [89]. Critical subunits of Swi/Snf complex miss or are disrupted in approximately 17% of all human adenocarcino‐ mas [90]. Another tumor suppressor gene is Ikaros, a molecule that plays a central role in lymphocyte development through its association with chromatin remodeling complexes [91]. Fusion protein BCR-ABL from preB lymphoblastic leukemia mediates an aberrant splicing of Ikaros, with consequences on cell differentiation [92].

(HAT) and lysine 4 methylation in the histone H3 (H3K4me) by histone methyltransferase (HMT), are generally correlated with active transcription of chromatin. In contrast, methyla‐ tion of lysine 9 and lysine 27 (H3K9, H3K27me) in the histone 3 have been reported as mark‐

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 161

During the last years, a major challenge in cancer biology is to elucidate how the histone

Histones methylation is a post translational modification that occurs at the lysine residues and is considered a reversible process [103]. Transcriptional activation or repression, corre‐ lates with different degrees of methylation of histones. The binding of one to three methyl groups at each lysine amino acid in the histone structure, give rise to unmethylated, mono‐ methylated, dimethylated and trimethylated degrees of methylation [104, 105]. The monomethylation state of histone has been reported to be associated with an open chromatin structure that lead to transcriptional activation. In contrast, the trimethylation state was shown to be associated with a condensed chromatin structure, which in turn inhibits tran‐

Some important exceptions from this rule have been reported by Strahl BD et al., they showed that H3K4 histone methylation state (mono-, di-, or tri-methylated level) is invaria‐ bly associated with active chromatin, while H3K9 trimethylation can be connected to both transcriptionally active and inactive chromatin [107]. To explain this exception from the general rule, Vakoc et al., described a mechanism by which an association between meH3K9 with RNA polymerase II complexes induces chromatin modification and tran‐ scriptional activation [108]. However, it is not fully understood why these markers differs

The binding of the methyl group of each lysine 4, 36 or 79 in the histone 3 (H3K4, H3K36, H3K79) and H4 (k20, H2BK5) induces trnascriptional activation. In contrast, the binding of three methyl group of lysine 9, 27 in the histone 3 (H3K9, H33K27) AND h4k20 was show to be associated with inhibiton of transcription [109, 110]. The histone modifications are arising from the action of enzymes which are responsible for methylation/demethylation activity in the pattern of histone H3 and H4. The enzymes involved in histone modifications are his‐ tone acetyltransferases (HATs) and histone deacetylases (HDACs), histona methyltransfer‐ ases (HMTs) and histone demethylases (HDMs). These enzymes add or remove acetyl or methyl groups, respectively [111, 112]. Several enzymes, like histone methyltransferase (HMTs), histone demethylases (HDMs) and histone deacetylases (HDACs), are connected

with each other to create a strong link between chromatin state and transcription.

to be associated with changes in H3K9 and H3K27 methylation patterns [113].

In addition to changes in histone acetylation, widespread changes in histone methylation patterns are described in cancer. Accordingly, in cancer, aberrant gene silencing was shown

A recent analysis in the context of histone modifications in cancer, illustrates different sce‐ narios such as histone methylation and its consequences, describing the role of histone methyltransferases (HMTs) and histone demethylation (HDMS) by adding or removing a

ers in transcriptionally silenced-chromatin [21, 22].

modifications in stem cells, influence carcinogenesis.

scription [106].

from the general rule.

Radiations have the ability to paradoxically induce or cure tumors. It seems that chromatin structure might be influenced by UV and gamma radiation. To study the changes in chro‐ matin pattern under irradiation conditions, Fluorescence In Situ Hybridization (FISH), com‐ bined with high-resolution confocal microscopy has been used [93, 94]. FISH studies were performed in leukemia cells for tumor suppressor gene TP53, revealing that TP53 genes are getting closer to each other, as well with the nuclear center within 2 hours of exposure to gamma-radiation, returning during the following 2 hours to its pre-irradiation conditions [95, 96]. There is increasing evidence that CSCs have a higher intrinsic radioresistance than non-CSC tumor cells [97], explaining the difference of CSCs and non-CSC in their response to cancer therapy.

Such repressive complex for chromatin is "Nucleosome Remodeling and Histone Deacety‐ lase" (NuRD) [98], suggested to play a role in acute promyelocytic leukemia (APL) [99]. Hu‐ man APL is characterized by PML-RARa translocation, which represses gene transcription through several distinct epigenetic mechanisms: DNA methylation, chromatin compaction, heterochromatinization, histone deacetylation, histone modification. NuRD complex is strongly implicated in the epigenetic silencing, by PML-RARa. Earlier findings regarding carcinogenesis, such as the combination of both genetic and epigenetic factors, were con‐ firmed. in this case, PML-RARa oncogenic fusion protein recruiting, induces DNA hyperme‐ thylation [100] and result in blocking of hematopoietic differentiation [101].

Covalent modification of histones is an important mechanism, involved in the epigenetic processes. Five types of histones are known to be involved in chromatin building: H1/H5, H2A, H2B, H3, and H4 [15, 102]. In their structure, histones have three distinct domains: a central globular conserved domain and two terminal domains; one short N-terminal tail and one longer C-terminal tail [54].

Generally, histone modifications affect gene transcription, DNA replication and DNA repair mechanisms. The post-translational modifications of the N-terminal tail domains include: methylation, acetylation, phosphorylation, citrullination, ADP-ribosylation, sumoylation and ubiquitination [18-20]. The most studied of these modifications are methylation and ace‐ tylation. Lysine residues acetylation in the histone H3 and H4 by histone acetyltransferase (HAT) and lysine 4 methylation in the histone H3 (H3K4me) by histone methyltransferase (HMT), are generally correlated with active transcription of chromatin. In contrast, methyla‐ tion of lysine 9 and lysine 27 (H3K9, H3K27me) in the histone 3 have been reported as mark‐ ers in transcriptionally silenced-chromatin [21, 22].

stood, but their transcriptional pathways are already described, including the cell cycle and p53 signaling [82], insulin signaling [83], and TGFb signaling [84], or signaling through sev‐ eral different nuclear hormone receptors [85]. The biological roles of Swi/Snf components and their involvement in human disease remain to be completed. It is also known that loss of Snf2h impairs embryonic development and differentiation [86] and contributes to tumor development [87]. These tumors are also characterized by polyploidy and chromosomal in‐ stability [88]. Since Swi/Snf complex plays also an important role in DNA double strand brake repair, alteration of its function may lead to genomic instability [89]. Critical subunits of Swi/Snf complex miss or are disrupted in approximately 17% of all human adenocarcino‐ mas [90]. Another tumor suppressor gene is Ikaros, a molecule that plays a central role in lymphocyte development through its association with chromatin remodeling complexes [91]. Fusion protein BCR-ABL from preB lymphoblastic leukemia mediates an aberrant

Radiations have the ability to paradoxically induce or cure tumors. It seems that chromatin structure might be influenced by UV and gamma radiation. To study the changes in chro‐ matin pattern under irradiation conditions, Fluorescence In Situ Hybridization (FISH), com‐ bined with high-resolution confocal microscopy has been used [93, 94]. FISH studies were performed in leukemia cells for tumor suppressor gene TP53, revealing that TP53 genes are getting closer to each other, as well with the nuclear center within 2 hours of exposure to gamma-radiation, returning during the following 2 hours to its pre-irradiation conditions [95, 96]. There is increasing evidence that CSCs have a higher intrinsic radioresistance than non-CSC tumor cells [97], explaining the difference of CSCs and non-CSC in their response

Such repressive complex for chromatin is "Nucleosome Remodeling and Histone Deacety‐ lase" (NuRD) [98], suggested to play a role in acute promyelocytic leukemia (APL) [99]. Hu‐ man APL is characterized by PML-RARa translocation, which represses gene transcription through several distinct epigenetic mechanisms: DNA methylation, chromatin compaction, heterochromatinization, histone deacetylation, histone modification. NuRD complex is strongly implicated in the epigenetic silencing, by PML-RARa. Earlier findings regarding carcinogenesis, such as the combination of both genetic and epigenetic factors, were con‐ firmed. in this case, PML-RARa oncogenic fusion protein recruiting, induces DNA hyperme‐

Covalent modification of histones is an important mechanism, involved in the epigenetic processes. Five types of histones are known to be involved in chromatin building: H1/H5, H2A, H2B, H3, and H4 [15, 102]. In their structure, histones have three distinct domains: a central globular conserved domain and two terminal domains; one short N-terminal tail and

Generally, histone modifications affect gene transcription, DNA replication and DNA repair mechanisms. The post-translational modifications of the N-terminal tail domains include: methylation, acetylation, phosphorylation, citrullination, ADP-ribosylation, sumoylation and ubiquitination [18-20]. The most studied of these modifications are methylation and ace‐ tylation. Lysine residues acetylation in the histone H3 and H4 by histone acetyltransferase

thylation [100] and result in blocking of hematopoietic differentiation [101].

splicing of Ikaros, with consequences on cell differentiation [92].

160 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

to cancer therapy.

one longer C-terminal tail [54].

During the last years, a major challenge in cancer biology is to elucidate how the histone modifications in stem cells, influence carcinogenesis.

Histones methylation is a post translational modification that occurs at the lysine residues and is considered a reversible process [103]. Transcriptional activation or repression, corre‐ lates with different degrees of methylation of histones. The binding of one to three methyl groups at each lysine amino acid in the histone structure, give rise to unmethylated, mono‐ methylated, dimethylated and trimethylated degrees of methylation [104, 105]. The monomethylation state of histone has been reported to be associated with an open chromatin structure that lead to transcriptional activation. In contrast, the trimethylation state was shown to be associated with a condensed chromatin structure, which in turn inhibits tran‐ scription [106].

Some important exceptions from this rule have been reported by Strahl BD et al., they showed that H3K4 histone methylation state (mono-, di-, or tri-methylated level) is invaria‐ bly associated with active chromatin, while H3K9 trimethylation can be connected to both transcriptionally active and inactive chromatin [107]. To explain this exception from the general rule, Vakoc et al., described a mechanism by which an association between meH3K9 with RNA polymerase II complexes induces chromatin modification and tran‐ scriptional activation [108]. However, it is not fully understood why these markers differs from the general rule.

The binding of the methyl group of each lysine 4, 36 or 79 in the histone 3 (H3K4, H3K36, H3K79) and H4 (k20, H2BK5) induces trnascriptional activation. In contrast, the binding of three methyl group of lysine 9, 27 in the histone 3 (H3K9, H33K27) AND h4k20 was show to be associated with inhibiton of transcription [109, 110]. The histone modifications are arising from the action of enzymes which are responsible for methylation/demethylation activity in the pattern of histone H3 and H4. The enzymes involved in histone modifications are his‐ tone acetyltransferases (HATs) and histone deacetylases (HDACs), histona methyltransfer‐ ases (HMTs) and histone demethylases (HDMs). These enzymes add or remove acetyl or methyl groups, respectively [111, 112]. Several enzymes, like histone methyltransferase (HMTs), histone demethylases (HDMs) and histone deacetylases (HDACs), are connected with each other to create a strong link between chromatin state and transcription.

In addition to changes in histone acetylation, widespread changes in histone methylation patterns are described in cancer. Accordingly, in cancer, aberrant gene silencing was shown to be associated with changes in H3K9 and H3K27 methylation patterns [113].

A recent analysis in the context of histone modifications in cancer, illustrates different sce‐ narios such as histone methylation and its consequences, describing the role of histone methyltransferases (HMTs) and histone demethylation (HDMS) by adding or removing a methyl group. It has been reported that the level of transcriptional activation is largely maintained by HMTs and HDMs which are involved in the histone methylation [103].

two different mechanisms: splitting and subsequent degradation of targeted RNAm or in‐ hibiting translation, both determining the stop or stimulation of cell reproduction [126-130].

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 163

The study on Caenorhabditis elegans (C. elegans) has permitted the clonation of first miR‐ NA, lin-4 and let-7 and their targets [133, 134]. This study discovered that the gene lin-14 was able to transcribe a precursor that matured to a 22 nucleotide mature RNA, which con‐ tained sequences partially complementary to multiple sequences in the 3' UTR of the lin-14 mRNA, ensuring inhibition of translation of lin-14 mRNA. In addition, in 2000, along with the discovery that gene let-7 repressed the genes lin-41, lin-14, lin-28, lin-42 and daf12 mRNA during transition in developmental stages in C. Elegans, it was also established that

More than 700 miRNAs have been identified in humans and over 800 more are predicted to exist. These molecules have an important role in cellular physiological processes (e.g. cell cy‐ cle, cell proliferation, apoptosis, cell differentiation and development), by implication in

In human embryonic stem cells and the differentiated embryonic bodies, over 100 miRNAs have been already described [101]. The self-renewal and pluripotency of embryonic stem cells are regulated by an array of protein-coding genes in a regulatory circuitry [135], which includes OCT4, SOX2, and KLF4 genes. Extensive studies have indicated the importance of OCT4 in self-renewal and pluripotency of embryonic stem cells [136, 137]. Multipotent cell lineages in early mouse development, have also been reported to be dependent on SOX2 function [138, 139] in the process of embryonic stem cells self-renewal and pluripotency. The miRNA genes are also connected to the transcriptional regulatory circuitry of embryonic

The three key proteins of pluripotent cells, Oct4, Nanog and SOX2, and TCF3 were found in the promoters of miRNA specific stem cells, but also in promoters of miRNA, which con‐ trols cell proliferation (mir 92 si let7g) and differentiation (e.g., mir-9 or mir-124a for neutral line). OCT4 was reported to bind and repress miR-145 promoter in human embryonic stem cells. On the other hand, inhibition of Oct 4 increases the activity of these miRNAs that in

miRNAs, occasionally causes DNA methylation of promoter sites and can regulate other epigenetic mechanisms. An altered miRNA gene methylation patterns in human cancers was reported to sustain in tumorigenesis. Half of these genes are associated with CpG is‐ lands and several studies indicated that miRNA gene methylation was often detectable, both in normal and malignant cells. Recent works have identified many types of miR‐ NA, which allow cancer cells to multiply indefinitely by avoiding natural cellular aging mechanisms, thus suggesting a close relation between cancer development and miRNA

There are several mechanisms which may lead to modification of mi RNA in cancer [142-145].

However, the entire mechanism of miRNA is not yet fully understood [131, 132].

non-coding RNA identified in 1993, was part of a wider phenomenon [133].

stem cells [140] and are overexpressed in their differentiating processes.

turn inhibit the stem cell renewal.

expression [141].

gene regulation; miRNA has been found to control about 30% of all human genes.

The histone lysine methyltransferase (HMT) that is responsible for the histone methylation, has a catalytically active site known as SET domain, which is formed by de 130 amino acid sequence. The major function of the SET domain is to modulate gene activity [114].

The binding of the methyl group at several lysine sites in histone H3 (H3K9, H3K27, H3K36, H3K79) and loss of acetylated H4 lysine 16 and H4 lysine 20 trimethylation have been re‐ ported to be associated with changes that occur during tumorigenesis[115]. The enzymes HDACs and HATs have been suggested to be responsible for these changes and are com‐ monly found to be altered in various forms of cancer [116].

Various observations suggest the presence of a novel chromatin pattern in embryonic stem cell, which consists of lysine 27 and lysine 4 tri-methylation superposition, termed "bivalent domains" [117].

The bivalent domains have been analysed by the genome mapping of histone methylation profiles in embryonic stem cell and was reported to include both active and repressive chro‐ matin marks. Developmentally, the "bivalent domains" is responsible for maintaining epi‐ genomic plasticity, enabling embryonic stem cells to regulate gene expression [117]. Bivalency is lost during stem cell differentiation, allowing epigenetic plasticity and lineage commitment. Epigenetic plasticity in association with bivalent gene promoters is suggested to induce a transcriptionally repressive and permissive histone mark in embryonic stem cells [117, 118].

In cancer, bivalency has been suggested to stigmatize specific genes for DNA methylation, inducing aberrant reprogramming [119-121]. In analogy with embrionic stem cells, bivalent gene promoters were reported to be DNA-methylated in cancer cells, suggesting the prove‐ nience of cancer cells from embryonic stem cells [122]. In absence of DNA methylation, the repressive H3K27 trimethylation mark was also demonstrated to induce gene silencing in cancer cells.

Chromatin regulating complexes are commonly observed in cancer, and is hypothesized to involve multiple mechanisms, including DNA methylation and Polycomb repressive com‐ plexes (PRCs). Chromatin regulating complexes including two families of Polycomb repres‐ sive complexes (PRC1 and PRC1), mediate trimethylation on H3K27 in cancer cells [123, 124]. PRC2 complex has also been reported to intermediate H3K27 trimethylation in embry‐ onic stem cell [125].

#### **4. miRNA and DNA methylation in cancer stem cells**

MicroRNAs (miRNAs) was first discovered in 1993 by Victor Ambros, Rosalind Lee and Rhonda Feinbaum. The recent definition of miRNA is: small non-coding RNA molecules (21-24 nucleotides long), implicated in posttranscriptional gene expression, regulation by two different mechanisms: splitting and subsequent degradation of targeted RNAm or in‐ hibiting translation, both determining the stop or stimulation of cell reproduction [126-130].

However, the entire mechanism of miRNA is not yet fully understood [131, 132].

methyl group. It has been reported that the level of transcriptional activation is largely maintained by HMTs and HDMs which are involved in the histone methylation [103].

The histone lysine methyltransferase (HMT) that is responsible for the histone methylation, has a catalytically active site known as SET domain, which is formed by de 130 amino acid

The binding of the methyl group at several lysine sites in histone H3 (H3K9, H3K27, H3K36, H3K79) and loss of acetylated H4 lysine 16 and H4 lysine 20 trimethylation have been re‐ ported to be associated with changes that occur during tumorigenesis[115]. The enzymes HDACs and HATs have been suggested to be responsible for these changes and are com‐

Various observations suggest the presence of a novel chromatin pattern in embryonic stem cell, which consists of lysine 27 and lysine 4 tri-methylation superposition, termed "bivalent

The bivalent domains have been analysed by the genome mapping of histone methylation profiles in embryonic stem cell and was reported to include both active and repressive chro‐ matin marks. Developmentally, the "bivalent domains" is responsible for maintaining epi‐ genomic plasticity, enabling embryonic stem cells to regulate gene expression [117]. Bivalency is lost during stem cell differentiation, allowing epigenetic plasticity and lineage commitment. Epigenetic plasticity in association with bivalent gene promoters is suggested to induce a transcriptionally repressive and permissive histone mark in embryonic stem

In cancer, bivalency has been suggested to stigmatize specific genes for DNA methylation, inducing aberrant reprogramming [119-121]. In analogy with embrionic stem cells, bivalent gene promoters were reported to be DNA-methylated in cancer cells, suggesting the prove‐ nience of cancer cells from embryonic stem cells [122]. In absence of DNA methylation, the repressive H3K27 trimethylation mark was also demonstrated to induce gene silencing in

Chromatin regulating complexes are commonly observed in cancer, and is hypothesized to involve multiple mechanisms, including DNA methylation and Polycomb repressive com‐ plexes (PRCs). Chromatin regulating complexes including two families of Polycomb repres‐ sive complexes (PRC1 and PRC1), mediate trimethylation on H3K27 in cancer cells [123, 124]. PRC2 complex has also been reported to intermediate H3K27 trimethylation in embry‐

MicroRNAs (miRNAs) was first discovered in 1993 by Victor Ambros, Rosalind Lee and Rhonda Feinbaum. The recent definition of miRNA is: small non-coding RNA molecules (21-24 nucleotides long), implicated in posttranscriptional gene expression, regulation by

**4. miRNA and DNA methylation in cancer stem cells**

sequence. The major function of the SET domain is to modulate gene activity [114].

monly found to be altered in various forms of cancer [116].

162 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

domains" [117].

cells [117, 118].

cancer cells.

onic stem cell [125].

The study on Caenorhabditis elegans (C. elegans) has permitted the clonation of first miR‐ NA, lin-4 and let-7 and their targets [133, 134]. This study discovered that the gene lin-14 was able to transcribe a precursor that matured to a 22 nucleotide mature RNA, which con‐ tained sequences partially complementary to multiple sequences in the 3' UTR of the lin-14 mRNA, ensuring inhibition of translation of lin-14 mRNA. In addition, in 2000, along with the discovery that gene let-7 repressed the genes lin-41, lin-14, lin-28, lin-42 and daf12 mRNA during transition in developmental stages in C. Elegans, it was also established that non-coding RNA identified in 1993, was part of a wider phenomenon [133].

More than 700 miRNAs have been identified in humans and over 800 more are predicted to exist. These molecules have an important role in cellular physiological processes (e.g. cell cy‐ cle, cell proliferation, apoptosis, cell differentiation and development), by implication in gene regulation; miRNA has been found to control about 30% of all human genes.

In human embryonic stem cells and the differentiated embryonic bodies, over 100 miRNAs have been already described [101]. The self-renewal and pluripotency of embryonic stem cells are regulated by an array of protein-coding genes in a regulatory circuitry [135], which includes OCT4, SOX2, and KLF4 genes. Extensive studies have indicated the importance of OCT4 in self-renewal and pluripotency of embryonic stem cells [136, 137]. Multipotent cell lineages in early mouse development, have also been reported to be dependent on SOX2 function [138, 139] in the process of embryonic stem cells self-renewal and pluripotency. The miRNA genes are also connected to the transcriptional regulatory circuitry of embryonic stem cells [140] and are overexpressed in their differentiating processes.

The three key proteins of pluripotent cells, Oct4, Nanog and SOX2, and TCF3 were found in the promoters of miRNA specific stem cells, but also in promoters of miRNA, which con‐ trols cell proliferation (mir 92 si let7g) and differentiation (e.g., mir-9 or mir-124a for neutral line). OCT4 was reported to bind and repress miR-145 promoter in human embryonic stem cells. On the other hand, inhibition of Oct 4 increases the activity of these miRNAs that in turn inhibit the stem cell renewal.

miRNAs, occasionally causes DNA methylation of promoter sites and can regulate other epigenetic mechanisms. An altered miRNA gene methylation patterns in human cancers was reported to sustain in tumorigenesis. Half of these genes are associated with CpG is‐ lands and several studies indicated that miRNA gene methylation was often detectable, both in normal and malignant cells. Recent works have identified many types of miR‐ NA, which allow cancer cells to multiply indefinitely by avoiding natural cellular aging mechanisms, thus suggesting a close relation between cancer development and miRNA expression [141].

There are several mechanisms which may lead to modification of mi RNA in cancer [142-145].

**•** *Chromosomal changes -* quantitative gene changes have been identified in approximately 283 miRNA, determined by either loosing heterozigotism by the action of a suppressive gene or by amplification of a chromosomal region of an oncogene either by chromosomal ruptures or translocations [146];

py. Epigenetic targeting may be realized by two classes of substances with antitumor effect in malignancies: the hypomethylating agents and histone deacetylase inhibitors [160].

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 165

DNA methylation is the most studied epigenetic marker. Abnormal DNA methylation of several regulatory genes is usually associated with cancer. The methylation process is rever‐ sible, therefore the reactivation of silenced genes can be realized using substances with hy‐

The new development cancer therapies are based on molecules that can inhibit the classes of DNA methyltransferases (DNMT), histone deacetylases (HDACs), histone acetyltransferases (HATs) and new substances that target chromatin and nucleosome remodeling proteins.

As mentioned before, DNMTs are enzymes that catalyze the reaction between methyl groups and pyrimidine base cytosine. The methyl group donor is S-adenosyl-L-methionine

The DNA hypomethylating agents are divided into two categories*:* nucleoside analogs drugs and non-nucleoside analogs drugs. The first description substances are 5-azacytidine (azacitidine, VidazaTM) and 5-aza-2'-deoxycytidine (decitabine, DacogenTM) that have the

**Figure 2.** Nucleoside analog drugs. Their structure allows incorporation into the DNA and subsequent hypomethilation.

Other substances from this group are: 1-β-D-arabinosyl-5-azacytidine (fazarabine) [166], di‐ hydro-5-azacytidine (DHAC), 5-fluoro-2'-deoxycytidine (FCDR) and zebularine [163]. Incor‐ poration into the DNA structure of nucleoside analogs is facilitated by their similar chemical

5-azacytidine is used as single-agent therapy or in combination with other therapies in treatment of myelodysplastic syndromes (MDS), acute myeloid leukemias (AML) and solid tumor. As associated substances are utilized valproic acid, cytarabine, entinostat,

5-aza-2'-deoxycytidine (DAC) has benefited as monotherapy in myelodysplastic syndromes, chronic myelomonocytic leukemia (CMML) and has been FDA approved on May 2006. The

DNMT inhibitors (DNMTi) can be natural or synthetic compounds [148, 162].

(SAM), which is the active form of amino acid methionine.

most powerful effect from nucleoside analogs drugs [1, 163-165].

**5.1. Targeting DNA methylation**

pomethylating activity [161].

structure.

etanercept etc [137, 164].


Oncogenic activity of miRNA, initially determined for mir-17-92 and mir-155, was further sustained by the discovery of other potentially oncogenic miRNA [150]. Therefore it is logi‐ cal that this classification of miRNA in oncogenes or tumor supressor genes may facilitate the identification of different tissues where they are expressed [151].

Embryonic stem cells gene expression of Oct4, Sox2, Klf4, and Nanog was observed in high‐ ly aggressive human tumors [152]. It has been reported that miR-200 known to mediate tran‐ scriptional repression, also play an important role in both cancer stem cells andembryonic stem cells [153, 154].

Several miRNAs have been reported to be overexpressed in human cancer. The mir-17-92 polycistron (cluster) is overexpressed in B-cell lymphoma [155, 156] and in testicular germ cell tumors miR-372 and miR-373 were identified as possible oncogenes [157].

Another theory supports the idea that some cancers such as Kaposi `s sarcoma, were in‐ duced by viral oncogenic miRNA [158]. It is clear that discovery of miRNA involvement in cancer stem cells function will be a crucial step in elucidating the process of oncogen‐ esis [159].

#### **5. Epigenetic targeting in cancer stem cells**

Recently, several epigenetic drugs targeting epigenetic mechanisms have been tested *in vivo* and *in vitro*. The epigenetic mechanisms comprise modifications of histones and DNA meth‐ ylation. Histone modifications includes several post translational modification of the: meth‐ ylation, acetylation, phosphorylation, ubiquitination, sumoylation; commonly found in tumor cells. Thus, epigenetic modifications targeting is an important event in cancer thera‐ py. Epigenetic targeting may be realized by two classes of substances with antitumor effect in malignancies: the hypomethylating agents and histone deacetylase inhibitors [160].

#### **5.1. Targeting DNA methylation**

**•** *Chromosomal changes -* quantitative gene changes have been identified in approximately 283 miRNA, determined by either loosing heterozigotism by the action of a suppressive gene or by amplification of a chromosomal region of an oncogene either by chromosomal

**•** *miRNA biosynthesis abnormalities*, mainly represented by gene amplification for proteins as Drosha (implicated in miRNA maturation process) or Ago2 (responsible for the interac‐

**•** *Epigenetic changes-* recent data suggest the implication of DNA methylation in the disor‐ der of miRNA expression. Gene analysis for miRNA established that these genes are usu‐ ally associate with CpG islands and thus represent candidate targets of the DNA methylation machinery. A high level of miRNA genes methylation exists both in normal and malignant cells. Epigenetic changes of chromatin, for instance histone deacetylation,

**•** *miRNA as oncogenes or tumor suppressors -* miRNA always acts as negative regulator of gene expression. In cancer, miRNA were classifies as miRNA with oncogenic effect (onco‐

Oncogenic activity of miRNA, initially determined for mir-17-92 and mir-155, was further sustained by the discovery of other potentially oncogenic miRNA [150]. Therefore it is logi‐ cal that this classification of miRNA in oncogenes or tumor supressor genes may facilitate

Embryonic stem cells gene expression of Oct4, Sox2, Klf4, and Nanog was observed in high‐ ly aggressive human tumors [152]. It has been reported that miR-200 known to mediate tran‐ scriptional repression, also play an important role in both cancer stem cells andembryonic

Several miRNAs have been reported to be overexpressed in human cancer. The mir-17-92 polycistron (cluster) is overexpressed in B-cell lymphoma [155, 156] and in testicular germ

Another theory supports the idea that some cancers such as Kaposi `s sarcoma, were in‐ duced by viral oncogenic miRNA [158]. It is clear that discovery of miRNA involvement in cancer stem cells function will be a crucial step in elucidating the process of oncogen‐

Recently, several epigenetic drugs targeting epigenetic mechanisms have been tested *in vivo* and *in vitro*. The epigenetic mechanisms comprise modifications of histones and DNA meth‐ ylation. Histone modifications includes several post translational modification of the: meth‐ ylation, acetylation, phosphorylation, ubiquitination, sumoylation; commonly found in tumor cells. Thus, epigenetic modifications targeting is an important event in cancer thera‐

cell tumors miR-372 and miR-373 were identified as possible oncogenes [157].

**5. Epigenetic targeting in cancer stem cells**

cause important alteration of miRNA expression as well [141];

mirs) and miRNA with suppressor effect (supressor mirs) [147-149].

the identification of different tissues where they are expressed [151].

ruptures or translocations [146];

164 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

tion with target messengers);

stem cells [153, 154].

esis [159].

DNA methylation is the most studied epigenetic marker. Abnormal DNA methylation of several regulatory genes is usually associated with cancer. The methylation process is rever‐ sible, therefore the reactivation of silenced genes can be realized using substances with hy‐ pomethylating activity [161].

The new development cancer therapies are based on molecules that can inhibit the classes of DNA methyltransferases (DNMT), histone deacetylases (HDACs), histone acetyltransferases (HATs) and new substances that target chromatin and nucleosome remodeling proteins. DNMT inhibitors (DNMTi) can be natural or synthetic compounds [148, 162].

As mentioned before, DNMTs are enzymes that catalyze the reaction between methyl groups and pyrimidine base cytosine. The methyl group donor is S-adenosyl-L-methionine (SAM), which is the active form of amino acid methionine.

The DNA hypomethylating agents are divided into two categories*:* nucleoside analogs drugs and non-nucleoside analogs drugs. The first description substances are 5-azacytidine (azacitidine, VidazaTM) and 5-aza-2'-deoxycytidine (decitabine, DacogenTM) that have the most powerful effect from nucleoside analogs drugs [1, 163-165].

**Figure 2.** Nucleoside analog drugs. Their structure allows incorporation into the DNA and subsequent hypomethilation.

Other substances from this group are: 1-β-D-arabinosyl-5-azacytidine (fazarabine) [166], di‐ hydro-5-azacytidine (DHAC), 5-fluoro-2'-deoxycytidine (FCDR) and zebularine [163]. Incor‐ poration into the DNA structure of nucleoside analogs is facilitated by their similar chemical structure.

5-azacytidine is used as single-agent therapy or in combination with other therapies in treatment of myelodysplastic syndromes (MDS), acute myeloid leukemias (AML) and solid tumor. As associated substances are utilized valproic acid, cytarabine, entinostat, etanercept etc [137, 164].

5-aza-2'-deoxycytidine (DAC) has benefited as monotherapy in myelodysplastic syndromes, chronic myelomonocytic leukemia (CMML) and has been FDA approved on May 2006. The drug has been associated with: carboplatin useful in solid tumors treatment [167], valproic acid in acute myeloid leukemias and advanced leukemia [168, 169], imatinib mesylate in chronic myelogenous leukemia (CML) [170] and IL-2 in metastatic melanoma, renal carcinoma [171].

Zebularine (2-pyrimidone-1-β- D-riboside) is other nucleoside analog with hypomethylation activity [172] and also implicated in tumor gene expression [173].

**Figure 3.** Structure of Zebularine. Zebularine is another nucleoside analog drug, with hypomethilation effect.

There are recent studies about another two molecules: NPEOC-DAC and SGI 110 (S110). NPEOC-DAC is the result of chemical reaction between azacytosine molecule and 2-(p-ni‐ trophenyl) ethoxycarbonyl, with reported effect on DNA methyltransferases inhibition. By‐ un et al. demonstrated that NPEOC-DAC inhibited DNA methylation in two cell lines of liver cancer. The authors, also showed that SGI 110 (S110) has a pronounced effect on DNA methylation inhibition [174].

The non-nucleoside analogs category contains compounds with hypomethylation effect. This group contains hydralazine (the widely known as vasodilatator), procainamide (antiarhythmic), RG108 and SGI-1027.

**Figure 4.** HDAC inhibitors. There are four classes of curently known HDAC inhibitors: short-chain fatty acids, hydroxa‐

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 167

Exemples of short-chain fatty acids are: sodium n-butyrate, sodium phenylacetate, phenyl‐ butyrate, valproate, substances that in millimolar concentrations are involved in inhibition the growth of some carcinomas but their mechanism of action is not fully understood [163,

One of the most studied agent from class of small fatty acids, is valproic acid (VPA), an anti‐ epileptic drug reported to target histone deacetylase. Numerous research studies *in vitro* demonstrated that VPA was implicated in hyperacetylation of histones H3 and H4 and also *in vivo* tests confirmed the drug inhibiting action of HDACs. VPA antitumor activity was de‐ manstrated by: cell growth inhibition, apoptosis inducing, antimetastatic and antiangiogene‐

mic acids, cyclic peptides, benzamides, with a great potential use as detection and prognosis markers.

sis effect, etc. These benefits lead to FDA approving of VPA [14, 180].

177-179].

Physiologically, acetylation of chromatin is realized by specific enzymes - histone deace‐ tylases and acetyltransferases. A possible change in their normal function can promote tumors.

#### **5.2. Targeting histone modification**

At first glance, HDACs are enzymes that play a role in elimination of acetyl radical just from lysine molecules of histones, but their actions is not limited to histones, they can also act on non-histone proteins [175].

HDAC inhibitors are classified into four classes, based on their chemical structure: shortchain fatty acids, hydroxamic acids, cyclic peptides, benzamides (hybrid molecules) [176].

drug has been associated with: carboplatin useful in solid tumors treatment [167], valproic acid in acute myeloid leukemias and advanced leukemia [168, 169], imatinib mesylate in chronic myelogenous leukemia (CML) [170] and IL-2 in metastatic melanoma, renal carcinoma [171]. Zebularine (2-pyrimidone-1-β- D-riboside) is other nucleoside analog with hypomethylation

**Figure 3.** Structure of Zebularine. Zebularine is another nucleoside analog drug, with hypomethilation effect.

methylation inhibition [174].

tumors.

arhythmic), RG108 and SGI-1027.

**5.2. Targeting histone modification**

non-histone proteins [175].

There are recent studies about another two molecules: NPEOC-DAC and SGI 110 (S110). NPEOC-DAC is the result of chemical reaction between azacytosine molecule and 2-(p-ni‐ trophenyl) ethoxycarbonyl, with reported effect on DNA methyltransferases inhibition. By‐ un et al. demonstrated that NPEOC-DAC inhibited DNA methylation in two cell lines of liver cancer. The authors, also showed that SGI 110 (S110) has a pronounced effect on DNA

The non-nucleoside analogs category contains compounds with hypomethylation effect. This group contains hydralazine (the widely known as vasodilatator), procainamide (anti-

Physiologically, acetylation of chromatin is realized by specific enzymes - histone deace‐ tylases and acetyltransferases. A possible change in their normal function can promote

At first glance, HDACs are enzymes that play a role in elimination of acetyl radical just from lysine molecules of histones, but their actions is not limited to histones, they can also act on

HDAC inhibitors are classified into four classes, based on their chemical structure: shortchain fatty acids, hydroxamic acids, cyclic peptides, benzamides (hybrid molecules) [176].

activity [172] and also implicated in tumor gene expression [173].

166 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 4.** HDAC inhibitors. There are four classes of curently known HDAC inhibitors: short-chain fatty acids, hydroxa‐ mic acids, cyclic peptides, benzamides, with a great potential use as detection and prognosis markers.

Exemples of short-chain fatty acids are: sodium n-butyrate, sodium phenylacetate, phenyl‐ butyrate, valproate, substances that in millimolar concentrations are involved in inhibition the growth of some carcinomas but their mechanism of action is not fully understood [163, 177-179].

One of the most studied agent from class of small fatty acids, is valproic acid (VPA), an anti‐ epileptic drug reported to target histone deacetylase. Numerous research studies *in vitro* demonstrated that VPA was implicated in hyperacetylation of histones H3 and H4 and also *in vivo* tests confirmed the drug inhibiting action of HDACs. VPA antitumor activity was de‐ manstrated by: cell growth inhibition, apoptosis inducing, antimetastatic and antiangiogene‐ sis effect, etc. These benefits lead to FDA approving of VPA [14, 180].

**6. Conclusions and perspectives**

cancer cells have been mentioned in several studies.

Grant support: 134/2011 UEFISCDI Romania

, Stefana Oana Purcaru1

ly target cancer stem cells.

**Acknowledgments**

**Author details**

Oana Daianu1,3, Bogdan Stoleru1

Ligia Gabriela Tataranu3

Anica Dricu1

Epigenetic is a heritage mechanism involved in the process of stem cells differentiation to more specialized cells. According to the cancer stem cell model, dysregulation of epigenetic mechanisms (i.e. DNA methylation and histone modification) in pluripotent stem cells ena‐ ble their transformation in cancer cells with high proliferation rates and poor prognosis.

DNA methylation is considered the most largely studied part of the epigenetic, but recent works associate the methylation with other epigenetic changes, such as histone modifica‐ tions, chromatin remodeling and microRNA, suggesting a reciprocal relationship between them in cancer cells. The similarities between chromatin regulation process in stem cells and

It is therefore important to understand the epigenetic alterations that take place in cancer cells compared with normal cells and the importance of these modifications in carcinogene‐ sis, according to the cancer stem cell theory. In addition, it is very useful to understand the potential of epigenetic marks in designing more effective treatment strategies that specifical‐

, Alice Sandra Buteica2

, Amelia Mihaela Dobrescu4

\*Address all correspondence to: anica.dricu@live.co.uk anicadricu@webmail.umfcv.ro

1 Department of Biochemistry, University of Medicine and Pharmacy of Craiova, Romania

2 Department of Pharmacology, University of Medicine and Pharmacy of Craiova, Romania

3 Department of Neurosurgery, "Bagdasar-Arseni" Emergency Hospital, Bucharest, Romania

4 Department of Medical Genetics,University of Medicine and Pharmacy of Craiova, Romania

5 Department of Microbiology, University of Medicine and Pharmacy of Craiova, Romania

, Daniela Elise Tache1

, Tiberiu Daianu5

,

and

DNA Methylation, Stem Cells and Cancer http://dx.doi.org/10.5772/53263 169

**Figure 5.** Structure of valproic acid. Valproic acid is a small fatty acid commonly used as an antiepileptic drug, but with recently emerged antitumor effects.

The class of hydroxamic acids include synthesized compounds such as: belinostat, panobino‐ stat, vorinostat (SAHA) etc. Belinostat and panobinostat, have been used in clinical trials to treat solid tumors and blood malignancies [181-183]; MDL and CML [184-186], vorinostat (SAHA) that was approved by FDA for the treatment of chronic T-cell lymphoma (CTCL) and used in clinical trials for hematologic malignancies, mesothelioma, breast and ovarian cancer, etc [175].

A natural compound from cyclic peptides class is romidepsin, also known as Istodax (FK228), which was clinical tested in various lymphomas. The drug was shown to induce apoptosis in different tumor cell lines, due to blocking of HDACs [187].

Hybrid molecules (i.e. benzamides) includes two synthetic compounds: Entinostat (MS-275) and Mocetinostat (MGCD 0103). The mechanism by which Entinostat induced cytotoxic ef‐ fect on tumor cells was suggested to be due to the upregulation of some tumor suppressor genes (p21]. Both Entinostat and Mocetinostat are currently approved by the FDA and are used in cancer treatment. Entinostat is used in the treatment of blood and lung tumor [181, 183] and Mocetinostatin in the treatment of chronic lymphocytic leukemia (CLL) [175].

HATs are a class of enzymes discovered twenty years ago, enzymes with demonstrated role in gene transcription [188]. HATs have been reported to be implicate in numerous types of diseases (i.e. viral infection, respiratory maladies, cancer etc). It has been suggested that the HATs enzymes may be used as biological markers for cancer prediction or recurrence [14]. Four families of HATs are known that share primary-structure homology: GNAT (Gcn5-re‐ lated N-acetyltransferase), p300/CBP and MYST, Rtt109 [189]. The HAT enzymes have vari‐ ous chemical structure and their classification is still unclear.

Histone methylation process plays an important task in epigenetic regulation, which lead to synthesizing of new target drugs for cancer therapy [163].

Researchers describe a class of enzymes called histone methyltransferases. This class of en‐ zymes includes lysine methyltransferases and arginine methyltransferases, both of them linked to many types of cancer.

There are 8 known lysine methyltransferases (KMT1-8) with suggested role in the epigenetic gene silencing in malignancies like: prostate, liver, colon, breast cancer [190, 191].

Few of the many types of arginine methyltransferases (PRMTs), are also closely linked to cancer [191].

Thus, the importance of DNMTs and HDACs, two classes of enzymes involved in epigenetic targeted therapy of malignant diseases, is obvious. The enzymes implicated in histone meth‐ ylation and demethylation are mainly attractive as validated targets for cancer therapy.

#### **6. Conclusions and perspectives**

H3C CH3

valproic acid

**Figure 5.** Structure of valproic acid. Valproic acid is a small fatty acid commonly used as an antiepileptic drug, but with

The class of hydroxamic acids include synthesized compounds such as: belinostat, panobino‐ stat, vorinostat (SAHA) etc. Belinostat and panobinostat, have been used in clinical trials to treat solid tumors and blood malignancies [181-183]; MDL and CML [184-186], vorinostat (SAHA) that was approved by FDA for the treatment of chronic T-cell lymphoma (CTCL) and used in clinical trials for hematologic malignancies, mesothelioma, breast and ovarian cancer, etc [175]. A natural compound from cyclic peptides class is romidepsin, also known as Istodax (FK228), which was clinical tested in various lymphomas. The drug was shown to induce

Hybrid molecules (i.e. benzamides) includes two synthetic compounds: Entinostat (MS-275) and Mocetinostat (MGCD 0103). The mechanism by which Entinostat induced cytotoxic ef‐ fect on tumor cells was suggested to be due to the upregulation of some tumor suppressor genes (p21]. Both Entinostat and Mocetinostat are currently approved by the FDA and are used in cancer treatment. Entinostat is used in the treatment of blood and lung tumor [181, 183] and Mocetinostatin in the treatment of chronic lymphocytic leukemia (CLL) [175].

HATs are a class of enzymes discovered twenty years ago, enzymes with demonstrated role in gene transcription [188]. HATs have been reported to be implicate in numerous types of diseases (i.e. viral infection, respiratory maladies, cancer etc). It has been suggested that the HATs enzymes may be used as biological markers for cancer prediction or recurrence [14]. Four families of HATs are known that share primary-structure homology: GNAT (Gcn5-re‐ lated N-acetyltransferase), p300/CBP and MYST, Rtt109 [189]. The HAT enzymes have vari‐

Histone methylation process plays an important task in epigenetic regulation, which lead to

Researchers describe a class of enzymes called histone methyltransferases. This class of en‐ zymes includes lysine methyltransferases and arginine methyltransferases, both of them

There are 8 known lysine methyltransferases (KMT1-8) with suggested role in the epigenetic

Few of the many types of arginine methyltransferases (PRMTs), are also closely linked to

Thus, the importance of DNMTs and HDACs, two classes of enzymes involved in epigenetic targeted therapy of malignant diseases, is obvious. The enzymes implicated in histone meth‐ ylation and demethylation are mainly attractive as validated targets for cancer therapy.

gene silencing in malignancies like: prostate, liver, colon, breast cancer [190, 191].

apoptosis in different tumor cell lines, due to blocking of HDACs [187].

ous chemical structure and their classification is still unclear.

synthesizing of new target drugs for cancer therapy [163].

linked to many types of cancer.

cancer [191].

recently emerged antitumor effects.

168 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

O OH

Epigenetic is a heritage mechanism involved in the process of stem cells differentiation to more specialized cells. According to the cancer stem cell model, dysregulation of epigenetic mechanisms (i.e. DNA methylation and histone modification) in pluripotent stem cells ena‐ ble their transformation in cancer cells with high proliferation rates and poor prognosis.

DNA methylation is considered the most largely studied part of the epigenetic, but recent works associate the methylation with other epigenetic changes, such as histone modifica‐ tions, chromatin remodeling and microRNA, suggesting a reciprocal relationship between them in cancer cells. The similarities between chromatin regulation process in stem cells and cancer cells have been mentioned in several studies.

It is therefore important to understand the epigenetic alterations that take place in cancer cells compared with normal cells and the importance of these modifications in carcinogene‐ sis, according to the cancer stem cell theory. In addition, it is very useful to understand the potential of epigenetic marks in designing more effective treatment strategies that specifical‐ ly target cancer stem cells.

#### **Acknowledgments**

Grant support: 134/2011 UEFISCDI Romania

#### **Author details**

Anica Dricu1 , Stefana Oana Purcaru1 , Alice Sandra Buteica2 , Daniela Elise Tache1 , Oana Daianu1,3, Bogdan Stoleru1 , Amelia Mihaela Dobrescu4 , Tiberiu Daianu5 and Ligia Gabriela Tataranu3

\*Address all correspondence to: anica.dricu@live.co.uk anicadricu@webmail.umfcv.ro

1 Department of Biochemistry, University of Medicine and Pharmacy of Craiova, Romania

2 Department of Pharmacology, University of Medicine and Pharmacy of Craiova, Romania

3 Department of Neurosurgery, "Bagdasar-Arseni" Emergency Hospital, Bucharest, Romania

4 Department of Medical Genetics,University of Medicine and Pharmacy of Craiova, Romania

5 Department of Microbiology, University of Medicine and Pharmacy of Craiova, Romania

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[185] Chen S, Ye J, Kijima I, Evans D. The HDAC inhibitor LBH589 (panobinostat) is an in‐ hibitory modulator of aromatase gene expression. Proc Natl Acad Sci U S A. 2010 Jun 15;107(24):11032-7.

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**Chapter 8**

**DNA Methylation in the Pathogenesis of Head and**

Head and neck cancer is the sixth most common cancer worldwide and one of the most ag‐ gressive malignancies in human population. The most common histologic type among the head and neck tumors are the squamous cell carcinomas (*SCC*). Despite the significant ef‐ forts committed during the last decades in its early detection, prevention and treatment, head and neck cancer prognosis remains very poor with the rising incidence in developed countries and younger population. Carcinogenesis of Head and Neck Squamous Cell Carci‐ noma (*HNSCC*) is a multistep process, which arises through an accumulation of genetic and epigenetic alterations. Although the impact of genetic changes in oral carcinogenesis is wellknown, over the last decade it has been demonstrated that epigenetic changes, especially

Head and Neck Squamous Cell Carcinoma is the sixth most common cancer in males and tenth in females worldwide [1]. Despite the fact that significant results have been achieved during the last decades in its early detection, prevention and treatment, the survival rate has remained less than 40%, and *HNSCC* remains one of the most aggressive malignancies. Fur‐ thermore, the incidence of this carcinoma is rising in developed countries and younger pop‐ ulation, particularly young women [2, 3]. Early stages of the disease are associated with minimal symptoms, thus small percentage of *HNSCC* has been diagnosed at an early clinical stage. Advanced stages respond poorly to current cancer therapies, with high incidence of

> © 2013 Magić et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

**Neck Cancer**

Zvonko Magić, Gordana Supić,

http://dx.doi.org/10.5772/51169

**1. Introduction**

Mirjana Branković-Magić and Nebojša Jović

Additional information is available at the end of the chapter

aberrant DNA methylation, play a significant role in *HNSCC*.

local and regional relapse and lymph node metastasis [2, 4, 5].

**1.1. Head and neck cancer – Etiology and risk factors**

### **DNA Methylation in the Pathogenesis of Head and Neck Cancer**

Zvonko Magić, Gordana Supić, Mirjana Branković-Magić and Nebojša Jović

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51169

#### **1. Introduction**

Head and neck cancer is the sixth most common cancer worldwide and one of the most ag‐ gressive malignancies in human population. The most common histologic type among the head and neck tumors are the squamous cell carcinomas (*SCC*). Despite the significant ef‐ forts committed during the last decades in its early detection, prevention and treatment, head and neck cancer prognosis remains very poor with the rising incidence in developed countries and younger population. Carcinogenesis of Head and Neck Squamous Cell Carci‐ noma (*HNSCC*) is a multistep process, which arises through an accumulation of genetic and epigenetic alterations. Although the impact of genetic changes in oral carcinogenesis is wellknown, over the last decade it has been demonstrated that epigenetic changes, especially aberrant DNA methylation, play a significant role in *HNSCC*.

#### **1.1. Head and neck cancer – Etiology and risk factors**

Head and Neck Squamous Cell Carcinoma is the sixth most common cancer in males and tenth in females worldwide [1]. Despite the fact that significant results have been achieved during the last decades in its early detection, prevention and treatment, the survival rate has remained less than 40%, and *HNSCC* remains one of the most aggressive malignancies. Fur‐ thermore, the incidence of this carcinoma is rising in developed countries and younger pop‐ ulation, particularly young women [2, 3]. Early stages of the disease are associated with minimal symptoms, thus small percentage of *HNSCC* has been diagnosed at an early clinical stage. Advanced stages respond poorly to current cancer therapies, with high incidence of local and regional relapse and lymph node metastasis [2, 4, 5].

© 2013 Magić et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Head and neck cancers include malignancies arising from different anatomical sites within the upper aero-digestive tract. Head and neck cancers are characterized by heterogeneous histology. The majority carcinomas that arise from squamous cell epithelia are head and neck squamous cell carcinomas (*HNSCC*), while other cancer types that can occur in the head and neck include thyroid cancer, malignant salivary gland tumors, lymphomas and sarcomas. *HNSCC* include cancers of the oral cavity, larynx, pharynx (oropharynx, hypo‐ pharynx, nasopharynx), and esophagus, [4, 5], Figure 1.

effect directly, either acting as a solvent of carcinogens from tobacco smoke or damaging the oral mucosa, that enhances the penetration of carcinogens from tobacco smoke [7]. Genetic polymorphisms of ethanol-metabolizing enzymes, including alcohol dehydrogenases (ADH) (which metabolizes alcohol into acetaldehyde), with a different ability to generate carcinogen acetaldehyde, may determine individual susceptibility to head and neck cancer. Acetaldehyde can form adducts with DNA, interfering with DNA synthesis and repair [7, 13]. "Fast-metabolizing" ADHs genotype was associated with the increase of *OSCC* and *HNSCC* risk [14, 15, 16]. By the contrast, in other studies "fast-metabolizing" ADHs geno‐ type was found to be associated with decreased risk of *HNSCC* [11, 17, 18]. Therefore, the mechanism by which smoking and alcohol causes increased risk for *HNSCC* and the role of

DNA Methylation in the Pathogenesis of Head and Neck Cancer

http://dx.doi.org/10.5772/51169

187

**Figure 1.** Diverse anatomical localization of Head and Neck Squamous Cell Carcinomas. Although head and neck can‐ cers are characterized by heterogeneous histology, the majority carcinomas arise from squamous cell epithelia of the oral cavity (tongue, buccal mucosa, floor of the mouth), oropharynx (soft palate, base of tongue, tonsils), hypophar‐

In addition to alcohol and tobacco exposure, human papilloma virus (HPV) infection has al‐ so a significant part in the etiology of head and neck cancer. HPV cancers predominantly arise from tongue and palatine tonsils within the oropharynx [5, 19]. Acting in synergy with tobacco use and heavy alcohol consumption, HPV infection with high-risk types is consid‐ ered as an etiological factor in the development of *HSCC* and *OSCC* [5]. Reported incidence

ynx, nasopharynx, larynx (supraglotis, glottis, subglotis) and esophagus.

**1.4. HPV infection in HNSCC**

alcohol and tobacco-related polymorphisms have not been fully elucidated.

#### **1.2. Risk factors for HNSCC**

Both environmental and genetic factors play an important role in the etiology of head and neck cancers but the causal relationship between environmental factors, lifestyle and tumor development is not yet fully elucidated. The mucosa of the upper aerodigestive tract is ex‐ posed to number of carcinogens attributed to cause genetic and epigenetic changes that ulti‐ mately lead to head and neck cancer development. *HNSCC* incidence is influenced by age, genetic factors, geographic region and different lifestyle factors, including alcohol, smoking, betel quid use, oral hygiene, and Human Papilloma Virus (HPV) infection [5]. Emerging evi‐ dences are indicating that environmental factors, such as smoking, alcohol and diet could directly or indirectly affect epigenetic mechanisms of gene expression regulation and DNA methylation in *HNSCC*.

#### **1.3. Tobacco and alcohol use in HNSCC**

Smoking and alcohol consumption are the major risk factors for head and neck cancers [6, 7]. In addition, the combination of both alcohol and tobacco use synergically increases the risk for *HNSCC*development more than 10 times [8]. Cigarette smoking is the major cause of lung cancer and is associated with head and neck, esophagus, bladder, breast and kidney cancer [9]. Increased risk with smoking may be due to the direct effect of tobacco carcino‐ gens or due to genetic polymorphisms in enzymes that activate or detoxify carcinogens.

Several carcinogens present in cigarette smoke are inactivated by a family of enzymes cyto‐ chrome P-450 (CYP), which convert carcinogens into reactivated intermediates. These inter‐ mediates form DNA adducts that need to be detoxified by a number of enzymes, including glutathione S-transferase [GST) [10]. Single nucleotide polymorphisms (SNPs) in these genes could be an alternative mechanism that modulates the effects of cigarette smoke. Even though alcohol and smoking are known risk factors, only a fraction of smokers and alcohol consumers develop *HNSCC,* suggesting that genetic susceptibility and interactions between genetic, epigenetic and environmental factors could play an important role in the etiology of *HNSCC* [11, 12].

Alcohol consumption has been associated with an increased risk of the head and neck, esophagus, liver, colorectal, and breast cancer [13]. Possible mechanisms by which alcohol exerts its harmful effect includes the genotoxic effect of ethanol metabolite acetaldehyde, production of reactive oxygen- and nitrogen species, changes in folate metabolism, genera‐ tion of DNA adducts and inhibition of DNA repair. Also, alcohol could exert its damaging effect directly, either acting as a solvent of carcinogens from tobacco smoke or damaging the oral mucosa, that enhances the penetration of carcinogens from tobacco smoke [7]. Genetic polymorphisms of ethanol-metabolizing enzymes, including alcohol dehydrogenases (ADH) (which metabolizes alcohol into acetaldehyde), with a different ability to generate carcinogen acetaldehyde, may determine individual susceptibility to head and neck cancer. Acetaldehyde can form adducts with DNA, interfering with DNA synthesis and repair [7, 13]. "Fast-metabolizing" ADHs genotype was associated with the increase of *OSCC* and *HNSCC* risk [14, 15, 16]. By the contrast, in other studies "fast-metabolizing" ADHs geno‐ type was found to be associated with decreased risk of *HNSCC* [11, 17, 18]. Therefore, the mechanism by which smoking and alcohol causes increased risk for *HNSCC* and the role of alcohol and tobacco-related polymorphisms have not been fully elucidated.

**Figure 1.** Diverse anatomical localization of Head and Neck Squamous Cell Carcinomas. Although head and neck can‐ cers are characterized by heterogeneous histology, the majority carcinomas arise from squamous cell epithelia of the oral cavity (tongue, buccal mucosa, floor of the mouth), oropharynx (soft palate, base of tongue, tonsils), hypophar‐ ynx, nasopharynx, larynx (supraglotis, glottis, subglotis) and esophagus.

#### **1.4. HPV infection in HNSCC**

Head and neck cancers include malignancies arising from different anatomical sites within the upper aero-digestive tract. Head and neck cancers are characterized by heterogeneous histology. The majority carcinomas that arise from squamous cell epithelia are head and neck squamous cell carcinomas (*HNSCC*), while other cancer types that can occur in the head and neck include thyroid cancer, malignant salivary gland tumors, lymphomas and sarcomas. *HNSCC* include cancers of the oral cavity, larynx, pharynx (oropharynx, hypo‐

Both environmental and genetic factors play an important role in the etiology of head and neck cancers but the causal relationship between environmental factors, lifestyle and tumor development is not yet fully elucidated. The mucosa of the upper aerodigestive tract is ex‐ posed to number of carcinogens attributed to cause genetic and epigenetic changes that ulti‐ mately lead to head and neck cancer development. *HNSCC* incidence is influenced by age, genetic factors, geographic region and different lifestyle factors, including alcohol, smoking, betel quid use, oral hygiene, and Human Papilloma Virus (HPV) infection [5]. Emerging evi‐ dences are indicating that environmental factors, such as smoking, alcohol and diet could directly or indirectly affect epigenetic mechanisms of gene expression regulation and DNA

Smoking and alcohol consumption are the major risk factors for head and neck cancers [6, 7]. In addition, the combination of both alcohol and tobacco use synergically increases the risk for *HNSCC*development more than 10 times [8]. Cigarette smoking is the major cause of lung cancer and is associated with head and neck, esophagus, bladder, breast and kidney cancer [9]. Increased risk with smoking may be due to the direct effect of tobacco carcino‐ gens or due to genetic polymorphisms in enzymes that activate or detoxify carcinogens.

Several carcinogens present in cigarette smoke are inactivated by a family of enzymes cyto‐ chrome P-450 (CYP), which convert carcinogens into reactivated intermediates. These inter‐ mediates form DNA adducts that need to be detoxified by a number of enzymes, including glutathione S-transferase [GST) [10]. Single nucleotide polymorphisms (SNPs) in these genes could be an alternative mechanism that modulates the effects of cigarette smoke. Even though alcohol and smoking are known risk factors, only a fraction of smokers and alcohol consumers develop *HNSCC,* suggesting that genetic susceptibility and interactions between genetic, epigenetic and environmental factors could play an important role in the etiology of

Alcohol consumption has been associated with an increased risk of the head and neck, esophagus, liver, colorectal, and breast cancer [13]. Possible mechanisms by which alcohol exerts its harmful effect includes the genotoxic effect of ethanol metabolite acetaldehyde, production of reactive oxygen- and nitrogen species, changes in folate metabolism, genera‐ tion of DNA adducts and inhibition of DNA repair. Also, alcohol could exert its damaging

pharynx, nasopharynx), and esophagus, [4, 5], Figure 1.

186 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**1.2. Risk factors for HNSCC**

methylation in *HNSCC*.

*HNSCC* [11, 12].

**1.3. Tobacco and alcohol use in HNSCC**

In addition to alcohol and tobacco exposure, human papilloma virus (HPV) infection has al‐ so a significant part in the etiology of head and neck cancer. HPV cancers predominantly arise from tongue and palatine tonsils within the oropharynx [5, 19]. Acting in synergy with tobacco use and heavy alcohol consumption, HPV infection with high-risk types is consid‐ ered as an etiological factor in the development of *HSCC* and *OSCC* [5]. Reported incidence of high-risk HPVs in oral carcinoma patients varied from 0% to 100% [20], depending on the methods used for HPV detection, tumor-host characteristics of the examined group of pa‐ tients, varying numbers of included tissue samples, but it is also affected with different dis‐ tribution of oncogenic HPVs in different world regions. The DNA of oncogenic HPVs is present in 20% of all *HNSSCs* and in nearly 60% of tonsillar cancers [5, 8]. There is increasing evidence that HPV-associated *HNSCC* carcinomas are distinct clinical and pathological tu‐ mor entity from alcohol and smoking-associated *HNSCC*s with regards to risk factors, tumor biology and progression [5]. Infection with oncogenic HPV types, predominantly HPV16 and HPV18 is associated with increased risk of *HNSSC* [5, 19].

recent studies indicate that dietary habits and fruit and vegetable intake could be associated with cancer risk [26, 27]. Increased fruit and vegetable consumption was recently associated with reduced occurrence of *HNSCC* [28]. However, the findings of association of diet and

DNA Methylation in the Pathogenesis of Head and Neck Cancer

http://dx.doi.org/10.5772/51169

189

**Figure 2.** Schematic presentation of synergistic effect of different genetic alterations in transformation of normal to malignant cell. Gene mutations can be inherited (BRCA1, BRCA2, RET, VHL, APC) and associated with a predisposition to certain cancer or cancer syndromes. Also, mutations can be acquired leading to the strong predisposition to malig‐ nant transformation of cells (K-*ras*, p53, HER2, *bcr-abl, bcl-2*, Rb). The human genome is characterized with enormous number of genetic polymorphisms that sometimes results with impaired function of the gene product, especially when cells are exposed to environmental or host risk factors. Hypermethylation of gene promoter sequences causes gene silencing that is of great importance for tumor suppressor genes. All these genetic alterations can impair func‐

Second half of 20th century and beginning of 21st century are marked with almost revolution‐ ary breakthrough in our understanding of human gene structure and functions. The rapid development of biotechniques led to the sequencing of complete human genome and later of genome and transcriptome of many human tumors [29, 30]. Even before these achievements, it was well accepted that malignant tumors, on the molecular level, are disease of genes. This knowledge led to enormous number of worldwide research projects and publications concerning association of gene alterations in tumor cells with cancerogenesis and early diag‐ nosis, tumor progression, tumor staging and response to therapy. In spite of many practical/

tion of other genes and further contribute to increased risk of malignant transformation.

**2. Molecular changes In** *HNSCC*

cancer risk are still confounding.

HPV infection causes deregulation of cell cycle and apoptosis by inactivation of *Rb* and *p53* tu‐ mor suppressor gene protein products, involved in the maintenance of genome stability. E6 protein of high risk HPVs affects the *p53* protein function by ubiquitin-dependent degrada‐ tion. Although *p53* tumor suppressor gene is the most often mutated gene in human oncology, in tumors with HPV etiology is not clear in which extent the mechanism of *p53* mutation is in‐ cluded in *p53* inactivation. It can be assumed that *p53* mutations occur most frequently in HPV negative than in the HPV positive head and neck cancers, but it is questionable if the presence of *p53* mutation in HPV infected tumors additionally influences prognosis of the patients. Nu‐ merous studies with conflicting results deal with the influence of oncogenic HPV types and prognosis of *HSCC*. The majority of them reported that *HSCC* patients with HPV infection have better prognosis than the patients who are HPV negative [5, 21, 22]. On the contrary, our previous results concerning *OSCC* patients in stage III of the disease showed worse overall and disease-free survival for patients infected with high-risk HPV types (HPV 16, 18 and 31) [23], indicating the presence of more aggressive disease in these patients. Our study comparing dis‐ ease-free interval (DFI) and overall survival (OS) between patients with HPV infection only and the patients with HPV infection and *p53* mutation showed significantly shorter diseasefree interval as well as overall survival in patients with both HPV infection and *p53* mutation. Presence of *p53* mutations in HPV infected tumors confer a higher risk of recurrence in this dis‐ ease. In addition, since all patients were treated with postoperative radiotherapy, shorter DFI indicate that the response to radiotherapy may be influenced by *p53* status [24]. However, it has been recently reported that it is unlikely that HPV infection plays significant role in mobile tongue carcinogenesis in young *OSCC* patients [25].

The role of HPV infection in the etiology, as well as in prognosis of head and neck cancers varies and it depends on head and neck cancer subtypes and different anatomic site of tu‐ mors. From that point of view, it is clear that HPV typing, together with other molecular markers may help in defining a particular group of tumors in regard to prognosis and re‐ sponse to anti cancer therapies.

#### **1.5. Diet and HNSCC**

Recent studies provide growing evidence that some dietary components, might affect the process of carcinogenesis. There is a growing body of evidence that bioactive food compo‐ nents, such as isothiocyanates from cruciferous vegetables (cauliflower, cabbage, and broc‐ coli), diallyl sulfide (an organosulphur compound from garlic), green tea flavonoids, folate, and selenium, effect on cancer might be mediated through epigenetic mechanisms. Several recent studies indicate that dietary habits and fruit and vegetable intake could be associated with cancer risk [26, 27]. Increased fruit and vegetable consumption was recently associated with reduced occurrence of *HNSCC* [28]. However, the findings of association of diet and cancer risk are still confounding.

**Figure 2.** Schematic presentation of synergistic effect of different genetic alterations in transformation of normal to malignant cell. Gene mutations can be inherited (BRCA1, BRCA2, RET, VHL, APC) and associated with a predisposition to certain cancer or cancer syndromes. Also, mutations can be acquired leading to the strong predisposition to malig‐ nant transformation of cells (K-*ras*, p53, HER2, *bcr-abl, bcl-2*, Rb). The human genome is characterized with enormous number of genetic polymorphisms that sometimes results with impaired function of the gene product, especially when cells are exposed to environmental or host risk factors. Hypermethylation of gene promoter sequences causes gene silencing that is of great importance for tumor suppressor genes. All these genetic alterations can impair func‐ tion of other genes and further contribute to increased risk of malignant transformation.

#### **2. Molecular changes In** *HNSCC*

of high-risk HPVs in oral carcinoma patients varied from 0% to 100% [20], depending on the methods used for HPV detection, tumor-host characteristics of the examined group of pa‐ tients, varying numbers of included tissue samples, but it is also affected with different dis‐ tribution of oncogenic HPVs in different world regions. The DNA of oncogenic HPVs is present in 20% of all *HNSSCs* and in nearly 60% of tonsillar cancers [5, 8]. There is increasing evidence that HPV-associated *HNSCC* carcinomas are distinct clinical and pathological tu‐ mor entity from alcohol and smoking-associated *HNSCC*s with regards to risk factors, tumor biology and progression [5]. Infection with oncogenic HPV types, predominantly HPV16

HPV infection causes deregulation of cell cycle and apoptosis by inactivation of *Rb* and *p53* tu‐ mor suppressor gene protein products, involved in the maintenance of genome stability. E6 protein of high risk HPVs affects the *p53* protein function by ubiquitin-dependent degrada‐ tion. Although *p53* tumor suppressor gene is the most often mutated gene in human oncology, in tumors with HPV etiology is not clear in which extent the mechanism of *p53* mutation is in‐ cluded in *p53* inactivation. It can be assumed that *p53* mutations occur most frequently in HPV negative than in the HPV positive head and neck cancers, but it is questionable if the presence of *p53* mutation in HPV infected tumors additionally influences prognosis of the patients. Nu‐ merous studies with conflicting results deal with the influence of oncogenic HPV types and prognosis of *HSCC*. The majority of them reported that *HSCC* patients with HPV infection have better prognosis than the patients who are HPV negative [5, 21, 22]. On the contrary, our previous results concerning *OSCC* patients in stage III of the disease showed worse overall and disease-free survival for patients infected with high-risk HPV types (HPV 16, 18 and 31) [23], indicating the presence of more aggressive disease in these patients. Our study comparing dis‐ ease-free interval (DFI) and overall survival (OS) between patients with HPV infection only and the patients with HPV infection and *p53* mutation showed significantly shorter diseasefree interval as well as overall survival in patients with both HPV infection and *p53* mutation. Presence of *p53* mutations in HPV infected tumors confer a higher risk of recurrence in this dis‐ ease. In addition, since all patients were treated with postoperative radiotherapy, shorter DFI indicate that the response to radiotherapy may be influenced by *p53* status [24]. However, it has been recently reported that it is unlikely that HPV infection plays significant role in mobile

The role of HPV infection in the etiology, as well as in prognosis of head and neck cancers varies and it depends on head and neck cancer subtypes and different anatomic site of tu‐ mors. From that point of view, it is clear that HPV typing, together with other molecular markers may help in defining a particular group of tumors in regard to prognosis and re‐

Recent studies provide growing evidence that some dietary components, might affect the process of carcinogenesis. There is a growing body of evidence that bioactive food compo‐ nents, such as isothiocyanates from cruciferous vegetables (cauliflower, cabbage, and broc‐ coli), diallyl sulfide (an organosulphur compound from garlic), green tea flavonoids, folate, and selenium, effect on cancer might be mediated through epigenetic mechanisms. Several

and HPV18 is associated with increased risk of *HNSSC* [5, 19].

188 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

tongue carcinogenesis in young *OSCC* patients [25].

sponse to anti cancer therapies.

**1.5. Diet and HNSCC**

Second half of 20th century and beginning of 21st century are marked with almost revolution‐ ary breakthrough in our understanding of human gene structure and functions. The rapid development of biotechniques led to the sequencing of complete human genome and later of genome and transcriptome of many human tumors [29, 30]. Even before these achievements, it was well accepted that malignant tumors, on the molecular level, are disease of genes. This knowledge led to enormous number of worldwide research projects and publications concerning association of gene alterations in tumor cells with cancerogenesis and early diag‐ nosis, tumor progression, tumor staging and response to therapy. In spite of many practical/ clinical results of these studies [Her-2/neu, *BRCA-1* and *BRCA-2* in Breast Cancer; *K-ras* and *EGFR* in Colon Cancer; *bcr-abl* in Chronic Myeloid Leukemia; *B-RAF* and *c-KIT* in Melano‐ ma; detection of dominant clone of B- and T-lymphocytes in tracking minimal residual dis‐ ease in lymphomas and leukemia's [29-33], still much more questions concerning significance of gene alterations for tumor development and treatment remains unanswered. In the last 10-15 years significant efforts are done to fulfill these gaps in our understanding of molecular pathogenesis of cancer by studying epigenetic changes of tumor tissues.

Genetic instability in regions leads to loss of chromosomal region that contains tumor suppres‐ sor genes. A high incidence of Loss of heterozygosity (LOH) observed at 9p, 8p, 3p, 9q, and 11q regions were associated with tumor stage, and poor histological differentiation in *HNSCC* [37]. Loss of heterozygosity (LOH) of 9p21 appears is an early and frequent event in head and neck carcinogenesis [37, 38-40]. The CDKN2A gene locus found in 9p21 region encodes two differ‐ ent transcripts, *p16* and *p14*ARF, which are responsible for G1 cell cycle regulation and MDM2 mediated degradation of *p53*. *P16* and *p14* are often inactivated in *HNSCC* through homozy‐

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Loss of chromosome region 3p is a common early genetic event in *HNSCC* [2, 40, 45]. Genet‐ ic alterations at 3p that are common in *HNSCC* and *OSCC* contains tumor suppressor genes Fragile histidine triad (*FHIT*) gene mapped at 3*p14*, Ras association family (*RASSF1A*) gene, mapped at 3p21, and Retinoic acid receptor B2 gene (*RARB2*), mapped to chromosome re‐ gion 3p24 [37, 40]. Amplification of 11q13 leads to overexpression of cyclin D1 [46], detected

There has been much research on the tumor suppressor gene *p53* and its role in *HNSCC* and *OSCC* carcinogenesis. The *p53* protein blocks cell division at the G1 to S boundary, stimu‐ lates DNA repair after DNA damage, and induces apoptosis. *P53* has been shown to be functionally inactivated in oral and head and neck tumors [48]. LOH of 17p13, which con‐ tains *p53* and point mutations of the *p53* are seen in more than 50% of *HNSCC* cases [39, 49]. Cigarette smoking has been associated with the mutation of *p53* in head and neck cancers [14]. Investigations about the prognostic value of *p53* status in head and neck cancer sub‐ types hardly can get a clear result due to the presence of large heterogeneity in regard to patients characteristics, and methods for *p53* detection. Meta-analysis of Tandon et al [50] pointed out that evidence about the prognostic value of *p53* in head and neck cancer squa‐ mous cell carcinomas has been inconclusive. However, determination of *p53* status of *HNSCC* may be of particular interests due to its possible role in prediction of the response to cisplatin based chemotherapy. Cisplatin chemotherapy is widely used chemotherapeutical approach in the treatment of head and neck cancers and data that patients with *p53* muta‐ tions respond better to cisplatin chemotherapy need further confirmation [51]. Chemothera‐ py responses in *HNSCC* patients is also influenced by polymorphism at codon 72 of *p53* gene. In patients with wild type *p53*, arginine at codon 72 is associated with better response to chemotherapy. On the contrary, if *p53* is mutated, proline at position 72 is better option [52]. Better understanding of cellular processes and their actors such as *p53* network and their relation to clinical settings will help in appropriate biomarker characterization. Epider‐ mal growth factor receptor (*EGFR*) is overexpressed in 90% of *HNSCC* [53], often due to am‐ plification [54]. Thus, frequent molecular abnormalities in head and neck squamous cell carcinoma include alterations in tumor suppressor genes *p16*INK4A, *p53*, *p14*ARF, *FHIT*, *RASSF1A*, *Rb*, cyclin D1, and activation of oncogenes, such as members of the *ras* gene fami‐

A large number of previous studies have also suggested the correlation between polymor‐ phisms of genes involved in cell cycle control, angiogenesis and metabolism of alcohol and

carcinogens and susceptibility to head and neck cancer [10, 16, 34-36].

gous deletion, by promoter methylation, and by point mutations [41-44].

in 30–60% of *HNSCC* and 40% of cases of oral squamous dysplasia [47].

ly, *c-myc*, and *EGFR* [2-4], Table 1.

These alterations are of special interest because epigenetic changes are tumor/tissue specific, although inheritable epigenetic alterations are under strong influence of environmental risk factors and patient behavior and they are potentially reversible. Intensive research of epige‐ netic alterations in human tumors and their association with environmental risk factors and patient behavior put an attention to another group of gene alterations, i.e. on inherited gene polymorphisms. These researches tried to elucidate connection of certain gene polymor‐ phisms with environmental and host risk factors and epigenetic alterations in different hu‐ man tumors and in *HNSCC* [12, 16, 34-36], Figure 2.

Carcinogenesis of *HNSCC* is a multistep process, which arises through an accumulation of genetic and epigenetic alterations. These genetic alterations result in inactivation of multiple tumor suppressor genes and activation of proto-oncogenes by deletions, point mutations, promoter methylation, and gene amplification [3], Table 1.


**Table 1.** Frequent genetic abnormalities in head and neck cancer.

Genetic instability in regions leads to loss of chromosomal region that contains tumor suppres‐ sor genes. A high incidence of Loss of heterozygosity (LOH) observed at 9p, 8p, 3p, 9q, and 11q regions were associated with tumor stage, and poor histological differentiation in *HNSCC* [37]. Loss of heterozygosity (LOH) of 9p21 appears is an early and frequent event in head and neck carcinogenesis [37, 38-40]. The CDKN2A gene locus found in 9p21 region encodes two differ‐ ent transcripts, *p16* and *p14*ARF, which are responsible for G1 cell cycle regulation and MDM2 mediated degradation of *p53*. *P16* and *p14* are often inactivated in *HNSCC* through homozy‐ gous deletion, by promoter methylation, and by point mutations [41-44].

clinical results of these studies [Her-2/neu, *BRCA-1* and *BRCA-2* in Breast Cancer; *K-ras* and *EGFR* in Colon Cancer; *bcr-abl* in Chronic Myeloid Leukemia; *B-RAF* and *c-KIT* in Melano‐ ma; detection of dominant clone of B- and T-lymphocytes in tracking minimal residual dis‐ ease in lymphomas and leukemia's [29-33], still much more questions concerning significance of gene alterations for tumor development and treatment remains unanswered. In the last 10-15 years significant efforts are done to fulfill these gaps in our understanding

of molecular pathogenesis of cancer by studying epigenetic changes of tumor tissues.

man tumors and in *HNSCC* [12, 16, 34-36], Figure 2.

190 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**LOH**

**Mutation**

**Amplification**

promoter methylation, and gene amplification [3], Table 1.

9p21-22/ *p16INK4a/p14ARF*

3p/ *RASSF1A, FHIT, RARB2*

**Table 1.** Frequent genetic abnormalities in head and neck cancer.

These alterations are of special interest because epigenetic changes are tumor/tissue specific, although inheritable epigenetic alterations are under strong influence of environmental risk factors and patient behavior and they are potentially reversible. Intensive research of epige‐ netic alterations in human tumors and their association with environmental risk factors and patient behavior put an attention to another group of gene alterations, i.e. on inherited gene polymorphisms. These researches tried to elucidate connection of certain gene polymor‐ phisms with environmental and host risk factors and epigenetic alterations in different hu‐

Carcinogenesis of *HNSCC* is a multistep process, which arises through an accumulation of genetic and epigenetic alterations. These genetic alterations result in inactivation of multiple tumor suppressor genes and activation of proto-oncogenes by deletions, point mutations,

> *HNSCC OSCC*

> *OSCC HNSCC*

17p13/*p53 HNSCC* 76% 39 11q *OSCC* 20-33% 37 13q14/*Rb HNSCC* 68% 39 8p *OSCC* 53-83% 37

9p21-22/ *p16 OSCC* 70% 41 5q21-22/*APC OSCC* 50% 37 17p13/*p53 HNSCC* 40-79% 49 11p15/*H-ras OSCC* 35-55% 37

11q13/(*PRAD-1/Cyclin D1/hst-1/int-2*) *HNSCC* 30–50% 46, 47 7p12/*EGFR OSCC* 30% 53, 54

70–80% 37, 38,39, 40

30–70% 37, 39, 40, 47

**Genetic changes Locus / gene Cancer type Frequency Ref.**

Loss of chromosome region 3p is a common early genetic event in *HNSCC* [2, 40, 45]. Genet‐ ic alterations at 3p that are common in *HNSCC* and *OSCC* contains tumor suppressor genes Fragile histidine triad (*FHIT*) gene mapped at 3*p14*, Ras association family (*RASSF1A*) gene, mapped at 3p21, and Retinoic acid receptor B2 gene (*RARB2*), mapped to chromosome re‐ gion 3p24 [37, 40]. Amplification of 11q13 leads to overexpression of cyclin D1 [46], detected in 30–60% of *HNSCC* and 40% of cases of oral squamous dysplasia [47].

There has been much research on the tumor suppressor gene *p53* and its role in *HNSCC* and *OSCC* carcinogenesis. The *p53* protein blocks cell division at the G1 to S boundary, stimu‐ lates DNA repair after DNA damage, and induces apoptosis. *P53* has been shown to be functionally inactivated in oral and head and neck tumors [48]. LOH of 17p13, which con‐ tains *p53* and point mutations of the *p53* are seen in more than 50% of *HNSCC* cases [39, 49]. Cigarette smoking has been associated with the mutation of *p53* in head and neck cancers [14]. Investigations about the prognostic value of *p53* status in head and neck cancer sub‐ types hardly can get a clear result due to the presence of large heterogeneity in regard to patients characteristics, and methods for *p53* detection. Meta-analysis of Tandon et al [50] pointed out that evidence about the prognostic value of *p53* in head and neck cancer squa‐ mous cell carcinomas has been inconclusive. However, determination of *p53* status of *HNSCC* may be of particular interests due to its possible role in prediction of the response to cisplatin based chemotherapy. Cisplatin chemotherapy is widely used chemotherapeutical approach in the treatment of head and neck cancers and data that patients with *p53* muta‐ tions respond better to cisplatin chemotherapy need further confirmation [51]. Chemothera‐ py responses in *HNSCC* patients is also influenced by polymorphism at codon 72 of *p53* gene. In patients with wild type *p53*, arginine at codon 72 is associated with better response to chemotherapy. On the contrary, if *p53* is mutated, proline at position 72 is better option [52]. Better understanding of cellular processes and their actors such as *p53* network and their relation to clinical settings will help in appropriate biomarker characterization. Epider‐ mal growth factor receptor (*EGFR*) is overexpressed in 90% of *HNSCC* [53], often due to am‐ plification [54]. Thus, frequent molecular abnormalities in head and neck squamous cell carcinoma include alterations in tumor suppressor genes *p16*INK4A, *p53*, *p14*ARF, *FHIT*, *RASSF1A*, *Rb*, cyclin D1, and activation of oncogenes, such as members of the *ras* gene fami‐ ly, *c-myc*, and *EGFR* [2-4], Table 1.

A large number of previous studies have also suggested the correlation between polymor‐ phisms of genes involved in cell cycle control, angiogenesis and metabolism of alcohol and carcinogens and susceptibility to head and neck cancer [10, 16, 34-36].

Although the impact of genetic changes in oral carcinogenesis is well-known, over the last decade it has been demonstrated that epigenetic changes, especially aberrant DNA methyla‐ tion, play a significant role in *HNSCC* [55]. Epigenetic modifications are heritable changes in gene expression that are not coded in the DNA sequence [56]. These changes are mitoticaly (clonally) heritable and potentially reversible, which provides large possibilities of epigenet‐ ic therapy. Additionally, epigenetic changes could be modulated by the nutrition, environ‐ mental and genetic factors and/or gene-by-environment interactions. Main mechanisms of epigenetic control in mammals include DNA methylation, histone modifications and RNA interference (RNA silencing).

chromatin condensation and suppression of DNA transcription. Thus, DNA methylation

Frequent hypermethylation of tumor-related genes were observed in cancer tissue of *OSCC* and *HNSCC*, in the normal adjacent mucous in the *OSCC* [58], dysplastic tissue [59], and leu‐ koplakia [60]. Promoter hypermethylation can also be detected in buccal swabs samples of tobacco and alcohol users [58], Thus, DNA methylation has been considered as an early

The list of genes that are found to be inactivated by DNA methylation events in *HNSCC* is growing rapidly and includes genes involved in the cell cycle control (*p14*, p15, *p16* and p53), DNA damage repair (*MGMT*, *hMLH1*, and ATM), apoptosis (*DAPK*, *RASSF1A*, and RARβ), *Wnt* signalling (*APC*, *RUNX3*, *WIF1*, *E-cad* and *DCC*) *SFRP* family genes, *TCF21*, etc.

9p21 Regulation of

the *Rb* pathway

apoptosis

*CDH1 E-cad*herin 16q22 Cell–cell adhesion 2-66% 44, 72, 81-88

10q24 Antagonists of

*Wnt* signaling and adhesion

the Wnt pathway

TGF beta-mediated cell cycle arrest

9p21 Pro-apoptosis 18-46% 75-77, 96

*HNSCC methylation*

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**Ref.**

10-70% 42, 96

23-80% 151

18-77% 42, 44, 61,

20-68% 44, 92-95

30-94% 97-99

68, 72, 78

and histone modifications are not isolated events, but highly coordinated [56].

**3.1. DNA methylation in** *HNSCC*

event in head and neck carcinogenesis.

**FUNCTION** *Gene* **Gene name Locus Gene Action**

kinase inhibitor 2B 9p21

protein kinase 1 19q34 p53-dependent

5q21- 22

Cyclin-dependent kinase inhibitor 2A (CDKN2A)

Alternative open reading frame (ARF) of INK4a locus

Death-associated

Adenomatous polyposis coli

Soluble frizzled receptor protein genes family

*p15* Cyclin-dependent

[42, 44, 55, 58-62], Table 2.

*p16*

*p14*

*DAPK1*

*APC*

*SFRP*

**Cell Cycle Control**

**Apoptosis**

*Wnt signaling pathway*

#### **3. DNA methylation**

DNA methylation of cytosine in a CpG dinucleotide is the key epigenetic modifica‐ tion in mammals. The covalent addition of a methyl group to the 5-carbon (C5) po‐ sition of cytosine that are located 5' to a guanosine base in a CpG dinucleotide are catalyzed by a family of enzymes DNA methyltransferases (*DNMTs*). CpG dinucleoti‐ des are asymmetrically distributed in the genome. Throughout the genome rare soli‐ tary CpGs are heavily methylated. In contrast, CpG clustered in small stretches of DNA (0.5-4kb regions), with greater than 50% GC content, are termed 'CpG is‐ lands'. Of approximately 50% the genes in the genome CpG islands are associated with promoter regions. In normal cells, most CpG sites outside of CpG islands are methylated, whereas CpG islands in gene promoters are usually unmethylated, inde‐ pendently of the gene transcription status [56]. During the process of cancerogenesis paradoxical changes in DNA methylation patterns occurs, with simultaneous global hy‐ pomethylation and regional hypermethylation changes. Global DNA hypomethylation may activate, 'unlock' repetitive elements that could affect genome stability or could lead to transcriptional activation of latent viruses or oncogenes. Simultaneously, region‐ al hypermethylation leads to transcriptional silencing of tumor suppressor genes. Cy‐ tosine methylation of CpG islands in the promoters of tumor suppressor genes causes their inactivation, transcriptional silencing and consequently malignant transforma‐ tion. Investigations showed that the number of cancer-related genes that are inactivat‐ ed by epigenetic modifications equals or even exceeds the number of genes inactivated by mutation. Furthermore, genetic and epigenetic changes have almost identical biolog‐ ical effect and pattern of gene expression [56]. Pattern of tumor suppressor genes hy‐ permethylation shows tumor-type specificity.

While *DNMT3A* and *DNMT3B* are mostly involved in *de novo* methylation, *DNMT1* is in‐ volved in the maintenance of DNA methylation after replication [56]. Several studies have shown *DNMT* overexpression (mainly *DNMT1* and *DNMT3B*) in cancer, including *HNSCC* [57]. The analysis of *DNMT* knockout cells revealed that individual enzymes also interact between each other, and interact with other chromatin modifying enzymes, histone deacety‐ lases (*HDAC*) and methyl-CpG binding proteins (MBPs). Methylation of cytosine within CpG islands is associated with binding of MBPs, which recruit *HDAC* to methylated DNA in regions of transcriptional silencing. Histone deacetylation, catalyzed by *HDACs*, leads to the chromatin condensation and suppression of DNA transcription. Thus, DNA methylation and histone modifications are not isolated events, but highly coordinated [56].

#### **3.1. DNA methylation in** *HNSCC*

Although the impact of genetic changes in oral carcinogenesis is well-known, over the last decade it has been demonstrated that epigenetic changes, especially aberrant DNA methyla‐ tion, play a significant role in *HNSCC* [55]. Epigenetic modifications are heritable changes in gene expression that are not coded in the DNA sequence [56]. These changes are mitoticaly (clonally) heritable and potentially reversible, which provides large possibilities of epigenet‐ ic therapy. Additionally, epigenetic changes could be modulated by the nutrition, environ‐ mental and genetic factors and/or gene-by-environment interactions. Main mechanisms of epigenetic control in mammals include DNA methylation, histone modifications and RNA

DNA methylation of cytosine in a CpG dinucleotide is the key epigenetic modifica‐ tion in mammals. The covalent addition of a methyl group to the 5-carbon (C5) po‐ sition of cytosine that are located 5' to a guanosine base in a CpG dinucleotide are catalyzed by a family of enzymes DNA methyltransferases (*DNMTs*). CpG dinucleoti‐ des are asymmetrically distributed in the genome. Throughout the genome rare soli‐ tary CpGs are heavily methylated. In contrast, CpG clustered in small stretches of DNA (0.5-4kb regions), with greater than 50% GC content, are termed 'CpG is‐ lands'. Of approximately 50% the genes in the genome CpG islands are associated with promoter regions. In normal cells, most CpG sites outside of CpG islands are methylated, whereas CpG islands in gene promoters are usually unmethylated, inde‐ pendently of the gene transcription status [56]. During the process of cancerogenesis paradoxical changes in DNA methylation patterns occurs, with simultaneous global hy‐ pomethylation and regional hypermethylation changes. Global DNA hypomethylation may activate, 'unlock' repetitive elements that could affect genome stability or could lead to transcriptional activation of latent viruses or oncogenes. Simultaneously, region‐ al hypermethylation leads to transcriptional silencing of tumor suppressor genes. Cy‐ tosine methylation of CpG islands in the promoters of tumor suppressor genes causes their inactivation, transcriptional silencing and consequently malignant transforma‐ tion. Investigations showed that the number of cancer-related genes that are inactivat‐ ed by epigenetic modifications equals or even exceeds the number of genes inactivated by mutation. Furthermore, genetic and epigenetic changes have almost identical biolog‐ ical effect and pattern of gene expression [56]. Pattern of tumor suppressor genes hy‐

While *DNMT3A* and *DNMT3B* are mostly involved in *de novo* methylation, *DNMT1* is in‐ volved in the maintenance of DNA methylation after replication [56]. Several studies have shown *DNMT* overexpression (mainly *DNMT1* and *DNMT3B*) in cancer, including *HNSCC* [57]. The analysis of *DNMT* knockout cells revealed that individual enzymes also interact between each other, and interact with other chromatin modifying enzymes, histone deacety‐ lases (*HDAC*) and methyl-CpG binding proteins (MBPs). Methylation of cytosine within CpG islands is associated with binding of MBPs, which recruit *HDAC* to methylated DNA in regions of transcriptional silencing. Histone deacetylation, catalyzed by *HDACs*, leads to the

interference (RNA silencing).

192 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**3. DNA methylation**

permethylation shows tumor-type specificity.

Frequent hypermethylation of tumor-related genes were observed in cancer tissue of *OSCC* and *HNSCC*, in the normal adjacent mucous in the *OSCC* [58], dysplastic tissue [59], and leu‐ koplakia [60]. Promoter hypermethylation can also be detected in buccal swabs samples of tobacco and alcohol users [58], Thus, DNA methylation has been considered as an early event in head and neck carcinogenesis.

The list of genes that are found to be inactivated by DNA methylation events in *HNSCC* is growing rapidly and includes genes involved in the cell cycle control (*p14*, p15, *p16* and p53), DNA damage repair (*MGMT*, *hMLH1*, and ATM), apoptosis (*DAPK*, *RASSF1A*, and RARβ), *Wnt* signalling (*APC*, *RUNX3*, *WIF1*, *E-cad* and *DCC*) *SFRP* family genes, *TCF21*, etc. [42, 44, 55, 58-62], Table 2.



poorly differentiated *HNSCC* [73]. *p16* hypermethylation has been associated with poor prognosis in *HNSCC* [55, 74] and *OSCC* [75]. Interestingly, *p16*I*NK4A* and *p14ARF* genes, transcribed from the same locus INK4A by alternate splicing, could have a diametrically op‐ posite clinical effect in oral cancer patients when methylated. Promoter methylation of *p16* was associated with increased disease recurrences and worse prognosis, whereas *p14* meth‐ ylation was strongly associated with lower disease recurrence and was found to be a good prognostic predictor for oral carcinoma [75]. However, other studies associated *p14* methyla‐ tion with tumor stage and lymph node involvement of *OSCC* [76] and poor prognosis [77]. These findings indicate that DNA methylation status of *INK4A/ARF* locus could be used as a prognostic biomarker for assessing the aggressiveness of disease in oral and head and neck

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*Death associated protein-kinase (DAPK)*, plays a critical role in apoptosis regulation in tu‐ mor development, and commonly is hypermethylated in *HNSCC* [42] and *OSCC* [68]. *DAPK* promoter methylation showed the positive correlation with lymph node involve‐ ment [42, 72, 78] and advanced disease stage in *HNSCC* [42, 72]. The presence of *DAPK* promoter hypermethylation detected in surgical margins was associated with the de‐ creased overall survival and was shown to be an independent prognostic factor for over‐

*RASSF1A* Ras association (RalGDS/AF-6) domain family member 1A, involved in RAS path‐ way regulation and tumor suppression, is frequently inactivated by promoter hypermethy‐ lation in *HNSCC* [45, 72]. *RASSF1A* methylation correlates with head and neck tumor stage [72], poor disease-free survival of *OSCC* [79] and lymph node metastasis in nasophageal car‐

*E-cadherin* (*E-cad*) has a dual role in a mammalian cell, as a Ca+2-dependent cell adhesion molecule, and as a part of the complex signaling Wnt pathway interacting with β- and αcatenine [80]. Hypermethylation of *E-cad* is a common event in *HNSCC* [44, 72, 81, 82]. Re‐ duced *E-cad* expression as a consequence of promoter hypermethylation leads to the development of the invasive phenotype in the *OSCC* [83], and is associated with lymph node metastasis in the *OSCC* [84] and *HNSCC* [85]. Hypermethylation of *E-cad* has previous‐ ly been associated with tumor stage of *HNSCC* [72] and nasopharyngeal carcinomas [86] and lymph node metastasis of oral [82] and nasopharyngeal carcinomas [87]. Down-regula‐ tion of CDH1 due to hypermethylation contributed to the progression of esophageal cancer [88], led to poor differentiation and the development of the invasive phenotype in the *OSCC* [83], and salivary gland carcinomas [89]. Furthermore, *E-cad* hypermethylation was associat‐ ed with poor survival in *the OSCC* [84] and *HNSCC* [85]. Recently published investigations showed that *E-cad* gene hypermethylation was associated with the decreased survival in *HNSCC* patients [90]. In addition, advanced *OSCC* patients with *E-cad* promoter methylation had significantly worse 3- and 5-year survival rates [44], and *E-cad* hypermethylation was found to be an independent prognostic factor in oral tongue carcinoma [81]. However, *HNSCC* patients with *E-cad* promoter methylation had lower rates of local recurrences and better disease-specific survival and outcome [91], which indicates that the role of *E-cad* gene

methylation in head and neck carcinomas has not been fully elucidated.

carcinoma patients.

cinomas [78].

all survival in *OSCC* patients [61].

**Table 2.** Genes commonly methylated in *HNSCC*.

#### **3.2. DNA methylation association with progression and prognosis in** *HNSCC*

A number of tumor suppressor genes hypermethylation has been associated with worse outcome in various cancer types. Hypermethylation of the *p16* and *WIF1* genes [63] and *RASSF1A* and *RUNX3* methylation status [64] were found to be independent prognos‐ tic factors non-small cell lung cancers. *DAPK* promoter hypermethylation has been asso‐ ciated with tumor aggressiveness and poor prognosis in lung cancer [65]. *E-cad*herin promoter hypermethylation was found to be the prognostic factor of worse prognosis in diffuse gastric cancer [66], and in non-small cell lung carcinoma [67]. Epigenetic in‐ activation of several genes by DNA methylation has been found to associate with *HNSCC* progression [44, 58, 61, 68].

*p16INK4a* is cyclin-dependent kinase inhibitor which regulates the *Rb* pathway, leading to in‐ hibition of cell cycle progression. An alternate spliced product of the same INK4a locus is another tumor suppressor gene *p14ARF* (Alternative open reading frame). Loss of *p16* in head and neck and oral tumors has been frequently reported [69, 70]. *p16* methylation was correlated with malignant transformation of oral epithelial dysplasia and is a potential bio‐ marker for prediction of prognosis of mild or moderate oral epithelial dysplasia, with the overall sensitivity and specificity of >60% [71]. Previously, *p16* methylation status has been correlated with tumor stage, lymph node metastasis and tumor size in *HNSCC* [72], and poorly differentiated *HNSCC* [73]. *p16* hypermethylation has been associated with poor prognosis in *HNSCC* [55, 74] and *OSCC* [75]. Interestingly, *p16*I*NK4A* and *p14ARF* genes, transcribed from the same locus INK4A by alternate splicing, could have a diametrically op‐ posite clinical effect in oral cancer patients when methylated. Promoter methylation of *p16* was associated with increased disease recurrences and worse prognosis, whereas *p14* meth‐ ylation was strongly associated with lower disease recurrence and was found to be a good prognostic predictor for oral carcinoma [75]. However, other studies associated *p14* methyla‐ tion with tumor stage and lymph node involvement of *OSCC* [76] and poor prognosis [77]. These findings indicate that DNA methylation status of *INK4A/ARF* locus could be used as a prognostic biomarker for assessing the aggressiveness of disease in oral and head and neck carcinoma patients.

**FUNCTION** *Gene* **Gene name Locus Gene Action**

factor 1 12q14

transcription factor 3 1p36

DNA methyltransferase 10q26 DNA repair

*WIF1* Wnt inhibitory

194 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Runt-related

O6-mehylguanine

Ras association (RalGDS/AF-6) domain family member 1A

Deleted in

Retinoic acid

*RUNX3*

*MGMT*

*RASSF1A*

*DCC*

**RARβ**

**Table 2.** Genes commonly methylated in *HNSCC*.

*HNSCC* progression [44, 58, 61, 68].

**DNA damage repair**

**Tumor suppressor** *HNSCC methylation*

Secreted Wnt antagonist

*Wnt* signaling inhibitor, TGF-β-

tumor suppression

for alkylated guanine

RAS pathway regulation and tumor suppression

Colorectal Cancer 18q21 Cell–cell adhesion 17-75% 108, 109

and apoptosis

induced

*hMLH1* human mutL homolog 1 3p21 DNA mismatch repair 8-76% 96, 106, 107

*FHIT* Fragile Histidine Triad 3*p14* Control of cell cycle 33-84% 96, 110

3p21

receptor beta 3p24 Regulatory protein

A number of tumor suppressor genes hypermethylation has been associated with worse outcome in various cancer types. Hypermethylation of the *p16* and *WIF1* genes [63] and *RASSF1A* and *RUNX3* methylation status [64] were found to be independent prognos‐ tic factors non-small cell lung cancers. *DAPK* promoter hypermethylation has been asso‐ ciated with tumor aggressiveness and poor prognosis in lung cancer [65]. *E-cad*herin promoter hypermethylation was found to be the prognostic factor of worse prognosis in diffuse gastric cancer [66], and in non-small cell lung carcinoma [67]. Epigenetic in‐ activation of several genes by DNA methylation has been found to associate with

*p16INK4a* is cyclin-dependent kinase inhibitor which regulates the *Rb* pathway, leading to in‐ hibition of cell cycle progression. An alternate spliced product of the same INK4a locus is another tumor suppressor gene *p14ARF* (Alternative open reading frame). Loss of *p16* in head and neck and oral tumors has been frequently reported [69, 70]. *p16* methylation was correlated with malignant transformation of oral epithelial dysplasia and is a potential bio‐ marker for prediction of prognosis of mild or moderate oral epithelial dysplasia, with the overall sensitivity and specificity of >60% [71]. Previously, *p16* methylation status has been correlated with tumor stage, lymph node metastasis and tumor size in *HNSCC* [72], and

**3.2. DNA methylation association with progression and prognosis in** *HNSCC*

**Ref.**

25-90% 62, 97, 102

18-36% 62, 99-101

10-57% 59, 44, 72,

10-76% 45, 72, 78,

53-88% 59, 78

103-105

79, 96

*Death associated protein-kinase (DAPK)*, plays a critical role in apoptosis regulation in tu‐ mor development, and commonly is hypermethylated in *HNSCC* [42] and *OSCC* [68]. *DAPK* promoter methylation showed the positive correlation with lymph node involve‐ ment [42, 72, 78] and advanced disease stage in *HNSCC* [42, 72]. The presence of *DAPK* promoter hypermethylation detected in surgical margins was associated with the de‐ creased overall survival and was shown to be an independent prognostic factor for over‐ all survival in *OSCC* patients [61].

*RASSF1A* Ras association (RalGDS/AF-6) domain family member 1A, involved in RAS path‐ way regulation and tumor suppression, is frequently inactivated by promoter hypermethy‐ lation in *HNSCC* [45, 72]. *RASSF1A* methylation correlates with head and neck tumor stage [72], poor disease-free survival of *OSCC* [79] and lymph node metastasis in nasophageal car‐ cinomas [78].

*E-cadherin* (*E-cad*) has a dual role in a mammalian cell, as a Ca+2-dependent cell adhesion molecule, and as a part of the complex signaling Wnt pathway interacting with β- and αcatenine [80]. Hypermethylation of *E-cad* is a common event in *HNSCC* [44, 72, 81, 82]. Re‐ duced *E-cad* expression as a consequence of promoter hypermethylation leads to the development of the invasive phenotype in the *OSCC* [83], and is associated with lymph node metastasis in the *OSCC* [84] and *HNSCC* [85]. Hypermethylation of *E-cad* has previous‐ ly been associated with tumor stage of *HNSCC* [72] and nasopharyngeal carcinomas [86] and lymph node metastasis of oral [82] and nasopharyngeal carcinomas [87]. Down-regula‐ tion of CDH1 due to hypermethylation contributed to the progression of esophageal cancer [88], led to poor differentiation and the development of the invasive phenotype in the *OSCC* [83], and salivary gland carcinomas [89]. Furthermore, *E-cad* hypermethylation was associat‐ ed with poor survival in *the OSCC* [84] and *HNSCC* [85]. Recently published investigations showed that *E-cad* gene hypermethylation was associated with the decreased survival in *HNSCC* patients [90]. In addition, advanced *OSCC* patients with *E-cad* promoter methylation had significantly worse 3- and 5-year survival rates [44], and *E-cad* hypermethylation was found to be an independent prognostic factor in oral tongue carcinoma [81]. However, *HNSCC* patients with *E-cad* promoter methylation had lower rates of local recurrences and better disease-specific survival and outcome [91], which indicates that the role of *E-cad* gene methylation in head and neck carcinomas has not been fully elucidated.

*APC* gene plays an integral role in *the Wnt* signaling pathway in binding and degrading of β-catenine. Hypermethylation of *the APC* gene promoter is frequent in *HNSCC* [92-94], and *OSCC* [44]. *APC* methylation status has previously been associated with worse prognosis of esophageal carcinoma [95]. However, in another study on esophageal cancer, hypermethyla‐ tion of the *APC* gene was related to a lower number of metastatic lymph nodes and better recurrence rates [96].

*FHIT* Fragile Histidine Triad gene is associated highly methylated in *HNSCC* [96], and asso‐

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*MINT1* and *MINT 31,* are the members of Methylated in tumor gene family, which methyla‐ tion was previously associated with invasiveness and poor survival in the *OSCC* [109].

A number of recent data suggests that promoter hypermethylation of specific genes does not occur independently or randomly, but concurrently, which indicate that during the tumor progression progressive accumulation of epigenetic alterations could occur. High degree of methylation of multiple genes CpG island regions, associated with microsatellite instability and *hMLH1* gene hypermethylation, has been defined as a CpG island methylator pheno‐ type (CIMP) in colorectal cancer, and is characterized by a poor outcome [111]. It has been suggested that a form of the CpG island methylator phenotype (CIMP) exists in other solid tumors, including *HNSCC* [112]. As opposed of CIMP positive colorectal cancers In oral can‐ cer CIMP is characterized by less aggressive tumor biology [113],. The panel of tumor-relat‐ ed genes used to classify multiple methylation and CIMP differs substantially between studies [12, 112, 113], which could likely affect the results of association with clinical param‐ eters and prognosis. Future extensive investigations are needed to establish a reliable set of reference cancer-related genes whose methylation status should be examined in the specific types of tumors. Further investigations are needed to better characterize the etiology of this methylation phenotype as well as to determine if this phenotype has important prognostic

The detection of aberrant DNA hypermethylation emerged as a potential biomarker strategy for early detection of various carcinomas. Since DNA methylation is an early event in tu‐ morigenesis of *HNSCC*, identification of methylation markers could provide great promise for early detection and treatment in *HNSCC* [68]. As opposed to advanced *HNSCC* diagno‐ sis, early *HNSCC* detection increases survival to 80%. Therefore, early detection markers may potentially serve as a predictive tool for diagnosis and recurrence of *HNSCC*. However, even though such markers provide great promise for early detection, epigenetic biomarkers

Routine oral visual screening could significantly reduce oral cavity mortality [116]. Howev‐ er, oral cavity screening will not identify cancers deep in the pharynx or larynx which re‐ quires special instrumentation and examination. In addition, screening of *HNSCC* should involve noninvasive detection of *OSCC* and *HNSCC*, such as detection in saliva and oral rinses, or minimally invasive detection, such as blood and serum analysis. Saliva presents an ideal means for *HNSCC* biomarker detection because of its proximity to the primary tumor

Soluble CD44 promoter gene hypermethylation detected in saliva samples has been indicat‐ ed as an early detection marker in *HNSCC* screening [119]. Recently, *NID2* and *HOXA9* pro‐ moter hypermethylation have been identified as biomarkers for prevention and early detection in oral carcinoma tissues and saliva [120]. Hypermethylation of multiple tumor-

site, availability of exfoliated cancer cells, and ease of sampling [117, 118].

ciated with poor prognosis in early stage esophageal squamous cell carcinoma [110].

or clinical use.

**3.3. DNA methylation in early detection of** *HNSCC*

have not yet been clinically implemented [114, 115].

*SFRP* (soluble frizzled receptor protein) family genes are involved in the inhibition of *Wnt* signaling. Promoters of SFRP-2, SFRP-4, SFRP-5 genes showed methylation in *OSCC*, where‐ as SFRP-1 was demethylated in oral cancer [97]. SFRP-1 has been associated with tumor grade in *OSCC* [98]. Hypermethylation of SFRP-1 was associated with an increased risk of esophageal cancer recurrence [99].

*RUNX3* Runt-related transcription factor 3 gene plays a role in the transforming growth fac‐ tor-beta (TGF-β) -induced tumor suppression pathway. *RUNX3* may have an oncogenic role in *HNSCC* and its expression may predict malignant behavior [100]. In contrast to *HNSCC*, in *OSCC* the *RUNX3* gene is downregulated due to promoter hypermethylation [101] and associated with lymph node involvement and tumor stage in tongue carcinomas [62]. Hy‐ permethylation of *RUNX3* detected in the plasma of esophageal cancer patients was signifi‐ cantly associated with an increased risk of cancer recurrence [99].

*WIF1* Wnt inhibitory factor 1, gene involved in the inhibition of *Wnt* signaling, is commonly methylated in the *HNSCC* [102] and *OSCC* [62, 97]. Its methylation status was correlated with lymph node metastasis in nasopharyngeal carcinoma [102].

*MGMT* O(6)-methylguanine-DNAmethyltransferase a DNA repair gene that removes muta‐ genic O6 -guanine adducts from DNA, is inactivated by hypermethylation in *HNSCC* [59] and *OSCC* [44]. *MGMT* promoter hypermethylation was associated with tumor stage of *HNSCC* [72] and with lymph node involvement in laryngeal carcinomas [103]. In addition, the methylation of *MGMT* was associated with poor survival and reduced disease-free sur‐ vival in *OSCC* [104] and increased recurrences rate and poor prognosis of *HNSCC* [105].

*hMLH1* mutL homolog 1 is involved in the DNA repair process. The *hMLH1* promoter meth‐ ylation occurred in high frequency of the majority of the early stage *OSCC* and in about half of the late stage carcinomas [106], indicating its potential role in the tumor progression. Pro‐ moter methylation of *hMLH1* gene is also a common event in *HNSCC*, and was correlated with poor survival in *HNSCC* [107].

*RARß* Retinoic acid receptor beta is commonly methylated in *HNSCC* [59] and associated with highly differentiated tumors, advanced tumor stage and the presence of lymph node metastasis of nasopharyngeal carcinomas [78].

*TIMP3* Tissue inhibitor of metalloproteinases 3 gene is involved in the inhibition of angio‐ genesis and tumor growth. Promoter methylation of TIMP3 predicts better outcome in *HNSCC* treated by radiotherapy [91].

*DCC* Deleted in colorectal cancer, tumor suppressor gene highly methylated in *HNSCC* [108] was correlated with mandibular invasion and poor survival in oral cancer [109].

*FHIT* Fragile Histidine Triad gene is associated highly methylated in *HNSCC* [96], and asso‐ ciated with poor prognosis in early stage esophageal squamous cell carcinoma [110].

*MINT1* and *MINT 31,* are the members of Methylated in tumor gene family, which methyla‐ tion was previously associated with invasiveness and poor survival in the *OSCC* [109].

A number of recent data suggests that promoter hypermethylation of specific genes does not occur independently or randomly, but concurrently, which indicate that during the tumor progression progressive accumulation of epigenetic alterations could occur. High degree of methylation of multiple genes CpG island regions, associated with microsatellite instability and *hMLH1* gene hypermethylation, has been defined as a CpG island methylator pheno‐ type (CIMP) in colorectal cancer, and is characterized by a poor outcome [111]. It has been suggested that a form of the CpG island methylator phenotype (CIMP) exists in other solid tumors, including *HNSCC* [112]. As opposed of CIMP positive colorectal cancers In oral can‐ cer CIMP is characterized by less aggressive tumor biology [113],. The panel of tumor-relat‐ ed genes used to classify multiple methylation and CIMP differs substantially between studies [12, 112, 113], which could likely affect the results of association with clinical param‐ eters and prognosis. Future extensive investigations are needed to establish a reliable set of reference cancer-related genes whose methylation status should be examined in the specific types of tumors. Further investigations are needed to better characterize the etiology of this methylation phenotype as well as to determine if this phenotype has important prognostic or clinical use.

#### **3.3. DNA methylation in early detection of** *HNSCC*

*APC* gene plays an integral role in *the Wnt* signaling pathway in binding and degrading of β-catenine. Hypermethylation of *the APC* gene promoter is frequent in *HNSCC* [92-94], and *OSCC* [44]. *APC* methylation status has previously been associated with worse prognosis of esophageal carcinoma [95]. However, in another study on esophageal cancer, hypermethyla‐ tion of the *APC* gene was related to a lower number of metastatic lymph nodes and better

*SFRP* (soluble frizzled receptor protein) family genes are involved in the inhibition of *Wnt* signaling. Promoters of SFRP-2, SFRP-4, SFRP-5 genes showed methylation in *OSCC*, where‐ as SFRP-1 was demethylated in oral cancer [97]. SFRP-1 has been associated with tumor grade in *OSCC* [98]. Hypermethylation of SFRP-1 was associated with an increased risk of

*RUNX3* Runt-related transcription factor 3 gene plays a role in the transforming growth fac‐ tor-beta (TGF-β) -induced tumor suppression pathway. *RUNX3* may have an oncogenic role in *HNSCC* and its expression may predict malignant behavior [100]. In contrast to *HNSCC*, in *OSCC* the *RUNX3* gene is downregulated due to promoter hypermethylation [101] and associated with lymph node involvement and tumor stage in tongue carcinomas [62]. Hy‐ permethylation of *RUNX3* detected in the plasma of esophageal cancer patients was signifi‐

*WIF1* Wnt inhibitory factor 1, gene involved in the inhibition of *Wnt* signaling, is commonly methylated in the *HNSCC* [102] and *OSCC* [62, 97]. Its methylation status was correlated

*MGMT* O(6)-methylguanine-DNAmethyltransferase a DNA repair gene that removes muta‐

and *OSCC* [44]. *MGMT* promoter hypermethylation was associated with tumor stage of *HNSCC* [72] and with lymph node involvement in laryngeal carcinomas [103]. In addition, the methylation of *MGMT* was associated with poor survival and reduced disease-free sur‐ vival in *OSCC* [104] and increased recurrences rate and poor prognosis of *HNSCC* [105].

*hMLH1* mutL homolog 1 is involved in the DNA repair process. The *hMLH1* promoter meth‐ ylation occurred in high frequency of the majority of the early stage *OSCC* and in about half of the late stage carcinomas [106], indicating its potential role in the tumor progression. Pro‐ moter methylation of *hMLH1* gene is also a common event in *HNSCC*, and was correlated

*RARß* Retinoic acid receptor beta is commonly methylated in *HNSCC* [59] and associated with highly differentiated tumors, advanced tumor stage and the presence of lymph node

*TIMP3* Tissue inhibitor of metalloproteinases 3 gene is involved in the inhibition of angio‐ genesis and tumor growth. Promoter methylation of TIMP3 predicts better outcome in

*DCC* Deleted in colorectal cancer, tumor suppressor gene highly methylated in *HNSCC* [108]

was correlated with mandibular invasion and poor survival in oral cancer [109].


cantly associated with an increased risk of cancer recurrence [99].

with lymph node metastasis in nasopharyngeal carcinoma [102].

recurrence rates [96].

genic O6

esophageal cancer recurrence [99].

196 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

with poor survival in *HNSCC* [107].

*HNSCC* treated by radiotherapy [91].

metastasis of nasopharyngeal carcinomas [78].

The detection of aberrant DNA hypermethylation emerged as a potential biomarker strategy for early detection of various carcinomas. Since DNA methylation is an early event in tu‐ morigenesis of *HNSCC*, identification of methylation markers could provide great promise for early detection and treatment in *HNSCC* [68]. As opposed to advanced *HNSCC* diagno‐ sis, early *HNSCC* detection increases survival to 80%. Therefore, early detection markers may potentially serve as a predictive tool for diagnosis and recurrence of *HNSCC*. However, even though such markers provide great promise for early detection, epigenetic biomarkers have not yet been clinically implemented [114, 115].

Routine oral visual screening could significantly reduce oral cavity mortality [116]. Howev‐ er, oral cavity screening will not identify cancers deep in the pharynx or larynx which re‐ quires special instrumentation and examination. In addition, screening of *HNSCC* should involve noninvasive detection of *OSCC* and *HNSCC*, such as detection in saliva and oral rinses, or minimally invasive detection, such as blood and serum analysis. Saliva presents an ideal means for *HNSCC* biomarker detection because of its proximity to the primary tumor site, availability of exfoliated cancer cells, and ease of sampling [117, 118].

Soluble CD44 promoter gene hypermethylation detected in saliva samples has been indicat‐ ed as an early detection marker in *HNSCC* screening [119]. Recently, *NID2* and *HOXA9* pro‐ moter hypermethylation have been identified as biomarkers for prevention and early detection in oral carcinoma tissues and saliva [120]. Hypermethylation of multiple tumorrelated genes (*RAR-β*, *DAPK*, *CDH1*, *p16* and *RASSF1A*) analyzed in combination could serve as a biomarker for early diagnosis of esophageal squamous cell carcinoma [57]. More‐ over, these multiple tumor-related gene hypermethylation were associated with the increase of *DNMT3b* expression in early stages of esophageal cancer [57].

**3.4. Predictive significance of DNA methylation**

ipgenetic agents can reverse drug resistance.

notecan in primary colon cancer [131].

way in human oral carcinoma cells [136].

gery alone [79].

Epigenetic alterations such as DNA methylation could be a marker of response to radio and/or chemotherapy in the treatment of cancer. DNA methylation could play an important role in the regulation of gene expression for genes involved in cell cycle and apoptosis, thus affecting the chemosensitivity of cancers. Treatment of cancer cells with demethylating agent could lead to re-expression of gene involved in the activation of the apoptotic process and restore the sensitivity to the chemotherapy. Targeting the epigenetic mechanisms of apoptosis related genes inactivation may increase the efficacy of chemotherapy in various cancer types. Studies in vitro and in model systems certainly suggest that treatment with ep‐

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199

The best known example of DNA promoter hypermethylation and response to chemothera‐ py is *MGMT* promoter methylation in glioma patients treated with alkylating agents [127]. The expression of DNA repair gene *MGMT*, which removes alkyl groups added to guanine in DNA, is controlled by its promoter methylation. A cell that expresses a low amount of *MGMT* is known to be more sensitive to the antiproliferative effects of alkylating agents. It has been shown that hypermethylation of *MGMT* is the predictor of good response to temo‐ zolomide therapy in gliomas [128], to carmustine therapy in gliomas [127], and to cyclo‐ phosphamide therapy in diffuse B cell lymphoma [129]. Acquired hypermethylation of DNA mismatch repair gene *hMLH1* (detectable in peripheral blood) during carboplatinum/ taxane therapy of ovarian cancer predicts poorer outcome [130]. In addition, *WRN* gene pro‐ moter hypermethylation was associated with hypersensitivity to topoisomerase inhibitor iri‐

It has been shown that DNA methylation could have the predictive significance in *OSCC* and *HNSCC*. Hypermethylation of *MGMT* was the predictor of good response to temozolo‐ mide therapy in oral squamous cell carcinoma [105]. GPx3 promoter hypermethylation was associated with tumorigenesis and chemotherapy response in *HNSCC* [132]. A correlation between methylation of mitotic checkpoint gene CHFR (checkpoint with ring finger) and sensitivity to microtubule inhibitors (docetaxel or paclitaxel) was observed in *OSCC* cells [133]. Downregulation of *SMG-1* (suppressor with morphogenetic effect on genitalia) due to promoter hypermethylation in HPV-Positive *HNSCC* resulted in increased radiation sensi‐ tivity and correlated with improved survival, whereas *SMG-1* overexpression protected HPV-positive tumor cells from irradiation [134]. DNA methylation could be a regulatory mechanism for chemosensitivity to 5-fluorouracil and cisplatin by zebularine, a novel DNA methyltransferase inhibitor, in oral squamous cell carcinomas [135]. It has been shown that zebularine suppresses the apoptotic potential of 5-fluorouracil via cAMP/PKA/CREB path‐

Radiotherapy is the standard adjuvant treatment for *OSCC* and the Ras/PI3K/AKT pathway plays an important role in *OSCC* radioresistance. A combination of *RASSF1A*, *RASSF2A*, and *HIN-1* methylation was found to be significantly associated with poor disease-free sur‐ vival in patients treated with radiotherapy after surgery but not in patients treated with sur‐

The development methylation-specific polymerase chain reaction (PCR) techniques has re‐ sulted in the identification of methylated genes specific to *HNSCC* detected in tumor sam‐ ples [68], including *DAPK* [42], and *RASSF1A* [72].

Saliva and oral rinses could be ideal diagnostic and predictive biofluids for head and neck cancer since they samples cells from the entire lesion and the entire oral cavity. The detec‐ tion of hypermethylated marker genes from oral rinse and saliva samples has a great poten‐ tial for the noninvasive detection of *OSCC* and *HNSCC* [118, 68, 121, 122, 123, 124]. DNA hypermethylation of *p16*, *MGMT*, and *DAPK* in saliva showed aberrant methylation in 56% of samples from *HNSCC* patients, and only in one of the 30 control subjects [68]. The aber‐ rant methylation of a combination of marker genes *E-cadherin*, transmembrane protein with epidermal growth factor-like and 2 follistatin-like domains 2 (*TMEFF2*), and *MGMT*, present in oral rinses was used to detect *OSCC* with >90% sensitivity and specificity [124].

Using the approach of gene selection according to previous studies on promoter methyla‐ tion in *HNSCC* Carvalho et al. analyzed both saliva and serum in 211 *HNSCC* patients and 527 normal controls, and showed high specificity of promoter hypermethylation in *HNSCC* patients compared with normal subjects (>90%); however, the sensitivity of this panel was 31.4% [118]. In another study with different approaches, genome-wide methylation array analysis of 807 cancer-associated genes, was conducted on the matched preoperative saliva, postoperative saliva, and oral cancer tissue, and compared to saliva of normal subjects. Mul‐ tiple potential diagnostic gene panels that consisted of 4 to 7 genes ranged in their sensitivi‐ ty from 62% to 77% and in their specificity from 83% to 100% [123], significantly higher than previous studies of aberrant methylation detected in saliva in head and neck cancer patients [121, 122, 118]. Using this approach, new genes could be discovered that can be used as a reliable biomarker for the early detection of oral and head and neck cancer.

The *KIF1A* (Kinesin family member 1A) gene, that encodes a microtubule-dependent molec‐ ular motor protein involved in organelle transport and cell division, has recently been asso‐ ciated with aberrant DNA methylation in *HNSCC* [125]. *EDNRB* (Endothelin receptor type B) is a G protein coupled receptor, which activates a phosphatidylinositol calcium second messenger system, has previously been associated with nasopharyngeal carcinomas [126] and *HNSCC* [125]. Promoter hypermethylation of *KIF1A* and *EDNRB* is a frequent event in primary *HNSCC*, and these genes are preferentially methylated in salivary rinses from *HNSCC* patients. *KIF1A* (97.8% specificity and 36.6% sensitivity) and *EDNRB* (93.2% specif‐ icity and 67.6% sensitivity) are highly sensitive markers that could potentially be used as bi‐ omarkers for *HNSCC* detection. In addition, combining the markers improves sensitivity while maintains good specificity (93.1% specificity and 77.4% sensitivity) [125].

#### **3.4. Predictive significance of DNA methylation**

related genes (*RAR-β*, *DAPK*, *CDH1*, *p16* and *RASSF1A*) analyzed in combination could serve as a biomarker for early diagnosis of esophageal squamous cell carcinoma [57]. More‐ over, these multiple tumor-related gene hypermethylation were associated with the increase

The development methylation-specific polymerase chain reaction (PCR) techniques has re‐ sulted in the identification of methylated genes specific to *HNSCC* detected in tumor sam‐

Saliva and oral rinses could be ideal diagnostic and predictive biofluids for head and neck cancer since they samples cells from the entire lesion and the entire oral cavity. The detec‐ tion of hypermethylated marker genes from oral rinse and saliva samples has a great poten‐ tial for the noninvasive detection of *OSCC* and *HNSCC* [118, 68, 121, 122, 123, 124]. DNA hypermethylation of *p16*, *MGMT*, and *DAPK* in saliva showed aberrant methylation in 56% of samples from *HNSCC* patients, and only in one of the 30 control subjects [68]. The aber‐ rant methylation of a combination of marker genes *E-cadherin*, transmembrane protein with epidermal growth factor-like and 2 follistatin-like domains 2 (*TMEFF2*), and *MGMT*, present

Using the approach of gene selection according to previous studies on promoter methyla‐ tion in *HNSCC* Carvalho et al. analyzed both saliva and serum in 211 *HNSCC* patients and 527 normal controls, and showed high specificity of promoter hypermethylation in *HNSCC* patients compared with normal subjects (>90%); however, the sensitivity of this panel was 31.4% [118]. In another study with different approaches, genome-wide methylation array analysis of 807 cancer-associated genes, was conducted on the matched preoperative saliva, postoperative saliva, and oral cancer tissue, and compared to saliva of normal subjects. Mul‐ tiple potential diagnostic gene panels that consisted of 4 to 7 genes ranged in their sensitivi‐ ty from 62% to 77% and in their specificity from 83% to 100% [123], significantly higher than previous studies of aberrant methylation detected in saliva in head and neck cancer patients [121, 122, 118]. Using this approach, new genes could be discovered that can be used as a

The *KIF1A* (Kinesin family member 1A) gene, that encodes a microtubule-dependent molec‐ ular motor protein involved in organelle transport and cell division, has recently been asso‐ ciated with aberrant DNA methylation in *HNSCC* [125]. *EDNRB* (Endothelin receptor type B) is a G protein coupled receptor, which activates a phosphatidylinositol calcium second messenger system, has previously been associated with nasopharyngeal carcinomas [126] and *HNSCC* [125]. Promoter hypermethylation of *KIF1A* and *EDNRB* is a frequent event in primary *HNSCC*, and these genes are preferentially methylated in salivary rinses from *HNSCC* patients. *KIF1A* (97.8% specificity and 36.6% sensitivity) and *EDNRB* (93.2% specif‐ icity and 67.6% sensitivity) are highly sensitive markers that could potentially be used as bi‐ omarkers for *HNSCC* detection. In addition, combining the markers improves sensitivity

in oral rinses was used to detect *OSCC* with >90% sensitivity and specificity [124].

reliable biomarker for the early detection of oral and head and neck cancer.

while maintains good specificity (93.1% specificity and 77.4% sensitivity) [125].

of *DNMT3b* expression in early stages of esophageal cancer [57].

ples [68], including *DAPK* [42], and *RASSF1A* [72].

198 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Epigenetic alterations such as DNA methylation could be a marker of response to radio and/or chemotherapy in the treatment of cancer. DNA methylation could play an important role in the regulation of gene expression for genes involved in cell cycle and apoptosis, thus affecting the chemosensitivity of cancers. Treatment of cancer cells with demethylating agent could lead to re-expression of gene involved in the activation of the apoptotic process and restore the sensitivity to the chemotherapy. Targeting the epigenetic mechanisms of apoptosis related genes inactivation may increase the efficacy of chemotherapy in various cancer types. Studies in vitro and in model systems certainly suggest that treatment with ep‐ ipgenetic agents can reverse drug resistance.

The best known example of DNA promoter hypermethylation and response to chemothera‐ py is *MGMT* promoter methylation in glioma patients treated with alkylating agents [127]. The expression of DNA repair gene *MGMT*, which removes alkyl groups added to guanine in DNA, is controlled by its promoter methylation. A cell that expresses a low amount of *MGMT* is known to be more sensitive to the antiproliferative effects of alkylating agents. It has been shown that hypermethylation of *MGMT* is the predictor of good response to temo‐ zolomide therapy in gliomas [128], to carmustine therapy in gliomas [127], and to cyclo‐ phosphamide therapy in diffuse B cell lymphoma [129]. Acquired hypermethylation of DNA mismatch repair gene *hMLH1* (detectable in peripheral blood) during carboplatinum/ taxane therapy of ovarian cancer predicts poorer outcome [130]. In addition, *WRN* gene pro‐ moter hypermethylation was associated with hypersensitivity to topoisomerase inhibitor iri‐ notecan in primary colon cancer [131].

It has been shown that DNA methylation could have the predictive significance in *OSCC* and *HNSCC*. Hypermethylation of *MGMT* was the predictor of good response to temozolo‐ mide therapy in oral squamous cell carcinoma [105]. GPx3 promoter hypermethylation was associated with tumorigenesis and chemotherapy response in *HNSCC* [132]. A correlation between methylation of mitotic checkpoint gene CHFR (checkpoint with ring finger) and sensitivity to microtubule inhibitors (docetaxel or paclitaxel) was observed in *OSCC* cells [133]. Downregulation of *SMG-1* (suppressor with morphogenetic effect on genitalia) due to promoter hypermethylation in HPV-Positive *HNSCC* resulted in increased radiation sensi‐ tivity and correlated with improved survival, whereas *SMG-1* overexpression protected HPV-positive tumor cells from irradiation [134]. DNA methylation could be a regulatory mechanism for chemosensitivity to 5-fluorouracil and cisplatin by zebularine, a novel DNA methyltransferase inhibitor, in oral squamous cell carcinomas [135]. It has been shown that zebularine suppresses the apoptotic potential of 5-fluorouracil via cAMP/PKA/CREB path‐ way in human oral carcinoma cells [136].

Radiotherapy is the standard adjuvant treatment for *OSCC* and the Ras/PI3K/AKT pathway plays an important role in *OSCC* radioresistance. A combination of *RASSF1A*, *RASSF2A*, and *HIN-1* methylation was found to be significantly associated with poor disease-free sur‐ vival in patients treated with radiotherapy after surgery but not in patients treated with sur‐ gery alone [79].

*TGF-β* signaling has been found to be disrupted in *HNSCC* progression [137], thus this path‐ way has been targeted for therapy [138]. Downregulation of disabled homolog 2 (*DAB2*) gene expression via promoter DNA methylation frequently occurs in *HNSCC* and acts as an independent predictor of metastasis and poor prognosis [139]. Epigenetic downregulation of *DAB2* switches *TGF-β* from a tumor suppressor to a tumor promoter, suggest a way to strat‐ ify patients with advanced SCC who may benefit from anti-TGF-β therapies [139].

The epidermal growth factor receptor (*EGFR*) has been extensively investigated and validat‐ ed as a therapeutic target in lung, colorectal, pancreatic, and head and neck cancers. Howev‐ er, patients with wild type *EGFR* obtain little sustained benefit from anti-*EGFR* monotherapy Broad restoration of tumor suppressor function by demethylation could en‐ hance the anti-proliferative and pro-apoptotic effect of *EGFR* blockade in solid malignancy. Re-expression of *p15*, *p21*, or *p27*, cell cycle inhibitors downstream of *EGFR*, or *PTEN*, a PI3K/Akt inhibitory protein, may have particular synergy with anti-*EGFR* therapy. Recent phase I study evaluated the combination of anti-*EGFR* erlotinib and 5-azacytidine in solid carcinomas showed the beneficial clinical effect in lung and head and neck cancers [147]. Ef‐ ficacy of erlotinib could be enhanced by concurrent hypomethylating therapy with 5-azacy‐ tidine, secondary to re-expression of tumor suppressors interacting with the *EGFR* signaling cascade. In recent study of resistance to anti-*EGFR* therapy agents, promoter methylation of commonly methylated genes was investigated in two parental non-small cell lung cancer (*NSCLC*) and *HNSCC* cell lines and their resistant derivatives to either erlotinib or cetuxi‐ mab [148]. It was found that *DAPK* gene promoter was hypermethylated in drug-resistant derivatives generated from both parental cell lines. Restoration of *DAPK* into the resistant *NSCLC* cells by stable transfection re-sensitized the cells to both erlotinib and cetuximab [148], thus indicating that *DAPK* promoter methylation could be a potential biomarker of

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201

These results demonstrate that DNA methylation could play an important role in both che‐ motherapy and radiotherapy resistance, and that gene silencing through promoter methyla‐ tion is one of the key mechanisms of developed resistance to anti-*EGFR* therapeutic agents. Epigenetic therapy could be a novel treatment to overcome chemo- and/or radiotherapy re‐

Epidemiological studies have reported the association of DNA methylation status in *HNSCC* with exposure to environmental factors, including tobacco smoke, alcohol intake, genotoxic

Recent studies have shown a correlation between tobacco and/or alcohol use and hyperme‐ thylation of tumor related genes. DNA global hypomethylation was associated with alcohol consumption [149]. *p16* methylation has been correlated with alcohol use and smoking [72]. Promoter methylation of *p16* was previously correlated with alcohol intake [150], while *RASSF1A* methylation was associated with tobacco use in *HNSCC* [72]. Smoking and drink‐ ing was associated with *p15* promoter methylation in the upper aerodigestive tract of healthy individuals and *HNSCC* patients [151]. Promoter methylation of *SFRP1* occurred more often in both heavy and light drinkers compared to nondrinkers in head and neck squ‐ amous cell carcinoma [152]. Significant association of tumor-associated genes hypermethyla‐

Recent studies are indicating that increased fruit and vegetable consumption are associated with reduced head and neck cancer risk [28]. There is a growing body of evidence that bio‐ active food components, such as isothiocyanates from cruciferous vegetables (cauliflower,

betel quid consumption, HPV infection, diet and environmental pollutants.

sistance and to improve the benefits of current therapies.

**3.6. DNA methylation in** *HNSCC* **and etiological factors**

tion with alcohol use was observed in *HNSCC* [153].

drug response.

Although methylation of certain genes appears to influence the sensitivity to chemothera‐ peutic drugs, the majority of studies were performed on cell line models, or a small number of subjects. In addition, combination therapies of epigenetic agents and standard chemother‐ apy/radiotherapy have to be carefully investigated due to potential harmful effects in the clinical application of *DNMT* inhibitors. Epigenetic profile in predicting the chemosensitivi‐ ty of individual cancers would contribute to personalized therapy [140]. Studies of pharma‐ coepigenomics will require large-scale analyses and genome-wide methylation analyses using microarrays and next-generation sequencers, necessary to confirm the usage of epige‐ netic changes in predicting responses to chemotherapeutic drugs.

#### **3.5. Epigenetic therapy of** *HNSCC*

Unlike mutations, epigenetic changes are reversible, which provides large possibilities of ep‐ igenetic therapy [141]. Demethylating agents that inhibit DNA methyltransferases *DNMTs*, could have a utility to effectively activate expression of previously epigenetically silenced genes. 5-azanucleosides, nucleoside analogs that inhibit *DNMTs*, with consequent hypome‐ thylation of DNA, have long been known to have DNA demethylating activities, but are too toxic for clinical use [141]. However, recent studies have shown that therapeutic efficacy could be achieved at low drug doses [142 -4]. 5-Azacytidine, a cytosine analog treatment has been tested in multiple cancer cell lines and are shown to reexpress methylated genes in myelodysplastic syndromes (MDS) [142, 143]. Low doses were used in a large trial in pa‐ tients with MDS, and showed an increase in the time of conversion of MDS to leukemia, and increased overall survival [144].

5-Azacytidine application resulted in partial demethylation of the *MGMT* and *RASSF1A* tu‐ mor suppressor genes and reduced proliferation of the tumor cells suggesting further inves‐ tigation of 5-azacytidine for *HNSCC* treatment [145]. Methylation status and expression of *HIC1*, a potential tumor suppressor gene, was restored after demethylation treatment of *HNSCC* cell lines with 5-azacytidine [146].

Epigenetic therapy in monotherapy could reactivate tumor suppressor genes or in combina‐ tion therapy may enhance the anti-proliferative effect of standard chemotherapy, such as cisplatin, 5-fluorouracil, etc. The low dose of a novel DNA methyltransferase inhibitor, ze‐ bularine may sensitize oral cancer cells to cisplatin, an important characteristic of solid can‐ cer treatment. DNA methylation could be a regulatory mechanism for dihydropyrimidine dehydrogenase (*DPD)*, known to be a principal factor in 5-fluorouracil (5-FU) resistance and that DPD activated by zebularine in *OSCC* could be an inhibiting factor in the response to apoptosis induced by 5-FU [135].

The epidermal growth factor receptor (*EGFR*) has been extensively investigated and validat‐ ed as a therapeutic target in lung, colorectal, pancreatic, and head and neck cancers. Howev‐ er, patients with wild type *EGFR* obtain little sustained benefit from anti-*EGFR* monotherapy Broad restoration of tumor suppressor function by demethylation could en‐ hance the anti-proliferative and pro-apoptotic effect of *EGFR* blockade in solid malignancy. Re-expression of *p15*, *p21*, or *p27*, cell cycle inhibitors downstream of *EGFR*, or *PTEN*, a PI3K/Akt inhibitory protein, may have particular synergy with anti-*EGFR* therapy. Recent phase I study evaluated the combination of anti-*EGFR* erlotinib and 5-azacytidine in solid carcinomas showed the beneficial clinical effect in lung and head and neck cancers [147]. Ef‐ ficacy of erlotinib could be enhanced by concurrent hypomethylating therapy with 5-azacy‐ tidine, secondary to re-expression of tumor suppressors interacting with the *EGFR* signaling cascade. In recent study of resistance to anti-*EGFR* therapy agents, promoter methylation of commonly methylated genes was investigated in two parental non-small cell lung cancer (*NSCLC*) and *HNSCC* cell lines and their resistant derivatives to either erlotinib or cetuxi‐ mab [148]. It was found that *DAPK* gene promoter was hypermethylated in drug-resistant derivatives generated from both parental cell lines. Restoration of *DAPK* into the resistant *NSCLC* cells by stable transfection re-sensitized the cells to both erlotinib and cetuximab [148], thus indicating that *DAPK* promoter methylation could be a potential biomarker of drug response.

These results demonstrate that DNA methylation could play an important role in both che‐ motherapy and radiotherapy resistance, and that gene silencing through promoter methyla‐ tion is one of the key mechanisms of developed resistance to anti-*EGFR* therapeutic agents. Epigenetic therapy could be a novel treatment to overcome chemo- and/or radiotherapy re‐ sistance and to improve the benefits of current therapies.

#### **3.6. DNA methylation in** *HNSCC* **and etiological factors**

*TGF-β* signaling has been found to be disrupted in *HNSCC* progression [137], thus this path‐ way has been targeted for therapy [138]. Downregulation of disabled homolog 2 (*DAB2*) gene expression via promoter DNA methylation frequently occurs in *HNSCC* and acts as an independent predictor of metastasis and poor prognosis [139]. Epigenetic downregulation of *DAB2* switches *TGF-β* from a tumor suppressor to a tumor promoter, suggest a way to strat‐

Although methylation of certain genes appears to influence the sensitivity to chemothera‐ peutic drugs, the majority of studies were performed on cell line models, or a small number of subjects. In addition, combination therapies of epigenetic agents and standard chemother‐ apy/radiotherapy have to be carefully investigated due to potential harmful effects in the clinical application of *DNMT* inhibitors. Epigenetic profile in predicting the chemosensitivi‐ ty of individual cancers would contribute to personalized therapy [140]. Studies of pharma‐ coepigenomics will require large-scale analyses and genome-wide methylation analyses using microarrays and next-generation sequencers, necessary to confirm the usage of epige‐

Unlike mutations, epigenetic changes are reversible, which provides large possibilities of ep‐ igenetic therapy [141]. Demethylating agents that inhibit DNA methyltransferases *DNMTs*, could have a utility to effectively activate expression of previously epigenetically silenced genes. 5-azanucleosides, nucleoside analogs that inhibit *DNMTs*, with consequent hypome‐ thylation of DNA, have long been known to have DNA demethylating activities, but are too toxic for clinical use [141]. However, recent studies have shown that therapeutic efficacy could be achieved at low drug doses [142 -4]. 5-Azacytidine, a cytosine analog treatment has been tested in multiple cancer cell lines and are shown to reexpress methylated genes in myelodysplastic syndromes (MDS) [142, 143]. Low doses were used in a large trial in pa‐ tients with MDS, and showed an increase in the time of conversion of MDS to leukemia, and

5-Azacytidine application resulted in partial demethylation of the *MGMT* and *RASSF1A* tu‐ mor suppressor genes and reduced proliferation of the tumor cells suggesting further inves‐ tigation of 5-azacytidine for *HNSCC* treatment [145]. Methylation status and expression of *HIC1*, a potential tumor suppressor gene, was restored after demethylation treatment of

Epigenetic therapy in monotherapy could reactivate tumor suppressor genes or in combina‐ tion therapy may enhance the anti-proliferative effect of standard chemotherapy, such as cisplatin, 5-fluorouracil, etc. The low dose of a novel DNA methyltransferase inhibitor, ze‐ bularine may sensitize oral cancer cells to cisplatin, an important characteristic of solid can‐ cer treatment. DNA methylation could be a regulatory mechanism for dihydropyrimidine dehydrogenase (*DPD)*, known to be a principal factor in 5-fluorouracil (5-FU) resistance and that DPD activated by zebularine in *OSCC* could be an inhibiting factor in the response to

ify patients with advanced SCC who may benefit from anti-TGF-β therapies [139].

netic changes in predicting responses to chemotherapeutic drugs.

200 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**3.5. Epigenetic therapy of** *HNSCC*

increased overall survival [144].

apoptosis induced by 5-FU [135].

*HNSCC* cell lines with 5-azacytidine [146].

Epidemiological studies have reported the association of DNA methylation status in *HNSCC* with exposure to environmental factors, including tobacco smoke, alcohol intake, genotoxic betel quid consumption, HPV infection, diet and environmental pollutants.

Recent studies have shown a correlation between tobacco and/or alcohol use and hyperme‐ thylation of tumor related genes. DNA global hypomethylation was associated with alcohol consumption [149]. *p16* methylation has been correlated with alcohol use and smoking [72]. Promoter methylation of *p16* was previously correlated with alcohol intake [150], while *RASSF1A* methylation was associated with tobacco use in *HNSCC* [72]. Smoking and drink‐ ing was associated with *p15* promoter methylation in the upper aerodigestive tract of healthy individuals and *HNSCC* patients [151]. Promoter methylation of *SFRP1* occurred more often in both heavy and light drinkers compared to nondrinkers in head and neck squ‐ amous cell carcinoma [152]. Significant association of tumor-associated genes hypermethyla‐ tion with alcohol use was observed in *HNSCC* [153].

Recent studies are indicating that increased fruit and vegetable consumption are associated with reduced head and neck cancer risk [28]. There is a growing body of evidence that bio‐ active food components, such as isothiocyanates from cruciferous vegetables (cauliflower, cabbage, and broccoli), diallyl sulfide, an organosulphur compound from garlic, isoflavone, phytosterole, folate, selenium, vitamin E, flavonoids might reduce cancer risk through epi‐ genetic mechanisms. The chemoprevention of cancer by natural compounds could be a promising approach with less side effects and toxicity. Main polyphenols with the proper‐ ties of *DNMT* inhibition are tea polyphenols, soy isoflavones, organosulphur compounds from garlic, and isothiocyanates from cruciferous vegetables. Dietary folate intake was asso‐ ciated with *p16* promoter methylation in head and neck squamous cell carcinoma [154]. Epi‐ gallocatechin-3-gallate (*EGCG*), the major polyphenol in green tea, is believed to be a key active ingredient with anti-cancer properties. *EGCG* is methylated by catechol-O-methyl‐ transferase and inhibits DNA methyltransferase (*DNMT*), thus reversing the hypermethyla‐ tion and inducing the re-expression of the silenced genes [141].

**Author details**

**References**

tics. *CA Cancer J Clin.*, 57(1).

*Engl J Med.*, 345(26), 1890-900.

pharyngeal cancer. *Ann Oncol.*, 14, 1677-1681.

meta-analysis. *Am J Epidemiol.*, 173(8), 847-57.

set of head and neck cancers. *J Natl Cancer Inst.*, 92, 709-720.

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Zvonko Magić1,2\*, Gordana Supić1,2, Mirjana Branković-Magić<sup>3</sup>

3 Institute for Oncology and Radiology of Serbia, Belgrade, Serbia

1 Institute for Medical Research, Military Medical Academy, Belgrade, Serbia

4 Clinic for Maxillofacial Surgery, Military Medical Academy, Belgrade, Serbia

2 Faculty of Medicine, Military Medical Academy, University of Defense, Belgrade, Serbia

[1] Jemal, A., Siegel, R., Ward, E., Murray, T., Xu, J., & Thun, M. J. (2007). Cancer statis‐

[2] Perez-Ordonez, B., Beauchemin, M., & Jordan, R. (2006). Molecular biology of squa‐

[3] Williams, H.K. (2000). Molecular pathogenesis of oral squamous carcinoma. *Mol*

[4] Forastiere, A., Koch, W., Trotti, A., & Sidransky, D. (2001). Head and neck cancer. *N*

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[6] Pelucchi, C., Talamini, R., Negri, E., et al. (2003). Folate intake and risk of oral and

[8] Gillison, M. L., Koch, W. M., Capone, R. B., Spafford, M., Westra, W. H., Wu, L., et al. (2000). Evidence for a causal association between human papillomavirus and a sub‐

[9] Doll, R. (1998). Uncovering the risks of smoking: historical perspective. *Stat. Meth.*

[10] Zhang, Z. J., Hao, K., Shi, R., Zhao, G., Jiang, G. X., Song, Y., Xu, X., & Ma, J. (2011). Glutathione S-transferase M1 (GSTM1) and glutathione S-transferase T1 (GSTT1) null polymorphisms, smoking, and their interaction in oral cancer: a HuGE review and

[7] Pöschl, G., & Seitz, H. K. (2004). Alcohol and cancer. *Alcohol Alcohol.*, 39(3), 155-65.

mous cell carcinoma of the head and neck. *J. Clin Pathol*, 57, 445-453.

\*Address all correspondence to: zvonkomag@yahoo.co.uk

and Nebojša Jović4,2

http://dx.doi.org/10.5772/51169

203

DNA Methylation in the Pathogenesis of Head and Neck Cancer

*EGCG* has been reported to reverse hypermethylation and reactivate several tumor suppres‐ sor genes in human esophageal squamous cell carcinoma cell lines [155]. The reversion-in‐ ducing cysteine-rich protein with Kazal motifs (*RECK*), a novel matrix metalloproteinases (*MMP*) inhibitor, is involved in the inhibition of tumor angiogenesis, invasion, and metasta‐ sis. *EGCG* treatment enhanced *RECK* expression by reversal of hypermethylation of *RECK* promoter and inhibiting *MMP* activities and invasion in *OSCC* cell lines [156]. Genistein, soy dietary isoflavone, reversed DNA hypermethylation and reactivated *RARβ*, *p16*, and *MGMT* in esophageal cancer cells [157]. However, the findings of association of diet and epigenetic modifications in the head and cancer and other carcinomas are still confounding.

#### **4. Conclusions**

Head and neck squamous cell carcinoma (*HNSCC*) is one of the most common and highly aggressive malignancies worldwide, despite the significant efforts committed in the last dec‐ ades in its detection, prevention and treatment. Therefore, early detection and better disease prediction via genetic and epigenetic biomarkers is crucial. However, very few reliable markers are currently known. Recent studies provide strong evidence that DNA methyla‐ tion could have an important role in head and neck cancer. The detection of promoter meth‐ ylation status may be a useful molecular marker for early detection of *HNSCC* from tissue, saliva, and serum samples and in real time analysis of margins during surgery. In addition, the creation of methylation gene panels could be useful for *HNSCC* screening in the timely inclusion of treatment and thorough surveillance during follow-up period. Unlike genetic changes, epigenetic modifications are heritable and potentially reversible, thus providing the potential for therapy. Frequent DNA methylation detected in *HNSCC* and association with tumor progression and survival indicates that DNA methylation plays an important role in head and neck carcinogenesis and may be a useful diagnostic marker and a potential therapeutic target for *HNSCC*.

#### **Author details**

cabbage, and broccoli), diallyl sulfide, an organosulphur compound from garlic, isoflavone, phytosterole, folate, selenium, vitamin E, flavonoids might reduce cancer risk through epi‐ genetic mechanisms. The chemoprevention of cancer by natural compounds could be a promising approach with less side effects and toxicity. Main polyphenols with the proper‐ ties of *DNMT* inhibition are tea polyphenols, soy isoflavones, organosulphur compounds from garlic, and isothiocyanates from cruciferous vegetables. Dietary folate intake was asso‐ ciated with *p16* promoter methylation in head and neck squamous cell carcinoma [154]. Epi‐ gallocatechin-3-gallate (*EGCG*), the major polyphenol in green tea, is believed to be a key active ingredient with anti-cancer properties. *EGCG* is methylated by catechol-O-methyl‐ transferase and inhibits DNA methyltransferase (*DNMT*), thus reversing the hypermethyla‐

*EGCG* has been reported to reverse hypermethylation and reactivate several tumor suppres‐ sor genes in human esophageal squamous cell carcinoma cell lines [155]. The reversion-in‐ ducing cysteine-rich protein with Kazal motifs (*RECK*), a novel matrix metalloproteinases (*MMP*) inhibitor, is involved in the inhibition of tumor angiogenesis, invasion, and metasta‐ sis. *EGCG* treatment enhanced *RECK* expression by reversal of hypermethylation of *RECK* promoter and inhibiting *MMP* activities and invasion in *OSCC* cell lines [156]. Genistein, soy dietary isoflavone, reversed DNA hypermethylation and reactivated *RARβ*, *p16*, and *MGMT* in esophageal cancer cells [157]. However, the findings of association of diet and epigenetic

Head and neck squamous cell carcinoma (*HNSCC*) is one of the most common and highly aggressive malignancies worldwide, despite the significant efforts committed in the last dec‐ ades in its detection, prevention and treatment. Therefore, early detection and better disease prediction via genetic and epigenetic biomarkers is crucial. However, very few reliable markers are currently known. Recent studies provide strong evidence that DNA methyla‐ tion could have an important role in head and neck cancer. The detection of promoter meth‐ ylation status may be a useful molecular marker for early detection of *HNSCC* from tissue, saliva, and serum samples and in real time analysis of margins during surgery. In addition, the creation of methylation gene panels could be useful for *HNSCC* screening in the timely inclusion of treatment and thorough surveillance during follow-up period. Unlike genetic changes, epigenetic modifications are heritable and potentially reversible, thus providing the potential for therapy. Frequent DNA methylation detected in *HNSCC* and association with tumor progression and survival indicates that DNA methylation plays an important role in head and neck carcinogenesis and may be a useful diagnostic marker and a potential

modifications in the head and cancer and other carcinomas are still confounding.

tion and inducing the re-expression of the silenced genes [141].

202 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**4. Conclusions**

therapeutic target for *HNSCC*.

Zvonko Magić1,2\*, Gordana Supić1,2, Mirjana Branković-Magić<sup>3</sup> and Nebojša Jović4,2


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from a tumor suppressor to a tumor promoter. *J Clin Invest.*, 120(8), 2842-57.

with tumorigenesis and chemotherapy response. *Cancer Lett.*, 309(1), 37-45.

ovarian cancer patients. *Clin Cancer Res.*, 10(13), 4420-6.

214 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

to microtubule inhibitors. *Cancer Biol Ther.*, 4(7), 773-80.

squamous cell carcinoma. *Int J Oncol*, 31(6), 1449-56.

sis: role of TGF-beta1. *Carcinogenesis.*, 23, 1569-75.

cancer therapy. *Cancer Sci.*, 100(5), 787-91.

tients as related to prognosis. *J Oral Pathol Med.*, 32(3), 139-45.

vival. *Clin Cancer Res.*, 18(5), 1257-1267.

*macol*, 64(2), 223-32.

8822-7.


but not SLC5A8, in head and neck squamous cell carcinomas. *Genes Chromosomes Cancer*, 49, 319-26.

**Section 5**

**Bacteria, Viruses and Metals Methylation: Risk**

**and Benefit for Human Health**


**Bacteria, Viruses and Metals Methylation: Risk and Benefit for Human Health**

but not SLC5A8, in head and neck squamous cell carcinomas. *Genes Chromosomes*

[154] Kraunz, K. S., Hsiung, D., Mc Clean, M. D., Liu, M., Osanyingbemi, J., Nelson, H. H., & Kelsey, K. T. (2006). Dietary folate is associated with p16 (INK4A) methylation in

[155] Fang, M. Z., Wang, Y. W., Ai, N., Hou, Z., Sun, Y., Lu, H., Welsh, W., & Yang, C. S. (2003). Tea polyphenol (\_)-epigallocatechin-3-gallate inhibits DNA methyltransferase and reactivates methylation-silenced genes in cancer cell lines. *Cancer Res.*, 63,

[156] Kato, K., Long, N. K., Makita, H., Toida, M., Yamashita, T., Hatakeyama, D., Hara, A., Mori, H., & Shibata, T. (2008). Effects of green tea polyphenol on methylation sta‐ tus of RECK gene and cancer cell invasion in oral squamous cell carcinoma cells. *Br J*

[157] Fang, M. Z., Chen, D., Sun, Y., Jin, Z., Christman, J. K., & Yang, C. S. (2005). Reversal of hypermethylation and reactivation of 16INK4aRARbeta, and MGMT genes by

genistein and other isoflavones from soy. *Clin Cancer Res.*, 11(19 Pt 1), 7033-41.

head and neck squamous cell carcinoma. *Int J Cancer.*, 119(7), 1553-7.

*Cancer*, 49, 319-26.

216 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

7563-7570.

*Cancer*, 99(4), 647-54.

**Chapter 9**

**Host-Mimicking Strategies in DNA Methylation for**

In 1928, Griffith [1] reported that soluble substances from virulent pneumococcal cells trans‐ formed non-virulent pneumococcus to virulent forms. This substance has now been demon‐ strated to be DNA [2-4]. This is considered to be the first report on genetic transformation of bacteria by exogenous DNA. Subsequently, natural competence of *Bacillus subtilis* was re‐ ported in 1958 by Young and Spizizen [5]. They also demonstrated genetic transformation of natural competent *B. subtilis* cells using exogenous DNA. It was in 1970 that genetic trans‐ formation of *Escherichia coli* using chemically competent cells was reported [6]. Thus, genetic transformation of common bacterial models was established at an early stage in the develop‐ ment of bacteriology. The alternative view is that bacterial models such as *B. subtilis* and *E. coli* have become the mainstay of this field because of high transformation ability. Genetic transformation techniques remain important for studying numerous bacteria and for the ad‐ vancement of bacteriology, biochemistry, applied microbiology, and microbial biotechnolo‐ gy. Moreover, recent developments in the search for new bacteria and genome sequencing have provided numerous effective bacteria that are useful for biological studies and indus‐ trial applications. With these developments, there is a greater demand for establishing ge‐

DNA introduction is an essential process for transforming target bacterium by exogenous DNA. Various methods for introducing DNA into bacteria have been developed to date, in‐ cluding chemotransformation, electroporation, sonopolation, tribos, and conjugational transfer [7]. In spite of these developments, it is often difficult to establish transformation methods for target bacterium. A possible reason is the difficulty faced while exploring suita‐ ble conditions for introducing DNA, which requires not only theoretical understanding but also a trial and error approach. Circumventing bacterial RM systems is a major challenge.

> © 2013 Suzuki; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

**Improved Bacterial Transformation**

Additional information is available at the end of the chapter

netic transformation methods for more bacteria.

Hirokazu Suzuki

**1. Introduction**

http://dx.doi.org/10.5772/51691

### **Host-Mimicking Strategies in DNA Methylation for Improved Bacterial Transformation**

Hirokazu Suzuki

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51691

#### **1. Introduction**

In 1928, Griffith [1] reported that soluble substances from virulent pneumococcal cells trans‐ formed non-virulent pneumococcus to virulent forms. This substance has now been demon‐ strated to be DNA [2-4]. This is considered to be the first report on genetic transformation of bacteria by exogenous DNA. Subsequently, natural competence of *Bacillus subtilis* was re‐ ported in 1958 by Young and Spizizen [5]. They also demonstrated genetic transformation of natural competent *B. subtilis* cells using exogenous DNA. It was in 1970 that genetic trans‐ formation of *Escherichia coli* using chemically competent cells was reported [6]. Thus, genetic transformation of common bacterial models was established at an early stage in the develop‐ ment of bacteriology. The alternative view is that bacterial models such as *B. subtilis* and *E. coli* have become the mainstay of this field because of high transformation ability. Genetic transformation techniques remain important for studying numerous bacteria and for the ad‐ vancement of bacteriology, biochemistry, applied microbiology, and microbial biotechnolo‐ gy. Moreover, recent developments in the search for new bacteria and genome sequencing have provided numerous effective bacteria that are useful for biological studies and indus‐ trial applications. With these developments, there is a greater demand for establishing ge‐ netic transformation methods for more bacteria.

DNA introduction is an essential process for transforming target bacterium by exogenous DNA. Various methods for introducing DNA into bacteria have been developed to date, in‐ cluding chemotransformation, electroporation, sonopolation, tribos, and conjugational transfer [7]. In spite of these developments, it is often difficult to establish transformation methods for target bacterium. A possible reason is the difficulty faced while exploring suita‐ ble conditions for introducing DNA, which requires not only theoretical understanding but also a trial and error approach. Circumventing bacterial RM systems is a major challenge.

© 2013 Suzuki; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

These systems defend bacteria against transformation by exogenous DNA, such as bacterio‐ phages, and effectively hamper genetic transformation by exogenous plasmids. RM systems selectively digest exogenous DNA by differentiating them from host-endogenous DNA on the basis of host-specific DNA methylation [8]. Therefore, DNA that imitates the methyla‐ tion patterns of the host bacterium (host-mimicking DNA) is incorporated into the bacteri‐ um without restriction. Thus, RM systems can be overcome by theoretical host-mimicking strategies (Figure 1), rather than exploring conditions for introducing DNA. This chapter ex‐ plains host-mimicking strategies and provides tips for establishing transformation methods for new bacteria.

RM systems selectively digest exogenous DNA and greatly influence the efficiency of genet‐ ic transformation by exogenous DNA. Numerous RM systems have been found in bacteria and archaea, and are classified into four main types (type I–IV) [10]. Their general properties are summarized in Table 1. The REBASE database (rebase.neb.com) [11] has accumulated large amounts of information about RM systems, including types, gene sequences, recogni‐ tion sites, and origin organisms. Among the four types of RM systems, type II consists of RE and MT. In this type, RE cuts exogenous DNA at specific sites but not endogenous DNA that has been methylated by MT (Figure 1). Type I and III systems also cut exogenous DNA by similar mechanisms, but comprise protein complexes of some subunits. Type IV is known as a modification-specific restriction system to cut DNA with heterologous modifica‐ tions. Intriguingly, RM systems behave as selfish elements like viruses and transposons [12], implying that RM systems have been irreversibly distributed in bacteria. The following sec‐

Host-Mimicking Strategies in DNA Methylation for Improved Bacterial Transformation

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221

Several DNA modifications have been elucidated and in almost all cases the modifica‐ tions that are involved in restriction are nucleobase methylations. The main forms of me‐


> **Restriction Methylation Machinery Cleavage site Machinery Nucleobase**

Type I R2M2S Variable M2S 6mA Type II RE Fixed MT 6mA, 5mC, 4mC Type III R2M2 Variable M2 6mA

**Table 1.** General properties of four types of RM systems.Type I comprises R, M, and S subunits. Methylation is performed by subunits M and S. Type III comprises R and M subunits. The M subunit alone catalyzes methylation. Type II comprises two independent proteins, RE and MT. Type IV comprises only RE and restricts DNA with heterologous


tions describe more details of RM systems.

thylated nucleobases are *N* <sup>6</sup>

or ghmC have been reported [19-21].

Type IV RE Variable

modifications. Methylation produces 6mA, 5mC, or 4mC.

**RM system**

*N* <sup>4</sup>

**2.1. DNA modifications involved in DNA restriction**

**Figure 1.** Host-mimicking strategies for circumventing restriction–modification (RM) systems in bacteria.RM systems serve to defend bacteria against invasion by exogenous DNA, and thereby hamper genetic transformation by exoge‐ nous plasmids. In typical RM systems, restriction endonuclease (RE) digests exogenous DNA but not endogenous DNA that has been methylated by cognate DNA methyltransferase (MT). Host-mimicking DNA that imitates the methyla‐ tion patterns of the bacterial host is recognized as endogenous DNA by the host because RM systems depend on hostspecific DNA methylation to distinguish between exogenous and endogenous DNA.

#### **2. RM systems**

In 1962, Arber and Dussoix [9] found that bacteriophage λ carried specificity for *E. coli* strains in which they were produced. For examples, bacteriophage λ from *E. coli* K-12 was efficiently transfected into K-12 and C strains, but not strains B and K(P1) (efficiencies were <10−4 fold). Meanwhile, bacteriophage λ produced by *E. coli* C efficiently transformed strain C, but not strains K-12, B, and K(P1) (efficiencies were <4×10−4 fold). Thus, bacteriophage λ vectors readily infected *E. coli* strains that produced them but the infection to other strains was "restricted". This occurrence, termed restriction, is explained by *E. coli* RM systems that act as host defense against exogenous DNA.

RM systems selectively digest exogenous DNA and greatly influence the efficiency of genet‐ ic transformation by exogenous DNA. Numerous RM systems have been found in bacteria and archaea, and are classified into four main types (type I–IV) [10]. Their general properties are summarized in Table 1. The REBASE database (rebase.neb.com) [11] has accumulated large amounts of information about RM systems, including types, gene sequences, recogni‐ tion sites, and origin organisms. Among the four types of RM systems, type II consists of RE and MT. In this type, RE cuts exogenous DNA at specific sites but not endogenous DNA that has been methylated by MT (Figure 1). Type I and III systems also cut exogenous DNA by similar mechanisms, but comprise protein complexes of some subunits. Type IV is known as a modification-specific restriction system to cut DNA with heterologous modifica‐ tions. Intriguingly, RM systems behave as selfish elements like viruses and transposons [12], implying that RM systems have been irreversibly distributed in bacteria. The following sec‐ tions describe more details of RM systems.

#### **2.1. DNA modifications involved in DNA restriction**

These systems defend bacteria against transformation by exogenous DNA, such as bacterio‐ phages, and effectively hamper genetic transformation by exogenous plasmids. RM systems selectively digest exogenous DNA by differentiating them from host-endogenous DNA on the basis of host-specific DNA methylation [8]. Therefore, DNA that imitates the methyla‐ tion patterns of the host bacterium (host-mimicking DNA) is incorporated into the bacteri‐ um without restriction. Thus, RM systems can be overcome by theoretical host-mimicking strategies (Figure 1), rather than exploring conditions for introducing DNA. This chapter ex‐ plains host-mimicking strategies and provides tips for establishing transformation methods

220 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 1.** Host-mimicking strategies for circumventing restriction–modification (RM) systems in bacteria.RM systems serve to defend bacteria against invasion by exogenous DNA, and thereby hamper genetic transformation by exoge‐ nous plasmids. In typical RM systems, restriction endonuclease (RE) digests exogenous DNA but not endogenous DNA that has been methylated by cognate DNA methyltransferase (MT). Host-mimicking DNA that imitates the methyla‐ tion patterns of the bacterial host is recognized as endogenous DNA by the host because RM systems depend on host-

In 1962, Arber and Dussoix [9] found that bacteriophage λ carried specificity for *E. coli* strains in which they were produced. For examples, bacteriophage λ from *E. coli* K-12 was efficiently transfected into K-12 and C strains, but not strains B and K(P1) (efficiencies were <10−4 fold). Meanwhile, bacteriophage λ produced by *E. coli* C efficiently transformed strain C, but not strains K-12, B, and K(P1) (efficiencies were <4×10−4 fold). Thus, bacteriophage λ vectors readily infected *E. coli* strains that produced them but the infection to other strains was "restricted". This occurrence, termed restriction, is explained by *E. coli* RM systems that

specific DNA methylation to distinguish between exogenous and endogenous DNA.

act as host defense against exogenous DNA.

for new bacteria.

**2. RM systems**

Several DNA modifications have been elucidated and in almost all cases the modifica‐ tions that are involved in restriction are nucleobase methylations. The main forms of me‐ thylated nucleobases are *N* <sup>6</sup> -methyladenine (6mA; Figure 2), 5-methylcytosine (5mC), and *N* <sup>4</sup> -methylcytosine (4mC). These modifications are performed for double-stranded DNA using methyltransferases or other methyltransfer machinery. In addition, some bacterio‐ phage DNA contain 5-hydroxymethylcytosine (5hmC) instead of cytosine. This modifica‐ tion is incorporated during phage DNA replication using 5-hydroxymethyldeoxycytidine triphosphate as the substrate [13]. The hydroxyl group is further glucosylated to pro‐ duce β-glucosyl-5-hydroxymethylcytosine (ghmC) in a phage-specific pattern by glucosyltrans‐ ferase [14]. This modification has various biological functions, including circumvention of restriction barriers of RM systems in phage hosts [13, 14]. The modified cytosine 5hmC is also found in mammalians [15-17]. Unlike phages, it is produced by oxidation of 5mC in double-stranded DNA [18]. In bacteria, there are no known type I–III RM systems involv‐ ing 5hmC or ghmC. However, several type IV systems that restrict DNA containing 5hmC and/ or ghmC have been reported [19-21].


**Table 1.** General properties of four types of RM systems.Type I comprises R, M, and S subunits. Methylation is performed by subunits M and S. Type III comprises R and M subunits. The M subunit alone catalyzes methylation. Type II comprises two independent proteins, RE and MT. Type IV comprises only RE and restricts DNA with heterologous modifications. Methylation produces 6mA, 5mC, or 4mC.

activity. In all cases reported so far, methylation by type I RM systems occurs in adenine to

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223

Type II RM systems are extremely diverse and are currently classified into 11 subfamilies [8]. Generally, these comprise two enzymes, RE and MT. Cleavages of exogenous DNA at unmethylated recognition sites is carried out by RE, which spares endogenous DNA that has been methylated by the cognate MT. Most REs require Mg2+ ions as a cofactor for cleav‐ age. Although RE may form monomers, dimers, or tetramers, it functions without forming complexes with the cognate MT. The recognition of cleavage sites is highly precise and rec‐ ognition sequences are often palindromic. Such sites include 5′-GAATTC-3′ for EcoRI and 5′-GGATCC-3′ for BamHI, which are cleaved symmetrically within the sites. Because of these useful properties, more than 3,500 REs have been characterized, and many are widely utilized in recombinant DNA technology [8]. The enzyme MT catalyzes methylation at rec‐ ognition sites using *S*-adenosyl-L-methionine as the substrate and generally acts as a mono‐ mer. The nucleobases produced are 6mA, 5mC, or 4mC. For example, EcoRI and BamHI

methyltransferases produce 5′-GA6mATTC-3′ and 5′-GGAT4mCC-3′, respectively.

Type III RM systems operate with multi-subunit machinery comprising two R subunits and two M subunits (R2M2) [33]. Subunit M contains recognition domain for binding to specific sites and also a methyltransferase domain. It can thereby bind at recognition sites independ‐ ently, and methylate DNA using *S*-adenosyl-L-methionine as a substrate. The nucleobase produced is 6mA in all cases reported so far. Unlike type I and II RM systems, full modifica‐ tion is actually hemimethylation (methylation on one strand). The recognition sequences are

Subunit R has an ATP dependent DNA helicase and endonuclease domains that are respon‐ sible for DNA cleavage. This subunit is unable to bind to DNA and therefore requires subu‐ nit M to cleave DNA. Two unmethylated sites that are inversely oriented (head-to-head orientation) serve as the target for DNA cleavage. Cleavage occurs at 25–27 bp downstream of one of the recognition sites, which is chosen randomly from the two sites. Even DNA with 3.5 kb between the two sites is cleaved. The cleavage requires ATP similar to type I RM systems. However, the amount of ATP consumed is only ~1% of that required for cleavage by type I RM systems. This fact makes it difficult to transpose the translocation model that is proposed in type I RM systems to type III RM systems. Thus, some alternative models have

Several enzymes specifically restrict modified DNA [10, 19-21, 26, 34-36]. These systems of‐ fer very efficient to defense from bacteriophages with highly-modified DNA. Among these, the enzymes for which cleavage sites are very specific and precise are classified into the M

asymmetric, such as 5′-CAGCAG-3′ for EcoP15I (methylated adenine is underlined).

produce 6mA.

**2.3. Type II RM systems**

**2.4. Type III RM systems**

now been proposed [33].

**2.5. Type IV restriction systems**

**Figure 2.** Chemical structures of modified nucleobases (modification moieties are indicated in red).

In addition to the methyl-based modifications described above, sulfur modification (phos‐ phorothioation) of DNA backbones has been observed [22, 23]. Notably, it is suggested that this modification is involved in DNA restriction in *Salmonella enterica* [24]. The gene cluster for this RM system consists of eight genes, of which four are involved in phosphorothioa‐ tion, while seven genes are essential for restricting unphosphorothioated DNA. Similar gene clusters, along with DNA containing phosphorothiol bonds, are found in many bacteria, im‐ plying that phosphorothioation is a widespread DNA modification [24, 25]. A type IV sys‐ tem that restricts phosphorothioated DNA has been also reported [26].

#### **2.2. Type I RM systems**

The *E. coli* strain K-12 harbors one type I RM system (EcoKI) encoded by the genes *hsdR* (R subunit), *hsdM* (M subunit), and *hsdS* (S subunit). This system constructs a multi-subunit complex that comprises two R subunits, two M subunits, and one S subunit (R2M2S), and scans double-stranded DNA after replication [27, 28]. When the complex recognizes DNA that is unmethylated at recognition sites, it acts as an ATP dependent endonuclease to digest DNA. The sequences of recognition sites are asymmetric but not palindromic. Examples in‐ clude 5′-AACN6GTGC-3′ and 5′-GCACN6GTT-3′ for EcoKI, 5′-TGAN8TGCT-3′ and 5′-AG‐ CAN8TCA-3′ for EcoBI, and 5′-TTAN7GTCY-3′ and 5′-RGACN7TAA-3′ for EcoDI (methylated adenine is underlined; R: A/G, Y: C/T, N: A/C/G/T) [29, 30].

The cleavage positions are distal from recognition sites and are variable. It is believed that the complex of type I RM system, while binding to recognition sites, translocates (or pulls) the DNA along in an ATP dependent fashion, and cleaves DNA when the translocation is impended by collision and/or by stalling with another translocating complex [27, 31, 32]. Electron microscopy analysis has been used to detect ATP-dependent formation of loop DNA during DNA cleavage by a type I RM complex [32]. The R subunit is responsible for ATP hydrolysis, translocation, and endonuclease activity, but not DNA binding. The bind‐ ing depends on subunits M and S, which are therefore essential for both endonuclease and methyltransferase activities.

The complex of type I RM system acts similar to methyltransferase when it recognizes DNA that is hemimethylated (methylated on one strand) at recognition sites [27]. The methylation is performed by M and S subunits using *S*-adenosyl-L-methionine as the methyl donor [8]. The M subunit has the binding site for *S*-adenosyl-L-methionine, while the S subunit is es‐ sential for determining recognition sites. The R subunit is unnecessary for methyltransferase activity. In all cases reported so far, methylation by type I RM systems occurs in adenine to produce 6mA.

#### **2.3. Type II RM systems**

**Figure 2.** Chemical structures of modified nucleobases (modification moieties are indicated in red).

222 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

tem that restricts phosphorothioated DNA has been also reported [26].

(methylated adenine is underlined; R: A/G, Y: C/T, N: A/C/G/T) [29, 30].

**2.2. Type I RM systems**

methyltransferase activities.

In addition to the methyl-based modifications described above, sulfur modification (phos‐ phorothioation) of DNA backbones has been observed [22, 23]. Notably, it is suggested that this modification is involved in DNA restriction in *Salmonella enterica* [24]. The gene cluster for this RM system consists of eight genes, of which four are involved in phosphorothioa‐ tion, while seven genes are essential for restricting unphosphorothioated DNA. Similar gene clusters, along with DNA containing phosphorothiol bonds, are found in many bacteria, im‐ plying that phosphorothioation is a widespread DNA modification [24, 25]. A type IV sys‐

The *E. coli* strain K-12 harbors one type I RM system (EcoKI) encoded by the genes *hsdR* (R subunit), *hsdM* (M subunit), and *hsdS* (S subunit). This system constructs a multi-subunit complex that comprises two R subunits, two M subunits, and one S subunit (R2M2S), and scans double-stranded DNA after replication [27, 28]. When the complex recognizes DNA that is unmethylated at recognition sites, it acts as an ATP dependent endonuclease to digest DNA. The sequences of recognition sites are asymmetric but not palindromic. Examples in‐ clude 5′-AACN6GTGC-3′ and 5′-GCACN6GTT-3′ for EcoKI, 5′-TGAN8TGCT-3′ and 5′-AG‐ CAN8TCA-3′ for EcoBI, and 5′-TTAN7GTCY-3′ and 5′-RGACN7TAA-3′ for EcoDI

The cleavage positions are distal from recognition sites and are variable. It is believed that the complex of type I RM system, while binding to recognition sites, translocates (or pulls) the DNA along in an ATP dependent fashion, and cleaves DNA when the translocation is impended by collision and/or by stalling with another translocating complex [27, 31, 32]. Electron microscopy analysis has been used to detect ATP-dependent formation of loop DNA during DNA cleavage by a type I RM complex [32]. The R subunit is responsible for ATP hydrolysis, translocation, and endonuclease activity, but not DNA binding. The bind‐ ing depends on subunits M and S, which are therefore essential for both endonuclease and

The complex of type I RM system acts similar to methyltransferase when it recognizes DNA that is hemimethylated (methylated on one strand) at recognition sites [27]. The methylation is performed by M and S subunits using *S*-adenosyl-L-methionine as the methyl donor [8]. The M subunit has the binding site for *S*-adenosyl-L-methionine, while the S subunit is es‐ sential for determining recognition sites. The R subunit is unnecessary for methyltransferase

Type II RM systems are extremely diverse and are currently classified into 11 subfamilies [8]. Generally, these comprise two enzymes, RE and MT. Cleavages of exogenous DNA at unmethylated recognition sites is carried out by RE, which spares endogenous DNA that has been methylated by the cognate MT. Most REs require Mg2+ ions as a cofactor for cleav‐ age. Although RE may form monomers, dimers, or tetramers, it functions without forming complexes with the cognate MT. The recognition of cleavage sites is highly precise and rec‐ ognition sequences are often palindromic. Such sites include 5′-GAATTC-3′ for EcoRI and 5′-GGATCC-3′ for BamHI, which are cleaved symmetrically within the sites. Because of these useful properties, more than 3,500 REs have been characterized, and many are widely utilized in recombinant DNA technology [8]. The enzyme MT catalyzes methylation at rec‐ ognition sites using *S*-adenosyl-L-methionine as the substrate and generally acts as a mono‐ mer. The nucleobases produced are 6mA, 5mC, or 4mC. For example, EcoRI and BamHI methyltransferases produce 5′-GA6mATTC-3′ and 5′-GGAT4mCC-3′, respectively.

#### **2.4. Type III RM systems**

Type III RM systems operate with multi-subunit machinery comprising two R subunits and two M subunits (R2M2) [33]. Subunit M contains recognition domain for binding to specific sites and also a methyltransferase domain. It can thereby bind at recognition sites independ‐ ently, and methylate DNA using *S*-adenosyl-L-methionine as a substrate. The nucleobase produced is 6mA in all cases reported so far. Unlike type I and II RM systems, full modifica‐ tion is actually hemimethylation (methylation on one strand). The recognition sequences are asymmetric, such as 5′-CAGCAG-3′ for EcoP15I (methylated adenine is underlined).

Subunit R has an ATP dependent DNA helicase and endonuclease domains that are respon‐ sible for DNA cleavage. This subunit is unable to bind to DNA and therefore requires subu‐ nit M to cleave DNA. Two unmethylated sites that are inversely oriented (head-to-head orientation) serve as the target for DNA cleavage. Cleavage occurs at 25–27 bp downstream of one of the recognition sites, which is chosen randomly from the two sites. Even DNA with 3.5 kb between the two sites is cleaved. The cleavage requires ATP similar to type I RM systems. However, the amount of ATP consumed is only ~1% of that required for cleavage by type I RM systems. This fact makes it difficult to transpose the translocation model that is proposed in type I RM systems to type III RM systems. Thus, some alternative models have now been proposed [33].

#### **2.5. Type IV restriction systems**

Several enzymes specifically restrict modified DNA [10, 19-21, 26, 34-36]. These systems of‐ fer very efficient to defense from bacteriophages with highly-modified DNA. Among these, the enzymes for which cleavage sites are very specific and precise are classified into the M subfamily of type II RM systems [21]. Examples include DpnI (recognition sequence: 5′- G6mATC-3′), GlaI (5′-G5mCG5mC-3′), BisI (5′-G5mCNGC-3′; N: A/G/C/T), and MspJI (5′-5mCNNR-3′; R: G/A). The enzymes with non-specific and variable cleavage sites are clas‐ sified as type IV restriction systems. The *E. coli* strain K-12 harbors three type IV systems encoded by *mcrA*, *mcrB*-*mcrC*, and *mrr*. The enzyme McrA recognizes 5′-Y5mCGR-3′ site (Y: C/T; R: G/A) [37], whereas McrBC recognizes pairs of 5′-R<sup>m</sup>C-3′ (mC: 5mC or 4mC) separated by 40–3000 bp, and cleaves DNA ~30 bp distal from one of the sites [36]. The Mrr system recognizes DNA containing 6mA, 5mC, or 4mC, but its recognition sites have not been well de‐ fined [36]. The enzyme SauUSI of *Staphylococcus aureus* recognizes 5′-S5mCNGS-3′ and 5′- S5hmCNGS-3′ (S: C/G; N: A/G/C/T) and cleaves at position 2–18 bp downstream of the recognition site [21]. The enzyme GmrSD restriction system of *E. coli* CT596 cuts DNA con‐ taining ghmC [20]. Thus, all species of modified nucleobases are potentially restricted by type IV restriction systems.

**3. Host-mimicking strategies**

(

Circumvention of RM systems is critical for establishing transformation methods for target bacteria (Table 2). This is true not only for bacteria but also archaea [38]. This section de‐ scribes some strategies for circumventing RM systems and focuses on host-mimicking. Be‐ cause all types of RM systems digest exogenous DNA after distinguishing it from endogenous DNA on the basis of host-specific methylation patterns, DNA modification that mimic these patterns evade digestion. A general flowchart for producing host-mimicking

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225

**Figure 3.** General flowchart for producing host-mimicking DNA to target bacteria.The chromatogram is the data from HPLC analysis of deoxynucleosides prepared from *G. kaustophilus* chromosomes [41]. It includes 2′-deoxyadenosine (dA), 2′-deoxycytidine (dC), 2′-deoxyguanosine (dG), 2′-deoxythymidine (dT), and *N* <sup>6</sup>-methyl-2′-deoxyadenosine

As mentioned earlier, type I–III RM systems involve DNA methylation in their functions. Therefore, the presence of methylated DNA in bacterial chromosomes indicates a type I–III RM system in the bacterium. Hence analysis of methylated DNA is an effective survey method to identify functional RM systems in target bacterium. Methylated DNA in chromo‐ somes can be analyzed using high-performance liquid chromatography (HPLC) [15, 18, 39-45]. This method determines the presence of deoxynucleosides and methylated deoxynu‐ cleosides. Deoxynucleosides are prepared by hydrolyzing chromosomal DNA with nuclease P1 and alkaline phosphatase [39-41], are separated using reverse mode C18-based silica col‐ umns, and are detected by ultraviolet absorption at 260 and/or 280 nm. Authentic 6mdA, 5mdC, and 4mdC are commercially available or can be prepared from methylated 2′-deoxynu‐

6mdA) but not 5-methyl-2′-deoxycytidine (5mdC) or *N* <sup>4</sup>-methyl-2′-deoxycytidine (4mdC).

**3.1. A brief survey of functional RM systems**

DNA is shown in Figure 3. The details are described in the following sections.


**Table 2.** Microbial transformation using host-mimicking DNA.DS: *in vivo* methylation in a strain that is related to the target bacterium and is deficient in restriction and proficient in methylation; EX: *in vitro* methylation using a crude extract of the target bacterium; GM: *in vivo* methylation using methyltransferase genes in the target bacterial genome; HG: *in vivo* methylation using heterologous genes; IG: *in vivo* methylation using *E. coli* intrinsic genes; MF: methyl-free DNA; MT: *in vitro* methylation using commercially available methyltransferases; and SD: transformation using DNA with specifically abolished recognition sites.

#### **3. Host-mimicking strategies**

subfamily of type II RM systems [21]. Examples include DpnI (recognition sequence: 5′- G6mATC-3′), GlaI (5′-G5mCG5mC-3′), BisI (5′-G5mCNGC-3′; N: A/G/C/T), and MspJI (5′-5mCNNR-3′; R: G/A). The enzymes with non-specific and variable cleavage sites are clas‐ sified as type IV restriction systems. The *E. coli* strain K-12 harbors three type IV systems encoded by *mcrA*, *mcrB*-*mcrC*, and *mrr*. The enzyme McrA recognizes 5′-Y5mCGR-3′ site (Y: C/T; R: G/A) [37], whereas McrBC recognizes pairs of 5′-R<sup>m</sup>C-3′ (mC: 5mC or 4mC) separated by 40–3000 bp, and cleaves DNA ~30 bp distal from one of the sites [36]. The Mrr system recognizes DNA containing 6mA, 5mC, or 4mC, but its recognition sites have not been well de‐ fined [36]. The enzyme SauUSI of *Staphylococcus aureus* recognizes 5′-S5mCNGS-3′ and 5′- S5hmCNGS-3′ (S: C/G; N: A/G/C/T) and cleaves at position 2–18 bp downstream of the recognition site [21]. The enzyme GmrSD restriction system of *E. coli* CT596 cuts DNA con‐ taining ghmC [20]. Thus, all species of modified nucleobases are potentially restricted by type

224 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Microbe Host-mimicking DNA Reference**

*Bacillus anthracis In vivo* (MF) Electroporation [50] *Bacillus cereus In vitro* (EX) Electroporation [51] *Bacillus weihenstephanensis In vitro* (EX) Electroporation [51] *Bifidobacterium adolescentis In vivo* (GM) Electroporation [52] *Bifidobacterium longum In vitro* (MT/SD) Electroporation [53] *Borrelia burgdorferi In vitro* (MT) Electroporation [54] *Clostridium acetobutylicum In vivo* (HG) Electroporation [55] *Clostridium difficile In vivo* (HG/SD) Conjugation [56] *Clostridium thermocellum In vivo* (IG) Electroporation [57] *Geobacillus kaustophilus In vivo* (GM/IG) Conjugation [41] *Helicobacter pylori In vitro* (EX) Competency [58] *Salmonella typhimurium In vivo* (DS) Competency [59] *Staphylococcus aureus In vivo* (DS) Electroporation [60] *Streptomyces avermitilis In vivo* (MF) Protoplast [61] *Streptomyces bambergiensis In vivo* (MF) Conjugation [62] *Streptomyces coelicolor In vivo* (MF) Protoplast [35]

*Streptomyces griseus* NRRL B-2682 *In vitro* (MT) Protoplast [64] *Streptomyces natalensis In vivo* (MF) Conjugation [65] *Sulfolobus acidocaldarius In vivo* (HG) Electroporation [38] *Thermoanaerobacter* sp. X514 *In vivo* (IG) Sonoporation [66]

**Table 2.** Microbial transformation using host-mimicking DNA.DS: *in vivo* methylation in a strain that is related to the target bacterium and is deficient in restriction and proficient in methylation; EX: *in vitro* methylation using a crude extract of the target bacterium; GM: *in vivo* methylation using methyltransferase genes in the target bacterial genome; HG: *in vivo* methylation using heterologous genes; IG: *in vivo* methylation using *E. coli* intrinsic genes; MF: methyl-free DNA; MT: *in vitro* methylation using commercially available methyltransferases; and SD: transformation

**Production Introduction**

*In vivo* (GM/HG) Protoplast [40] *In vivo* (DS) Protoplast [63]

IV restriction systems.

*Streptomyces griseus* IFO 13350

using DNA with specifically abolished recognition sites.

Circumvention of RM systems is critical for establishing transformation methods for target bacteria (Table 2). This is true not only for bacteria but also archaea [38]. This section de‐ scribes some strategies for circumventing RM systems and focuses on host-mimicking. Be‐ cause all types of RM systems digest exogenous DNA after distinguishing it from endogenous DNA on the basis of host-specific methylation patterns, DNA modification that mimic these patterns evade digestion. A general flowchart for producing host-mimicking DNA is shown in Figure 3. The details are described in the following sections.

**Figure 3.** General flowchart for producing host-mimicking DNA to target bacteria.The chromatogram is the data from HPLC analysis of deoxynucleosides prepared from *G. kaustophilus* chromosomes [41]. It includes 2′-deoxyadenosine (dA), 2′-deoxycytidine (dC), 2′-deoxyguanosine (dG), 2′-deoxythymidine (dT), and *N* <sup>6</sup>-methyl-2′-deoxyadenosine ( 6mdA) but not 5-methyl-2′-deoxycytidine (5mdC) or *N* <sup>4</sup>-methyl-2′-deoxycytidine (4mdC).

#### **3.1. A brief survey of functional RM systems**

As mentioned earlier, type I–III RM systems involve DNA methylation in their functions. Therefore, the presence of methylated DNA in bacterial chromosomes indicates a type I–III RM system in the bacterium. Hence analysis of methylated DNA is an effective survey method to identify functional RM systems in target bacterium. Methylated DNA in chromo‐ somes can be analyzed using high-performance liquid chromatography (HPLC) [15, 18, 39-45]. This method determines the presence of deoxynucleosides and methylated deoxynu‐ cleosides. Deoxynucleosides are prepared by hydrolyzing chromosomal DNA with nuclease P1 and alkaline phosphatase [39-41], are separated using reverse mode C18-based silica col‐ umns, and are detected by ultraviolet absorption at 260 and/or 280 nm. Authentic 6mdA, 5mdC, and 4mdC are commercially available or can be prepared from methylated 2′-deoxynu‐ cleoside-5′-triphosphate by dephosphorylation using *E. coli* alkaline phosphatase [40]. Note that this analysis requires large amounts of DNA (>10 µg). Contamination with RNA, pro‐ teins, and chemicals that absorb ultraviolet radiation may also complicate accurate analysis. In some reports [18, 42, 43], deoxynucleosides are more accurately identified by combining HPLC and mass spectrometry analysis. In addition, immunochemical methods are available for analyzing methylated DNA [46-49]. Several anti-5mC antibodies have been developed and are commercially available. Although commercially available antibodies against other methylated DNA are limited, Kong *et al*. [49] have reported successful production of rabbit polyclonal antibodies against 6mA and 4mC.

cific sites, or differential DNA methylation in chromosomes, there are a few methods that

Host-Mimicking Strategies in DNA Methylation for Improved Bacterial Transformation

http://dx.doi.org/10.5772/51691

227

In *S. griseus*, bisulfite-based analysis of a plasmid library isolated from this bacterium (Fig‐ ure 4A) was used to determine the two consensus sites 5′-GAG5mCTC-3′ and 5′-GC5mCGGC-3′ [40]. Note that this method is not employed for methylation analysis in bacteria that cannot be transformed with exogenous plasmids, and in bacteria that have methylated nucleobases other than 5mC. For determining 6mA consensus sites in *G. kaustophilus*, chromosome diges‐ tion using methyl-sensitive restriction enzymes (Figure 4B) was used to reveal 5′-GG6mATC-3′ and 5′-G6mATCC-3′ site [41]. Methods using methyl-sensitive restriction enzymes help iden‐ tify several methylation species, including 6mA, 5mC, and 4mC. However, methylation sites that can be identified by these methods are limited due to the lack of commercially available restriction enzymes. Recently, direct detection using real-time DNA sequencing has been reported [74, 75]. This method potentially enables the exhaustive determination of all meth‐ ylation sites in chromosomal DNA. Although this method requires special equipments, which has limited availability, it may become one of the most promising methods for methylation analysis in the future. Also, the author has now developed a versatile immunological meth‐

If the target bacterium contains no or negligible methylated DNA, methyl-free DNA should be used as host-mimicking DNA because the bacterium may harbor type IV restriction sys‐ tems, as exemplified by transformation of *S. avermitilis* [61], *S. coelicolor* [35], and *S. natalensis* [65]. Methyl-free DNA can be readily produced using *E. coli* strains deficient in DNA meth‐

also used for efficient transformation of *S. bambergiensis* [62] and *B. anthracis* [50], respective‐ ly. Because Hsd mediated methylation is of low frequency in *E. coli*, deficiency of this meth‐

If the target bacterium contains considerable methylated DNA, host-mimicking DNA that reconstitutes the methylation pattern of the target bacterium should be used for transforma‐ tion. There are two main approaches to produce heterologous methylation of DNA. One is *in vitro* methylation using methyltransferases [53, 54, 64], such as Dam (catalyzing 5′- G6mATC-3′ methylation), M.TaqI (5′-TCG6mA-3′), M.AluI (5′-AG5mCT-3′), M.BamHI (5′- GGATC4mC-3′), M.SssI (5′-5mCG-3′), M.EcoRI (5′-GA6mATTC-3′), M.CviPI (5′-GC5m-3′), M.HaeIII (5′-GGC5mC-3′), M.HhaI (5′-G5mCGC-3′), M.HpaII (5′-C5mCGG-3′), and M.MspI (5′-5mCCGG-3′). This approach is very simple and effective but has low cost-performance and low versatility due to the limited number of commercially available methyltransferases. In transformations of *B. cereus* [51], *B. weihenstephanensis* [51], and *H. pylori* [58], crude ex‐ tracts prepared from the respective bacterium were used for DNA methylation. This method has high cost-performance and high versatility but may not be efficient because of low methyltransferase activity in crude extracts and DNA degradation by nucleases. Moreover,

(Table 4). Methyl-restrained DNA from *E. coli* GM2929 and SCS110 (*dam* <sup>−</sup> *dcm* <sup>−</sup> *hsd* <sup>+</sup>

), such as *E. coli* IR27, ET12567, IBEC58, and HST04

) were

exhaustively determine methylated consensus sites in chromosomes as follows.

od for determining consensus sequences with methylated nucleobases.

yltion may not be essential for circumventing type IV systems.

type I and III methylation pattern cannot be achieved by this method.

**3.3. Production of host-mimicking DNA**

yltransferase genes (*dam* <sup>−</sup> *dcm* <sup>−</sup> *hsd* <sup>−</sup>


**Table 3.** Relative coefficients for determining deoxynucleoside molar ratios by HPLC analysis; [Relative amount of deoxynucleoside] = [Coefficient] × [Peak area on HPLC chromatogram].

Analysis of DNA using HPLC allows determination of deoxynucleoside composition and methylation frequency in chromosomes. The coefficients in Table 3 may be used for estima‐ tion. For example, HPLC analysis of *S. griseus* chromosomes revealed 5mdC but not 6mdA or 4mdC [40]. The composition ratio of 5mdC to dC was 0.7 mol%, suggesting that *S. griseus* pos‐ sesses approximately one 5mC per 0.5 kb of chromosomal DNA with 67% GC content. The chromosome of *G. kaustophilus* contains 6mdA and the composition ratio to dA is 2.0 mol% [41]. This suggests that *G. kaustophilus* possesses approximately one 6mA per 0.1 kb of chro‐ mosomal DNA with 52% GC content. Although DNA methylation unrelated to RM systems have been observed, such as *E. coli* Dam and Dcm methylation [67, 68], high frequency methylations imply that the bacteria may harbor a considerable type I–III RM system. Mean‐ while, the chromosomes of *S. avermitilis*, *S. coelicolor*, and *S. lividans* contain no methylated DNA (my unpublished data). This observation suggests that these bacteria harbor no type I– III RM systems. However, the possibility remains that a functional type IV system exists in these species. Potent methyl-specific restrictions have been observed in *S. avermitilis* [61], *S. coelicolor* [35], *S. bambergiensis* [62], and *S. natalensis* [65].

#### **3.2. Methylation site analysis**

When significant DNA methylation is observed in the target bacterium, preliminary deter‐ mination of DNA methylation sites is generally required to produce host-mimicking DNA. Recent epigenetic studies have developed many methods to analyze DNA methylation [29, 30, 69-76]. Although most of these studies aimed to analyze 5-methylation of cytosine at spe‐ cific sites, or differential DNA methylation in chromosomes, there are a few methods that exhaustively determine methylated consensus sites in chromosomes as follows.

In *S. griseus*, bisulfite-based analysis of a plasmid library isolated from this bacterium (Fig‐ ure 4A) was used to determine the two consensus sites 5′-GAG5mCTC-3′ and 5′-GC5mCGGC-3′ [40]. Note that this method is not employed for methylation analysis in bacteria that cannot be transformed with exogenous plasmids, and in bacteria that have methylated nucleobases other than 5mC. For determining 6mA consensus sites in *G. kaustophilus*, chromosome diges‐ tion using methyl-sensitive restriction enzymes (Figure 4B) was used to reveal 5′-GG6mATC-3′ and 5′-G6mATCC-3′ site [41]. Methods using methyl-sensitive restriction enzymes help iden‐ tify several methylation species, including 6mA, 5mC, and 4mC. However, methylation sites that can be identified by these methods are limited due to the lack of commercially available restriction enzymes. Recently, direct detection using real-time DNA sequencing has been reported [74, 75]. This method potentially enables the exhaustive determination of all meth‐ ylation sites in chromosomal DNA. Although this method requires special equipments, which has limited availability, it may become one of the most promising methods for methylation analysis in the future. Also, the author has now developed a versatile immunological meth‐ od for determining consensus sequences with methylated nucleobases.

#### **3.3. Production of host-mimicking DNA**

cleoside-5′-triphosphate by dephosphorylation using *E. coli* alkaline phosphatase [40]. Note that this analysis requires large amounts of DNA (>10 µg). Contamination with RNA, pro‐ teins, and chemicals that absorb ultraviolet radiation may also complicate accurate analysis. In some reports [18, 42, 43], deoxynucleosides are more accurately identified by combining HPLC and mass spectrometry analysis. In addition, immunochemical methods are available for analyzing methylated DNA [46-49]. Several anti-5mC antibodies have been developed and are commercially available. Although commercially available antibodies against other methylated DNA are limited, Kong *et al*. [49] have reported successful production of rabbit

> dC 2.34 1.74 5mdC 3.51 1.77 4mdC 1.77 1.57 dA 1.08 6.42 6mdA 1.15 1.93

**Table 3.** Relative coefficients for determining deoxynucleoside molar ratios by HPLC analysis; [Relative amount of

Analysis of DNA using HPLC allows determination of deoxynucleoside composition and methylation frequency in chromosomes. The coefficients in Table 3 may be used for estima‐ tion. For example, HPLC analysis of *S. griseus* chromosomes revealed 5mdC but not 6mdA or 4mdC [40]. The composition ratio of 5mdC to dC was 0.7 mol%, suggesting that *S. griseus* pos‐ sesses approximately one 5mC per 0.5 kb of chromosomal DNA with 67% GC content. The chromosome of *G. kaustophilus* contains 6mdA and the composition ratio to dA is 2.0 mol% [41]. This suggests that *G. kaustophilus* possesses approximately one 6mA per 0.1 kb of chro‐ mosomal DNA with 52% GC content. Although DNA methylation unrelated to RM systems have been observed, such as *E. coli* Dam and Dcm methylation [67, 68], high frequency methylations imply that the bacteria may harbor a considerable type I–III RM system. Mean‐ while, the chromosomes of *S. avermitilis*, *S. coelicolor*, and *S. lividans* contain no methylated DNA (my unpublished data). This observation suggests that these bacteria harbor no type I– III RM systems. However, the possibility remains that a functional type IV system exists in these species. Potent methyl-specific restrictions have been observed in *S. avermitilis* [61], *S.*

When significant DNA methylation is observed in the target bacterium, preliminary deter‐ mination of DNA methylation sites is generally required to produce host-mimicking DNA. Recent epigenetic studies have developed many methods to analyze DNA methylation [29, 30, 69-76]. Although most of these studies aimed to analyze 5-methylation of cytosine at spe‐

**Relative coefficient Detection at 260 nm Detection at 280 nm**

polyclonal antibodies against 6mA and 4mC.

226 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

deoxynucleoside] = [Coefficient] × [Peak area on HPLC chromatogram].

*coelicolor* [35], *S. bambergiensis* [62], and *S. natalensis* [65].

**3.2. Methylation site analysis**

**Deoxynucleoside**

If the target bacterium contains no or negligible methylated DNA, methyl-free DNA should be used as host-mimicking DNA because the bacterium may harbor type IV restriction sys‐ tems, as exemplified by transformation of *S. avermitilis* [61], *S. coelicolor* [35], and *S. natalensis* [65]. Methyl-free DNA can be readily produced using *E. coli* strains deficient in DNA meth‐ yltransferase genes (*dam* <sup>−</sup> *dcm* <sup>−</sup> *hsd* <sup>−</sup> ), such as *E. coli* IR27, ET12567, IBEC58, and HST04 (Table 4). Methyl-restrained DNA from *E. coli* GM2929 and SCS110 (*dam* <sup>−</sup> *dcm* <sup>−</sup> *hsd* <sup>+</sup> ) were also used for efficient transformation of *S. bambergiensis* [62] and *B. anthracis* [50], respective‐ ly. Because Hsd mediated methylation is of low frequency in *E. coli*, deficiency of this meth‐ yltion may not be essential for circumventing type IV systems.

If the target bacterium contains considerable methylated DNA, host-mimicking DNA that reconstitutes the methylation pattern of the target bacterium should be used for transforma‐ tion. There are two main approaches to produce heterologous methylation of DNA. One is *in vitro* methylation using methyltransferases [53, 54, 64], such as Dam (catalyzing 5′- G6mATC-3′ methylation), M.TaqI (5′-TCG6mA-3′), M.AluI (5′-AG5mCT-3′), M.BamHI (5′- GGATC4mC-3′), M.SssI (5′-5mCG-3′), M.EcoRI (5′-GA6mATTC-3′), M.CviPI (5′-GC5m-3′), M.HaeIII (5′-GGC5mC-3′), M.HhaI (5′-G5mCGC-3′), M.HpaII (5′-C5mCGG-3′), and M.MspI (5′-5mCCGG-3′). This approach is very simple and effective but has low cost-performance and low versatility due to the limited number of commercially available methyltransferases. In transformations of *B. cereus* [51], *B. weihenstephanensis* [51], and *H. pylori* [58], crude ex‐ tracts prepared from the respective bacterium were used for DNA methylation. This method has high cost-performance and high versatility but may not be efficient because of low methyltransferase activity in crude extracts and DNA degradation by nucleases. Moreover, type I and III methylation pattern cannot be achieved by this method.

were initially introduced and propagated in the restriction-deficient strains LB5000 [59] and RN4220 [60], respectively, and were then used for transformation of other strains. In *S. griseus* transformation, mutant HH1 that reduces restriction activity compared to the wild-type has been used [63]. This approach enables production of perfect host-mimick‐ ing DNA, although it is not easy to find a strain that is both deficient in restriction and

Host-Mimicking Strategies in DNA Methylation for Improved Bacterial Transformation

**Strain Relevant genotype Reference**

(StrR) [40]

LT

http://dx.doi.org/10.5772/51691

229

IR21 *e*14<sup>−</sup>(*mcrA*−) Δ*dam*::*metB* Δ(*mrr*-*hsdRMS*-*mcrBC*)114::*IS10 rpsL*104 (StrR) [41] IR24 *e*14<sup>−</sup>(*mcrA*−) Δ*dcm*::*lacZ* Δ(*mrr*-*hsdRMS*-*mcrBC*)114::*IS10 rpsL*104 (StrR) [41]

ET12567 *dam*-13::*Tn9* (CmR) *dcm*-6 *hsdRMzjj*-202::*Tn10* (TetR) *rpsL*136 (StrR) [77] IBEC58 Δ*dam* Δ*dcm* Δ*hsdRMS* [35] HST04 Δ(*mrr*-*hsdRMS*-*mcrBC*) Δ*mcrA dam dcm rpsL* (StrR) TB JM110 *dam dcm rpsL* (StrR) AT SCS110 *dam dcm rpsL* (StrR) *endA* AT

GM48 *dam*-3 *dcm*-6 CGSC GM272 *dam*-3 *dcm*-6 *hsdS*21 CGSC GM2929 *dam*-13::*Tn9* (CmR) *dcm*-6 *hsdR*2 *mcrA mcrB rpsL*136 (StrR) CGSC

**Table 4.** *E. coli* strains deficient in genes involved in DNA methylation and/or DNA restriction.CmR: chloramphenicol resistance; TetR: tetracycline resistance; StrR: streptomycin resistance; TB: Takara Bio Inc. (www.takara-bio.com); AT: Agilent Technologies Inc. (home.agilent.com); LT: Life Technologies Corporation (www.lifetechnologies.com); and

In addition to host-mimicking strategies, there are some simple approaches for circumvent‐ ing RM systems. One is to abolish sites recognized by RM systems. In *Clostridium difficile* transformation, five *Cdi*I sites in plasmids were abolished and used to demonstrate im‐ proved efficiency [56]. Similarly, a plasmid with three abolished *Sac*II sites was used for effi‐ cient transformation of *Bifidobacterium longum* [53]. When methyltransferase enzymes or genes are unavailable, this approach can be an effective alternative. One other approach is to reduce the restriction activity in the target bacterium temporarily by heat treatment. In *B. amyloliquefaciens* transformation, heat treatment at 46°C for 6 min increased transformation efficiency [78]. More forcible heat treatment (higher temperature and longer time) inacti‐ vates RM systems more efficiently but concurrently reduces viability of cells. Although this approach is very simple and is available for most bacteria, the heat conditions required for

IR27 *<sup>e</sup>*14<sup>−</sup>(*mcrA*−) Δ*dam*::*metB* Δ*dcm*::*lacZ* Δ(*mrr*-*hsdRMS*-*mcrBC*)114::*IS10 rpsL*<sup>104</sup>

*dam dcm* Δ(*mrr*-*hsdRMS*-*mcrBC*)102::*Tn10* (TetR)

*rps*L (StrR) *endA*

CGSC: The Coli Genetic Stock Center (cgsc.biology.yale.edu).

**3.4. Alternative methods for circumventing RM systems**

proficient in methylation.

INV110

**Figure 4.** Methylation site analysis in target bacterium. (A) Bisulfite-based analysis to determine 5mC consensus sites. Bisulfite treatment converts methyl-free cytosine to uracil without affecting 5mC. Therefore, 5mC positions can be deter‐ mined by comparing bisulfite-treated and -untreated DNA sequences. (B) Chromosomal digestion by methyl-sensitive restriction enzymes is used to analyze 5′-G6mATC-3′ methylation. The restriction enzyme DpnI cuts 5′-G6mATC-3′ but not 5′-GATC-3′, DpnII cuts 5′-GATC-3′ but not 5′-G6mATC-3′, and Sau3AI cuts 5′-GATC-3′ and 5′-G6mATC-3′.

Another approach is *in vivo* methylation by expressing methyltransferase genes in *E. coli* cells [38, 40, 41, 52, 55, 56]. Type II and III methylation can be reconstituted by expressing MT and M subunit genes, respectively, and type I methylation can be reconstituted by si‐ multaneous expression of M and S subunit genes. Numerous gene sequences of methyl‐ transferases are accumulated in the REBASE database along with their methylation sites [11]. Either plasmids or chromosomal integration can be used as expression vectors for methyltransferase genes. When the genome sequence of the target bacterium has been deter‐ mined, methyltransferase genes in the genome may be used for *in vivo* methylation [40, 41, 52]. Methylation site analysis is not essential for this approach; however, functional expres‐ sion of methyltransferase genes requires confirmation by HPLC analysis of recombinant *E. coli* chromosomes. A methyltransferase gene of *S. griseus*, responsible for 5′-GAG5mCTC-3′ methylation, was found to be nonfunctional in *E. coli*. Hence, an alternative methyltransfer‐ ase gene from *S. achromogenes* (*M.SacI*) was used for DNA methylation [40]. The *E. coli* host used to produce host-mimicking DNA must be a methylation-deficient strain (Table 4) be‐ cause the target bacterium may have type IV systems in addition to type I–III RM systems, as exemplified by *G. kaustophilus* transformation [41]. In addition, it is desirable that the *E. coli* host is deficient in type IV system genes (*mcrA*, *mcrBC*, and *mrr*) because these may re‐ strict heterologous methylation in *E. coli* cells. In this regard, *E. coli* strains IR27 and HST04 are appropriate for producing host-mimicking DNA through *in vivo* methylation. Although *in vivo* methylation may be more complicated than *in vitro* methylation, this approach often has excellent cost-performance, versatility, and efficiency.

A derivative of the target bacterium with methylation activity and reduced restriction activity can also be used for the production of host-mimicking DNA. In transformations of *Salmonella typhimurium* and *Staphylococcus aureus*, plasmids isolated from *E. coli* strains were initially introduced and propagated in the restriction-deficient strains LB5000 [59] and RN4220 [60], respectively, and were then used for transformation of other strains. In *S. griseus* transformation, mutant HH1 that reduces restriction activity compared to the wild-type has been used [63]. This approach enables production of perfect host-mimick‐ ing DNA, although it is not easy to find a strain that is both deficient in restriction and proficient in methylation.


**Table 4.** *E. coli* strains deficient in genes involved in DNA methylation and/or DNA restriction.CmR: chloramphenicol resistance; TetR: tetracycline resistance; StrR: streptomycin resistance; TB: Takara Bio Inc. (www.takara-bio.com); AT: Agilent Technologies Inc. (home.agilent.com); LT: Life Technologies Corporation (www.lifetechnologies.com); and CGSC: The Coli Genetic Stock Center (cgsc.biology.yale.edu).

#### **3.4. Alternative methods for circumventing RM systems**

**Figure 4.** Methylation site analysis in target bacterium. (A) Bisulfite-based analysis to determine 5mC consensus sites. Bisulfite treatment converts methyl-free cytosine to uracil without affecting 5mC. Therefore, 5mC positions can be deter‐ mined by comparing bisulfite-treated and -untreated DNA sequences. (B) Chromosomal digestion by methyl-sensitive restriction enzymes is used to analyze 5′-G6mATC-3′ methylation. The restriction enzyme DpnI cuts 5′-G6mATC-3′ but

Another approach is *in vivo* methylation by expressing methyltransferase genes in *E. coli* cells [38, 40, 41, 52, 55, 56]. Type II and III methylation can be reconstituted by expressing MT and M subunit genes, respectively, and type I methylation can be reconstituted by si‐ multaneous expression of M and S subunit genes. Numerous gene sequences of methyl‐ transferases are accumulated in the REBASE database along with their methylation sites [11]. Either plasmids or chromosomal integration can be used as expression vectors for methyltransferase genes. When the genome sequence of the target bacterium has been deter‐ mined, methyltransferase genes in the genome may be used for *in vivo* methylation [40, 41, 52]. Methylation site analysis is not essential for this approach; however, functional expres‐ sion of methyltransferase genes requires confirmation by HPLC analysis of recombinant *E. coli* chromosomes. A methyltransferase gene of *S. griseus*, responsible for 5′-GAG5mCTC-3′ methylation, was found to be nonfunctional in *E. coli*. Hence, an alternative methyltransfer‐ ase gene from *S. achromogenes* (*M.SacI*) was used for DNA methylation [40]. The *E. coli* host used to produce host-mimicking DNA must be a methylation-deficient strain (Table 4) be‐ cause the target bacterium may have type IV systems in addition to type I–III RM systems, as exemplified by *G. kaustophilus* transformation [41]. In addition, it is desirable that the *E. coli* host is deficient in type IV system genes (*mcrA*, *mcrBC*, and *mrr*) because these may re‐ strict heterologous methylation in *E. coli* cells. In this regard, *E. coli* strains IR27 and HST04 are appropriate for producing host-mimicking DNA through *in vivo* methylation. Although *in vivo* methylation may be more complicated than *in vitro* methylation, this approach often

A derivative of the target bacterium with methylation activity and reduced restriction activity can also be used for the production of host-mimicking DNA. In transformations of *Salmonella typhimurium* and *Staphylococcus aureus*, plasmids isolated from *E. coli* strains

not 5′-GATC-3′, DpnII cuts 5′-GATC-3′ but not 5′-G6mATC-3′, and Sau3AI cuts 5′-GATC-3′ and 5′-G6mATC-3′.

228 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

has excellent cost-performance, versatility, and efficiency.

In addition to host-mimicking strategies, there are some simple approaches for circumvent‐ ing RM systems. One is to abolish sites recognized by RM systems. In *Clostridium difficile* transformation, five *Cdi*I sites in plasmids were abolished and used to demonstrate im‐ proved efficiency [56]. Similarly, a plasmid with three abolished *Sac*II sites was used for effi‐ cient transformation of *Bifidobacterium longum* [53]. When methyltransferase enzymes or genes are unavailable, this approach can be an effective alternative. One other approach is to reduce the restriction activity in the target bacterium temporarily by heat treatment. In *B. amyloliquefaciens* transformation, heat treatment at 46°C for 6 min increased transformation efficiency [78]. More forcible heat treatment (higher temperature and longer time) inacti‐ vates RM systems more efficiently but concurrently reduces viability of cells. Although this approach is very simple and is available for most bacteria, the heat conditions required for inactivating RM systems are not predictable and thereby may have to be determined by re‐ peated trials.

[3] McCarty, M., & Avery, O. T. (1946). Studies on the Chemical Nature of the Substance Inducing Transformation of Pneumococcal Types: II. Effect of Desoxyribonuclease on

Host-Mimicking Strategies in DNA Methylation for Improved Bacterial Transformation

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231

[4] McCarty, M., & Avery, O. T. (1946). Studies on the Chemical Nature of the Substance Inducing Transformation of Pneumococcal types: III. An Improved Method for the Isolation of the Transforming Substance and its Application to Pneumo coccus Types

[5] Spizizen, J. (1958). Transformation of Biochemically Deficient Strains of Bacillus sub‐

[6] Mandel, M., & Higa, A. (1970). Calcium-dependent Bacteriophage DNA Infection. *J.*

[7] Aune, T. E., & Aachmann, F. L. (2010). Methodologies to Increase the Transformation Efficiencies and the Range of Bacteria that can be Transformed. *Appl. microbiol. bio‐*

[8] Roberts, R. J., Belfort, M., Bestor, T., Bhagwat, A. S., Bickle, T. A., Bitinaite, J., Blu‐ menthal, R. M., Degtyarev, S. K., Dryden, D. T. F., Dybvig, K., Firman, K., Gromova, E. S., Gumport, R. I., Halford, S. E., Hattman, S., Heitman, J., Hornby, D. P., Janulai‐ tis, A., Jeltsch, A., Josephsen, J., Kiss, A., Klaenhammer, T. R., Kobayashi, I., Kong, H., Krüger, D. H., Lacks, S., Marinus, M. G., Miyahara, M., Morgan, R. D., Murray, N. E., Nagaraja, V., Piekarowicz, A., Pingoud, A., Raleigh, E., Rao, D. N., Reich, N., Repin, V. E., Selker, E. U., Shaw, P. C., Stein, D. C., Stoddard, B. L., Szybalski, W., Trautner, T. A., Van Etten, J. L., Vitor, J. M. B., Wilson, G. G., & Xu, S. Y. (2003). A Nomencla‐ ture for Restriction Enzymes, DNA Methyltransferases, Homing Endonucleases and

[9] Arber, W., & Dussoix, D. (1962). Host Specificity of DNA Produced by Escherichia coli: I. Host Controlled Modification of Bacteriophage. *J. mol. biol.*, 5, 18-36.

[10] Roberts, R. J., Vincze, T., Posfai, J., & Macelis, D. (2003). REBASE: Restriction En‐

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[12] Kobayashi, I. (2001). Behavior of Restriction-Modification Systems as Selfish Mobile Elements and their Impact on Genome Evolution. *Nucleic acids res.*, 29, 3742-3756. [13] Snyder, L., Gold, L., & Kutter, E. (1976). A Gene of Bacteriophage T4 whose Product Prevents True Late Transcription on Cytosine-containing T4 DNA. *Proc. natl. acad.*

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the Biological Activity of the Transforming Substance. *J. exp. med.*, 83, 89-96.

II, III, and VI. *J. exp. med.*, 83, 97-104.

their Genes. *Nucleic acids res.*, 31, 1805-1812.

*mol. biol.*, 53, 159-162.

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234-D236.

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#### **4. Conclusion**

In this chapter, RM systems are reviewed and some approaches to produce host-mimicking DNA are described. Analysis of chromosomal DNA using HPLC is a simple method to elu‐ cidate functional RM systems in target bacterium, and is therefore highly recommended for establishing bacterial transformation methods. When negligible DNA methylation is ob‐ served, methyl-free DNA is suitable for transformation of the bacterium. On the other hand, when significant DNA methylation is observed, a host-mimicking strategy involving meth‐ ylation needs to be utilized. One weak point of this strategy is that there are no methods to exhaustively, readily, and rapidly determine methylation sites in target bacterium. When this analytical method becomes more widespread, this strategy will become a crucial techni‐ que for establishing efficient bacterial transformation methods.

#### **Acknowledgements**

This work was supported by Grant-in-Aid for Young Scientists (B) of Japan Society for the Promotion of Science (20780080 and in part 23750083). The author would like to thank Ena‐ go (www.enago.jp) for the English language review.

#### **Author details**

Hirokazu Suzuki\*

Address all correspondence to: hirokap@xpost.plala.or.jp

Faculty of Agriculture, Kyushu University, Japan

#### **References**


[3] McCarty, M., & Avery, O. T. (1946). Studies on the Chemical Nature of the Substance Inducing Transformation of Pneumococcal Types: II. Effect of Desoxyribonuclease on the Biological Activity of the Transforming Substance. *J. exp. med.*, 83, 89-96.

inactivating RM systems are not predictable and thereby may have to be determined by re‐

In this chapter, RM systems are reviewed and some approaches to produce host-mimicking DNA are described. Analysis of chromosomal DNA using HPLC is a simple method to elu‐ cidate functional RM systems in target bacterium, and is therefore highly recommended for establishing bacterial transformation methods. When negligible DNA methylation is ob‐ served, methyl-free DNA is suitable for transformation of the bacterium. On the other hand, when significant DNA methylation is observed, a host-mimicking strategy involving meth‐ ylation needs to be utilized. One weak point of this strategy is that there are no methods to exhaustively, readily, and rapidly determine methylation sites in target bacterium. When this analytical method becomes more widespread, this strategy will become a crucial techni‐

This work was supported by Grant-in-Aid for Young Scientists (B) of Japan Society for the Promotion of Science (20780080 and in part 23750083). The author would like to thank Ena‐

[1] Griffith, F. (1928). The Significance of Pneumococcal Types. *J. hyg. (Lond)*, 27, 113-159. [2] Avery, O. T., MacLeod, C. M., & McCarty, M. (1944). Studies on the Chemical Nature of the Substance Inducing Transformation of Pneumococcal Types: Induction of Transformation by a Desoxyribonucleic Acid Fraction Isolated from Pneumococcus

que for establishing efficient bacterial transformation methods.

go (www.enago.jp) for the English language review.

230 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Address all correspondence to: hirokap@xpost.plala.or.jp

Faculty of Agriculture, Kyushu University, Japan

Type III. *J. exp. med.*, 79, 137-158.

peated trials.

**4. Conclusion**

**Acknowledgements**

**Author details**

Hirokazu Suzuki\*

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**Chapter 10**

**Messenger RNA Cap Methylation in Vesicular**

**Negative‐Sense RNA Virus**

Additional information is available at the end of the chapter

DA due to their potential as agents of agricultural bioterrorism [5].

Jianrong Li and Yu Zhang

http://dx.doi.org/10.5772/54598

**1. Introduction**

**Stomatitis Virus, a Prototype of Non‐Segmented**

The non‐segmented negative‐sense (NNS) RNA viruses encompass a wide range of signifi‐ cant human, animal, and plant pathogens including several National Institute of Allergy and Infectious Diseases (NIAID) Category A and C biodefense pathogens. The NNS RNA viruses are classified into four families: the *Rhabdoviridae,* as exemplified by rabies virus and vesicular stomatitis virus (VSV); the *Paramyxoviridae,* as exemplified by human respiratory syncytial virus (RSV), human metapneumovirus (hMPV), human parainfluenza virus type 3 (PIV3), measles virus, mump virus, Newcastle disease virus (NDV), Nipah virus, and Hen‐ dra virus; the *Filoviridae,* as exemplified by Ebola and Marburg viruses; and the *Bornaviridae*, as exemplified by Borna disease virus. For many of these viruses, there are no effective vac‐ cines or anti-viral drugs. RSV, hMPV, and PIV3 account for more than 70% of acute viral respiratory diseases, especially in infants, children, and the elderly [1, 2]. hPIV 1-3 have been recognized as the causative agents of croup since the late 1950's [3]. In addition, measles re‐ mains a major killer of children worldwide, despite successful vaccination programs in de‐ veloped countries [4]. The most virulent strains of NDV, the viscerotropic velogenic strains (often called "exotic" NDV), are classified as High Consequence Livestock Pathogens by US‐

Messenger RNA modification is the essential issue in NNS RNA virus gene expression and replication. During viral RNA synthesis, NNS RNA viruses produce capped, methylated, and polyadenylated mRNAs [6-8]. Cap formation is essential for mRNA stability, efficient transla‐ tion, and gene expression [9-11]. It is now firmly established that mRNA capping and methyla‐ tion in NNS RNA viruses evolves in a mechanism distinct to their hosts [12-19]. Thus, mRNA cap formation is an attractive antiviral target for NNS RNA viruses. For decades, VSV has been

and reproduction in any medium, provided the original work is properly cited.

© 2013 Li and Zhang; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,


## **Messenger RNA Cap Methylation in Vesicular Stomatitis Virus, a Prototype of Non‐Segmented Negative‐Sense RNA Virus**

Jianrong Li and Yu Zhang

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/54598

#### **1. Introduction**

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The non‐segmented negative‐sense (NNS) RNA viruses encompass a wide range of signifi‐ cant human, animal, and plant pathogens including several National Institute of Allergy and Infectious Diseases (NIAID) Category A and C biodefense pathogens. The NNS RNA viruses are classified into four families: the *Rhabdoviridae,* as exemplified by rabies virus and vesicular stomatitis virus (VSV); the *Paramyxoviridae,* as exemplified by human respiratory syncytial virus (RSV), human metapneumovirus (hMPV), human parainfluenza virus type 3 (PIV3), measles virus, mump virus, Newcastle disease virus (NDV), Nipah virus, and Hen‐ dra virus; the *Filoviridae,* as exemplified by Ebola and Marburg viruses; and the *Bornaviridae*, as exemplified by Borna disease virus. For many of these viruses, there are no effective vac‐ cines or anti-viral drugs. RSV, hMPV, and PIV3 account for more than 70% of acute viral respiratory diseases, especially in infants, children, and the elderly [1, 2]. hPIV 1-3 have been recognized as the causative agents of croup since the late 1950's [3]. In addition, measles re‐ mains a major killer of children worldwide, despite successful vaccination programs in de‐ veloped countries [4]. The most virulent strains of NDV, the viscerotropic velogenic strains (often called "exotic" NDV), are classified as High Consequence Livestock Pathogens by US‐ DA due to their potential as agents of agricultural bioterrorism [5].

Messenger RNA modification is the essential issue in NNS RNA virus gene expression and replication. During viral RNA synthesis, NNS RNA viruses produce capped, methylated, and polyadenylated mRNAs [6-8]. Cap formation is essential for mRNA stability, efficient transla‐ tion, and gene expression [9-11]. It is now firmly established that mRNA capping and methyla‐ tion in NNS RNA viruses evolves in a mechanism distinct to their hosts [12-19]. Thus, mRNA cap formation is an attractive antiviral target for NNS RNA viruses. For decades, VSV has been

© 2013 Li and Zhang; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

used as a model to understand the replication and gene expression of NNS RNA viruses. Most of our understanding of mRNA modifications of NNS RNA viruses comes from studies of VSV, a prototype of the *Rhabdoviridae* family. Using VSV as a model, we will discuss (i) the un‐ usual mechanism of mRNA capping and cap methylation; (ii) the impact of viral mRNA cap methylation in viral life cycle and viral pathogenesis; and (iii) the applications of viral mRNA cap methylation in the development of novel vaccines and broadly-active anti-viral agents.

#### **2. Overview diagram of VSV mRNA synthesis and modifications**

#### **2.1. The structure of VSV virions**

VSV virions are bullet-shaped particles 170 nm in length and 80 nm in diameter (Fig.1). Among NNS RNA viruses, VSV has the simplest RNA genome consisting of 11,161 nucleotides (nt) or‐ ganized into five VSV genes encoding nucleocapsid (N), phospho- (P), matrix (M), glyco- (G), and large (L) proteins, and leader and trailer regulatory sequences arranged in the order 3'- (leader), N, P, M, G, L, (trailer)-5'[20-23]. Like all NNS RNA viruses, the genome is encapsidated with the N protein to form a nuclease-resistant helical N-RNA complex that is the functional template for mRNA synthesis as well as genomic RNA replication. The N-RNA complex is tight‐ ly associated with the viral RNA-dependent RNA polymerase (RdRp), which is comprised of the 241-kDa L protein catalytic subunit and the 29-kDa essential P protein cofactor, and results in the assembly of a viral ribonucleoprotein (RNP) complex [24, 25]. This structure contains the minimum virus encoded components of the VSV RNA synthesis machinery [26]. The RNP com‐ plex is further surrounded by the M protein which plays a crucial role in virus assembly, bud‐ ding, and maintenance of the structural integrity of the virus particle [27]. The outer membrane of virion is the envelope composed of a cellular lipid bilayer. The transmembrane G protein is anchored in the viral envelope, which is essential for receptor binding and cell entry [28].

**Figure 1. VSV virion structure and genome organization.** VSV encodes five structural proteins**:** nucleocapsid (N), phospho- (P), matrix (M), glyco- (G), and large (L) proteins. The VSV genome is arranged in the order 3'-(leader), N, P,

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**Figure 2. Overview diagram of VSV life cycle**. Steps of virus life cycle: attachment, endocytosis, uncoating, genome

replication, mRNA transcription, viral protein translation, viral assembly, and budding are shown.

M, G, L, (trailer)-5'.

#### **2.2. VSV life cycle**

The overview picture of VSV life cycle is depicted in Fig.2. Upon attaching to an unknown cell receptor(s), VSV enters host cells via receptor mediated endocytosis [29]. Following low pH triggered fusion and uncoating, the RNP complex is delivered into the cytoplasm where RNA synthesis and viral replication occur [30]. During primary transcription, the input RdRp recog‐ nizes the specific signals in the N-RNA template to transcribe six discrete RNAs: a 47-nucleo‐ tide leader RNA (Le+), which is neither capped nor polyadenylated, and 5 mRNAs that are capped and methylated at the 5' end and polyadenylated at the 3'end. These mature mRNAs are then translated by host ribosomes to yield functional viral proteins which are required for viral genome replication. During replication, the RdRP initiates at the extreme 3' end of the ge‐ nome and synthesizes a full-length complementary antigenome, which subsequently serves as template for synthesis of full-length progeny genomes. These progeny genomes can then be utilized as templates for secondary transcription, or assembled into infectious particles. Final‐ ly, viral proteins and genomic RNA are assembled into complete virus particles and the virus exits the cell by budding through the plasma membrane.

Messenger RNA Cap Methylation in Vesicular Stomatitis Virus, a Prototype of Non‐Segmented Negative‐Sense RNA Virus http://dx.doi.org/10.5772/54598 239

used as a model to understand the replication and gene expression of NNS RNA viruses. Most of our understanding of mRNA modifications of NNS RNA viruses comes from studies of VSV, a prototype of the *Rhabdoviridae* family. Using VSV as a model, we will discuss (i) the un‐ usual mechanism of mRNA capping and cap methylation; (ii) the impact of viral mRNA cap methylation in viral life cycle and viral pathogenesis; and (iii) the applications of viral mRNA cap methylation in the development of novel vaccines and broadly-active anti-viral agents.

VSV virions are bullet-shaped particles 170 nm in length and 80 nm in diameter (Fig.1). Among NNS RNA viruses, VSV has the simplest RNA genome consisting of 11,161 nucleotides (nt) or‐ ganized into five VSV genes encoding nucleocapsid (N), phospho- (P), matrix (M), glyco- (G), and large (L) proteins, and leader and trailer regulatory sequences arranged in the order 3'- (leader), N, P, M, G, L, (trailer)-5'[20-23]. Like all NNS RNA viruses, the genome is encapsidated with the N protein to form a nuclease-resistant helical N-RNA complex that is the functional template for mRNA synthesis as well as genomic RNA replication. The N-RNA complex is tight‐ ly associated with the viral RNA-dependent RNA polymerase (RdRp), which is comprised of the 241-kDa L protein catalytic subunit and the 29-kDa essential P protein cofactor, and results in the assembly of a viral ribonucleoprotein (RNP) complex [24, 25]. This structure contains the minimum virus encoded components of the VSV RNA synthesis machinery [26]. The RNP com‐ plex is further surrounded by the M protein which plays a crucial role in virus assembly, bud‐ ding, and maintenance of the structural integrity of the virus particle [27]. The outer membrane of virion is the envelope composed of a cellular lipid bilayer. The transmembrane G protein is

anchored in the viral envelope, which is essential for receptor binding and cell entry [28].

exits the cell by budding through the plasma membrane.

The overview picture of VSV life cycle is depicted in Fig.2. Upon attaching to an unknown cell receptor(s), VSV enters host cells via receptor mediated endocytosis [29]. Following low pH triggered fusion and uncoating, the RNP complex is delivered into the cytoplasm where RNA synthesis and viral replication occur [30]. During primary transcription, the input RdRp recog‐ nizes the specific signals in the N-RNA template to transcribe six discrete RNAs: a 47-nucleo‐ tide leader RNA (Le+), which is neither capped nor polyadenylated, and 5 mRNAs that are capped and methylated at the 5' end and polyadenylated at the 3'end. These mature mRNAs are then translated by host ribosomes to yield functional viral proteins which are required for viral genome replication. During replication, the RdRP initiates at the extreme 3' end of the ge‐ nome and synthesizes a full-length complementary antigenome, which subsequently serves as template for synthesis of full-length progeny genomes. These progeny genomes can then be utilized as templates for secondary transcription, or assembled into infectious particles. Final‐ ly, viral proteins and genomic RNA are assembled into complete virus particles and the virus

**2. Overview diagram of VSV mRNA synthesis and modifications**

**2.1. The structure of VSV virions**

238 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**2.2. VSV life cycle**

**Figure 1. VSV virion structure and genome organization.** VSV encodes five structural proteins**:** nucleocapsid (N), phospho- (P), matrix (M), glyco- (G), and large (L) proteins. The VSV genome is arranged in the order 3'-(leader), N, P, M, G, L, (trailer)-5'.

**Figure 2. Overview diagram of VSV life cycle**. Steps of virus life cycle: attachment, endocytosis, uncoating, genome replication, mRNA transcription, viral protein translation, viral assembly, and budding are shown.

#### **2.3. VSV mRNA synthesis and modifications**

Our current understanding of VSV mRNA synthesis and modification can be summarized as follows. In response to a specific promoter element provided by the genomic leader re‐ gion, the polymerase initiates mRNA synthesis at the first gene-start sequence to synthesize N gene. The nascent mRNA is capped through an unconventional mechanism in which the GDP: polyribonucleotidyltransferase (PRNTase) of L transfers a monophosphate RNA onto a GDP acceptor through a covalent protein-RNA intermediate [12, 13, 16]. Following cap ad‐ dition, VSV mRNAs are sequentially methylated at ribose 2'-O position and G-N-7 position, which is distinct from all known methylation reactions [17, 18]. Unlike traditional cap form‐ ing enzymes, the VSV capping and methylation machinery requires *cis-*acting signals in the RNA [12, 18, 31, 32]. When encountering a gene-end sequence, L polyadenylates and termi‐ nates mRNA synthesis by a programmed stuttering of the polymerase on a U7 tract [33, 34]. Termination at the end of the N gene is essential for the polymerase to initiate synthesis at the start of the next gene, to produce the P mRNA. During transcription, the RdRp complex transcribes the viral genome into five mRNAs in a sequential and gradient manner, such that 3′ proximal genes are transcribed more abundantly than 3′ distal genes [21-23]. This gradient of transcription reflects a poorly understood transcriptional attenuation event that is localized to the gene junction regions. Using this fashion, VSV produces five capped, me‐ thylated, and polyadenylated mRNAs, N, P, M, G, and L.

tive strand RdRPs. Mutations to the GDN motif of the VSV L protein inactivated polymerase function [40]. The mRNA capping enzyme maps to CR V [13, 16], and the capping activities of L differ from those of other viruses and their eukaryotic hosts. Specifically, an RNA:GDP PRNTase activity present within CR V transfers 5′ monophosphate RNA onto a GDP acceptor through a covalent L-pRNA intermediate [12, 13, 16]. The mRNA cap methyltransferases (MTase) map to CR VI [14, 17, 19]. Like the unconventional capping enzyme, methylation of the VSV mRNA cap structure is also unique in that mRNA cap is modified by a dual specificity MTase activity within CR VI whereby ribose 2′-O methylation precedes and facilitates subse‐ quent guanine-N-7 (G-N-7) methylation [17, 18]. Although functions have not yet to be as‐ signed to the other three conserved regions (CRs I, II, and IV), experiments with Sendai virus (SeV) have implicated CR I in binding P protein and CR II in binding the RNA template [41, 42].

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**Figure 3. Conserved regions in L proteins of NNS RNA viruses**. Six conserved regions numbered I to VI (CRs I–VI) in L pro‐ tein are shown. Signature motifs for nucleotide polymerization, mRNA cap addition, and cap methylation are shown.

The location of the nucleotide polymerization, capping, and cap methylation activities with‐ in separate regions of L has led to the notion that L protein may be organized as a series of independent structural domains. Consistent with this idea, a fragment containing CRs V and VI of the SeV L protein were expressed independently and shown to retain the ability to methylate short RNAs that corresponded to the 5′ end of SeV mRNA [43]. In addition, re‐ combinant VSV and measles virus can be recovered from infectious cDNA clones by insert‐ ing the coding sequence of green fluorescent protein between CR V and VI in L gene, suggesting that L protein folds and functions as a series of independent globular domains [44, 45]. Interestingly, mutations to a variable region between CRs V and VI (residues 1450– 1481) affect mRNA cap MTase activity, feasibly suggesting that mutation to this hinge re‐ gion may affect a conformational change in CR VI [46]. More recently, the molecular archi‐ tecture of VSV L protein has been revealed using negative stain electron microscopy (EM) in combination with proteolytic digestion and deletion mapping [37]. It was found that VSV L protein is organized into a ring domain containing the RNA polymerase and an appendage of three globular domains containing the cap-forming activities. The capping enzyme maps to a globular domain, which is juxtaposed to the ring, and the cap methyltransferase maps to a more distal and flexibly connected globule. Interestingly, upon binding to P protein, L protein undergoes a significant structural rearrangement that may facilitate the coordination

between mRNA synthesis and capping apparatus [37, 47].

#### **3. Large (L) polymerase protein, the multifunctional protein that modifies viral mRNA**

All NNS RNA viruses encode a large (L) polymerase protein, a multifunctional protein rang‐ ing from 220-250kDa in molecular weight. The L protein contains enzymatic activities for nu‐ cleotide polymerization, mRNA cap addition, cap methylation, and polyadenylation. To date, the structure of L protein, or L protein fragments, has not been determined for any of the NNS RNA viruses. Amino acid sequence alignment between the L proteins of representative mem‐ bers of each family within NNS RNA viruses has identified six conserved regions numbered I to VI (CRs I–VI) (Fig.3) [35]. Thus, there is a general assent that the enzymatic activities of L pro‐ tein are located in these conserved regions. For the last four decades, VSV L protein has been used a model to understand the different activities of NNS RNA virus L proteins because it is the only member of this order of viruses for which robust transcription can be reconstituted *in vitro* [6, 7, 21, 36]. In addition, the VSV L protein can be highly expressed in recombinant expres‐ sion systems, such as E.coli and insect cells. The purified VSV L protein retains all the enzymat‐ ic activities that can modify short virus-specific mRNA *in trans* [12, 16, 18, 37]. In recent years, many breakthroughs have been made in the characterization of the function of VSV L protein and the enzymatic activities have been mapped at the single amino acid level (Fig.3). Within the primary sequence of L are six conserved regions shared among all NNS RNA virus L proteins. The RdRP activity has been identified in CR III and this region is also required for polyadenyla‐ tion [38-40]. Consistent with this, a GDN motif is conserved in CR III of all NNS RNA virus L proteins and is functionally equivalent to the GDD polymerization motif characteristic of posi‐ tive strand RdRPs. Mutations to the GDN motif of the VSV L protein inactivated polymerase function [40]. The mRNA capping enzyme maps to CR V [13, 16], and the capping activities of L differ from those of other viruses and their eukaryotic hosts. Specifically, an RNA:GDP PRNTase activity present within CR V transfers 5′ monophosphate RNA onto a GDP acceptor through a covalent L-pRNA intermediate [12, 13, 16]. The mRNA cap methyltransferases (MTase) map to CR VI [14, 17, 19]. Like the unconventional capping enzyme, methylation of the VSV mRNA cap structure is also unique in that mRNA cap is modified by a dual specificity MTase activity within CR VI whereby ribose 2′-O methylation precedes and facilitates subse‐ quent guanine-N-7 (G-N-7) methylation [17, 18]. Although functions have not yet to be as‐ signed to the other three conserved regions (CRs I, II, and IV), experiments with Sendai virus (SeV) have implicated CR I in binding P protein and CR II in binding the RNA template [41, 42].

**2.3. VSV mRNA synthesis and modifications**

240 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

thylated, and polyadenylated mRNAs, N, P, M, G, and L.

**modifies viral mRNA**

**3. Large (L) polymerase protein, the multifunctional protein that**

All NNS RNA viruses encode a large (L) polymerase protein, a multifunctional protein rang‐ ing from 220-250kDa in molecular weight. The L protein contains enzymatic activities for nu‐ cleotide polymerization, mRNA cap addition, cap methylation, and polyadenylation. To date, the structure of L protein, or L protein fragments, has not been determined for any of the NNS RNA viruses. Amino acid sequence alignment between the L proteins of representative mem‐ bers of each family within NNS RNA viruses has identified six conserved regions numbered I to VI (CRs I–VI) (Fig.3) [35]. Thus, there is a general assent that the enzymatic activities of L pro‐ tein are located in these conserved regions. For the last four decades, VSV L protein has been used a model to understand the different activities of NNS RNA virus L proteins because it is the only member of this order of viruses for which robust transcription can be reconstituted *in vitro* [6, 7, 21, 36]. In addition, the VSV L protein can be highly expressed in recombinant expres‐ sion systems, such as E.coli and insect cells. The purified VSV L protein retains all the enzymat‐ ic activities that can modify short virus-specific mRNA *in trans* [12, 16, 18, 37]. In recent years, many breakthroughs have been made in the characterization of the function of VSV L protein and the enzymatic activities have been mapped at the single amino acid level (Fig.3). Within the primary sequence of L are six conserved regions shared among all NNS RNA virus L proteins. The RdRP activity has been identified in CR III and this region is also required for polyadenyla‐ tion [38-40]. Consistent with this, a GDN motif is conserved in CR III of all NNS RNA virus L proteins and is functionally equivalent to the GDD polymerization motif characteristic of posi‐

Our current understanding of VSV mRNA synthesis and modification can be summarized as follows. In response to a specific promoter element provided by the genomic leader re‐ gion, the polymerase initiates mRNA synthesis at the first gene-start sequence to synthesize N gene. The nascent mRNA is capped through an unconventional mechanism in which the GDP: polyribonucleotidyltransferase (PRNTase) of L transfers a monophosphate RNA onto a GDP acceptor through a covalent protein-RNA intermediate [12, 13, 16]. Following cap ad‐ dition, VSV mRNAs are sequentially methylated at ribose 2'-O position and G-N-7 position, which is distinct from all known methylation reactions [17, 18]. Unlike traditional cap form‐ ing enzymes, the VSV capping and methylation machinery requires *cis-*acting signals in the RNA [12, 18, 31, 32]. When encountering a gene-end sequence, L polyadenylates and termi‐ nates mRNA synthesis by a programmed stuttering of the polymerase on a U7 tract [33, 34]. Termination at the end of the N gene is essential for the polymerase to initiate synthesis at the start of the next gene, to produce the P mRNA. During transcription, the RdRp complex transcribes the viral genome into five mRNAs in a sequential and gradient manner, such that 3′ proximal genes are transcribed more abundantly than 3′ distal genes [21-23]. This gradient of transcription reflects a poorly understood transcriptional attenuation event that is localized to the gene junction regions. Using this fashion, VSV produces five capped, me‐

**Figure 3. Conserved regions in L proteins of NNS RNA viruses**. Six conserved regions numbered I to VI (CRs I–VI) in L pro‐ tein are shown. Signature motifs for nucleotide polymerization, mRNA cap addition, and cap methylation are shown.

The location of the nucleotide polymerization, capping, and cap methylation activities with‐ in separate regions of L has led to the notion that L protein may be organized as a series of independent structural domains. Consistent with this idea, a fragment containing CRs V and VI of the SeV L protein were expressed independently and shown to retain the ability to methylate short RNAs that corresponded to the 5′ end of SeV mRNA [43]. In addition, re‐ combinant VSV and measles virus can be recovered from infectious cDNA clones by insert‐ ing the coding sequence of green fluorescent protein between CR V and VI in L gene, suggesting that L protein folds and functions as a series of independent globular domains [44, 45]. Interestingly, mutations to a variable region between CRs V and VI (residues 1450– 1481) affect mRNA cap MTase activity, feasibly suggesting that mutation to this hinge re‐ gion may affect a conformational change in CR VI [46]. More recently, the molecular archi‐ tecture of VSV L protein has been revealed using negative stain electron microscopy (EM) in combination with proteolytic digestion and deletion mapping [37]. It was found that VSV L protein is organized into a ring domain containing the RNA polymerase and an appendage of three globular domains containing the cap-forming activities. The capping enzyme maps to a globular domain, which is juxtaposed to the ring, and the cap methyltransferase maps to a more distal and flexibly connected globule. Interestingly, upon binding to P protein, L protein undergoes a significant structural rearrangement that may facilitate the coordination between mRNA synthesis and capping apparatus [37, 47].

#### **4. An unconventional mRNA capping mechanism in VSV**

#### **4.1. Conventional mechanism of mRNA capping in eukaryotic cells**

In eukaryotic cells, capping of mRNA is an early posttranscriptional event that is essential for subsequent processing, nuclear export, stability, and translation of mRNA [11, 48]. Cap formation is mediated by a series of enzymatic reactions (Fig.4A). First, the 5' triphosphate end of the nascent mRNA chain (5'pppN-RNA) is hydrolyzed by an RNA triphosphatase (RTPase) to yield the diphosphate 5' ppN-RNA. Second, an RNA guanylyltransferase (GTase) reacts with GTP to form a covalent enzyme-GMP intermediate and transfers GMP to 5'ppN-RNA via a 5′-5′ triphosphate linkage to yield 5' GpppN-RNA. Typically, RNA GTases contain a signature Kx[D/N]G motif that functions as an active site for the capping reaction [11, 48, 49]. A lysine residue within Kx[D/N]G motif forms the enzyme-GMP cova‐ lent intermediate, prior to its transfer onto the diphosphate RNA acceptor [11, 48, 50]. This mRNA capping reaction is conserved among all eukaryotes.

unusual VSV capping enzyme caps RNA in a sequence specific manner [12]. Specifically, VSV L protein efficiently capped pppApApCpApG (the mRNA gene start sequence), but not pppApCpGpApA (the leader RNA start sequence). In addition, the VSV L protein was not able to cap ppApApCpApG, suggesting that the L protein specifically recognizes the 5′-triphos‐ phorylated AACAG. Further mutagenesis analysis has shown that the APuCNG (Pu, purine) sequence acts as a *cis*-acting element for the RNA capping reaction for the VSV L protein [12].

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**Figure 4. Comparison of mRNA cap formation in eukaryotic cells, WNV, and VSV. (A) Cellular mRNA cap forma‐ tion.** First, pppNNNN-RNA is hydrolyzed by an RNA triphosphatase (RTPase) to yield the diphosphate ppNNNN-RNA. Second, an RNA guanylyltransferase (GTase) transfers GMP to ppNNNN-RNA to yield GpppNNNN-RNA. Third, GpppNNNN-RNA is methylated by G-N-7 MTase to yield 7mGpppNNNN-RNA. Fourth, 7mGpppN-RNA is further methy‐ lated by a 2'-O MTase to yield 7mGpppNmNNN-RNA. **(B) WNV mRNA cap formation.** First, pppNNNN-RNA is hydro‐ lyzed by viral NS3 protein (RTPase) to yield ppNNNN-RNA. Second, viral NS5 protein (GTase) transfers GMP to ppNNNN-RNA to yield GpppNNNN-RNA. Third, GpppNNNN-RNA is methylated by N-terminus of NS5 (G-N-7 MTase) to yield 7mGpppNNNN-RNA. Fourth, 7mGpppN-RNA is further methylated by N-terminus of NS5 (2'-O MTase) to yield

Viruses are highly diverse in capping their mRNA. Many DNA viruses (such as vaccinia vi‐ rus and baculovirus), double stranded RNA viruses (such as reovirus, rotavirus, and blue‐ tongue virus), and single strand positive RNA viruses (such as West Nile virus, Fig.4B) utilize the conventional eukaryotic capping pathway [51-56]. It has been suggested that Kx[V/L/I]S motif serve as the GTase active site for reovirus and rotavirus [57]. Other viruses have evolved different mechanisms for acquiring their cap. For example, influenza virus and hantavirus furnish their mRNA with this structure by a cap-snatching mechanism, in which the viral polymerase steals host cell mRNA caps to prime viral mRNA synthesis [58, 59]. The alphaviruses, such as Sindbis, have evolved S-adenosyl-L-methionine(SAM*)*-de‐ pendent GTase activities that utilize distinct motifs (such as HxH motif) to transfer 7mGp through a covalent histidine Gp intermediate to form the 7mGpppN cap [60].

#### **4.2. An unconventional mRNA capping mechanism in VSV**

In the early 1970's, it was suggested that the cap structure of NNS viral mRNAs was formed by a mechanism which was unique from eukaryotic cap formation. For VSV [6], RSV [61], and spring viremia of carp virus [62], the two italicized phosphates of the 5′G*pp*p5′NpNpN triphosphate bridge have been shown to be derived from a GDP donor, rather than GMP. However, further studies on this mechanism have been seriously hampered due to the fact that the VSV capping events are tightly coupled to transcription and the capping machinery does not respond to exog‐ enous transcripts. In 2007, this unique capping mechanism was revealed using a novel *trans* cap‐ ping assay, in which a short mRNA corresponding to the first 5-nt of VSV gene start sequence was capped by a highly purified L protein in *trans* [12]. Specifically, capping of VSV mRNA was achieved by a novel polyribonucelotidyltransferase (PRNTase) which transferred a monophos‐ phate RNA onto a GDP acceptor through a covalent L-RNA intermediate (Fig. 4C). In the first step, a GTPase associated with the VSV L protein removes the γ-phosphate group of GTP to gen‐ erate GDP, an RNA acceptor. In the second step, the PRNTase activity of the L protein specifical‐ ly transfers a 5′-monophosphorylated (p-) RNA moiety of pppRNA with the conserved VSV mRNA-start sequence (AACAG) to GDP to yield a GpppA capped mRNA. Interestingly, this unusual VSV capping enzyme caps RNA in a sequence specific manner [12]. Specifically, VSV L protein efficiently capped pppApApCpApG (the mRNA gene start sequence), but not pppApCpGpApA (the leader RNA start sequence). In addition, the VSV L protein was not able to cap ppApApCpApG, suggesting that the L protein specifically recognizes the 5′-triphos‐ phorylated AACAG. Further mutagenesis analysis has shown that the APuCNG (Pu, purine) sequence acts as a *cis*-acting element for the RNA capping reaction for the VSV L protein [12].

**4. An unconventional mRNA capping mechanism in VSV**

In eukaryotic cells, capping of mRNA is an early posttranscriptional event that is essential for subsequent processing, nuclear export, stability, and translation of mRNA [11, 48]. Cap formation is mediated by a series of enzymatic reactions (Fig.4A). First, the 5' triphosphate end of the nascent mRNA chain (5'pppN-RNA) is hydrolyzed by an RNA triphosphatase (RTPase) to yield the diphosphate 5' ppN-RNA. Second, an RNA guanylyltransferase (GTase) reacts with GTP to form a covalent enzyme-GMP intermediate and transfers GMP to 5'ppN-RNA via a 5′-5′ triphosphate linkage to yield 5' GpppN-RNA. Typically, RNA GTases contain a signature Kx[D/N]G motif that functions as an active site for the capping reaction [11, 48, 49]. A lysine residue within Kx[D/N]G motif forms the enzyme-GMP cova‐ lent intermediate, prior to its transfer onto the diphosphate RNA acceptor [11, 48, 50]. This

Viruses are highly diverse in capping their mRNA. Many DNA viruses (such as vaccinia vi‐ rus and baculovirus), double stranded RNA viruses (such as reovirus, rotavirus, and blue‐ tongue virus), and single strand positive RNA viruses (such as West Nile virus, Fig.4B) utilize the conventional eukaryotic capping pathway [51-56]. It has been suggested that Kx[V/L/I]S motif serve as the GTase active site for reovirus and rotavirus [57]. Other viruses have evolved different mechanisms for acquiring their cap. For example, influenza virus and hantavirus furnish their mRNA with this structure by a cap-snatching mechanism, in which the viral polymerase steals host cell mRNA caps to prime viral mRNA synthesis [58, 59]. The alphaviruses, such as Sindbis, have evolved S-adenosyl-L-methionine(SAM*)*-de‐ pendent GTase activities that utilize distinct motifs (such as HxH motif) to transfer 7mGp

In the early 1970's, it was suggested that the cap structure of NNS viral mRNAs was formed by a mechanism which was unique from eukaryotic cap formation. For VSV [6], RSV [61], and spring viremia of carp virus [62], the two italicized phosphates of the 5′G*pp*p5′NpNpN triphosphate bridge have been shown to be derived from a GDP donor, rather than GMP. However, further studies on this mechanism have been seriously hampered due to the fact that the VSV capping events are tightly coupled to transcription and the capping machinery does not respond to exog‐ enous transcripts. In 2007, this unique capping mechanism was revealed using a novel *trans* cap‐ ping assay, in which a short mRNA corresponding to the first 5-nt of VSV gene start sequence was capped by a highly purified L protein in *trans* [12]. Specifically, capping of VSV mRNA was achieved by a novel polyribonucelotidyltransferase (PRNTase) which transferred a monophos‐ phate RNA onto a GDP acceptor through a covalent L-RNA intermediate (Fig. 4C). In the first step, a GTPase associated with the VSV L protein removes the γ-phosphate group of GTP to gen‐ erate GDP, an RNA acceptor. In the second step, the PRNTase activity of the L protein specifical‐ ly transfers a 5′-monophosphorylated (p-) RNA moiety of pppRNA with the conserved VSV mRNA-start sequence (AACAG) to GDP to yield a GpppA capped mRNA. Interestingly, this

through a covalent histidine Gp intermediate to form the 7mGpppN cap [60].

**4.1. Conventional mechanism of mRNA capping in eukaryotic cells**

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mRNA capping reaction is conserved among all eukaryotes.

**4.2. An unconventional mRNA capping mechanism in VSV**

**Figure 4. Comparison of mRNA cap formation in eukaryotic cells, WNV, and VSV. (A) Cellular mRNA cap forma‐ tion.** First, pppNNNN-RNA is hydrolyzed by an RNA triphosphatase (RTPase) to yield the diphosphate ppNNNN-RNA. Second, an RNA guanylyltransferase (GTase) transfers GMP to ppNNNN-RNA to yield GpppNNNN-RNA. Third, GpppNNNN-RNA is methylated by G-N-7 MTase to yield 7mGpppNNNN-RNA. Fourth, 7mGpppN-RNA is further methy‐ lated by a 2'-O MTase to yield 7mGpppNmNNN-RNA. **(B) WNV mRNA cap formation.** First, pppNNNN-RNA is hydro‐ lyzed by viral NS3 protein (RTPase) to yield ppNNNN-RNA. Second, viral NS5 protein (GTase) transfers GMP to ppNNNN-RNA to yield GpppNNNN-RNA. Third, GpppNNNN-RNA is methylated by N-terminus of NS5 (G-N-7 MTase) to yield 7mGpppNNNN-RNA. Fourth, 7mGpppN-RNA is further methylated by N-terminus of NS5 (2'-O MTase) to yield 7mGpppNmNNN-RNA. **(C) VSV mRNA cap formation.** First, a GTPase (CR V of L protein) removes the γ-phosphate group of GTP to generate GDP. Second, a polyribonucelotidyltransferase (PRNTase) (CR V of L protein) transfers a monophosphate RNA onto a GDP acceptor through a covalent L-RNA intermediate to GpppAACAG-RNA. Third, the cap structure is methylated a 2'-O MTase (CR VI of L protein) to yield GpppAmACAG-RNA. Fourth, GpppAmACAG-RNA is further methylated a G-N-7 MTase (CR VI of L protein) to yield 7mGpppAmACAG-RNA.

(MTase) to yield 7mGpppN-RNA (cap 0). Second, the G-N-7 methylated cap structure can then be further methylated by a ribose-2'-O (2'-O) MTase to yield 7mGpppNm-RNA (cap 1). During mRNA cap methylation, S-adenosyl-L-methionine (SAM) serves as the methyl do‐ nor, and the by-product S-adenosyl-homocysteine (SAH) is the competitive inhibitor of the SAM-dependent MTase. These mRNA cap methylation reactions are conserved among all eukaryotes. In this conventional methylation reaction, G-N-7 methylation occurs prior to 2'- O methylation and the two methylase activities are carried out by two separate enzymes,

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Many viruses encode their own mRNA cap methylaion machinery, the best-studied exam‐ ple of which is the poxvirus vaccinia virus. For vaccinia virus, the G-N-7 and 2'-O MTase activities are encoded by two separate viral proteins, D12L and VP39 [65, 68-70]. In the case of reovirus, G-N-7 and 2'-O MTases are catalyzed by two separate domains of the same viral polymerase protein [55, 71]. For VSV, G-N-7 and 2'-O MTases are accomplished by a single region (CR VI) located in the C terminus of viral polymerase protein, L (Fig.4C) [14, 17, 19]. Soon after the discovery of the dual MTase activities of VSV, the N terminus of flaviviruses polymerase protein (NS5) was found to encode both G-N-7 and 2'-O MTases (Fig.4B) [72-74]. In addition to this unusual dual MTase activity of CR VI, the order of mRNA cap methylation in VSV is unconventional in which 2'-O methylation precedes and facilitates the G-N-7 methylation [17, 18]. This is contrast to all known mRNA cap methylation reactions

**5.2. A single MTase catalytic site in CR-VI of L protein essential for both G-N-7 and 2'-O**

The SAM-dependent MTase superfamily contains a series of conserved motifs (X and I to VIII) [75]. The crystal structure of several known 2'-O MTases including *E. coli* heat shockinduced methyltransferase RrmJ/FtsJ and vaccinia virus VP39 have been solved[67, 68, 70, 76]. In RrmJ, a catalytic tetrad of residues: K38, D124, K164, and E199 formed the active site of 2'-O MTase [67, 76]. Site-directed mutagenesis of RrmJ found that a catalytic triad of resi‐ dues K38, D124, and K164 are essential for 2'-O MTase whereas E199 plays only a minor role in the methyltransferase reaction *in vitro*. In vaccinia virus VP39, four amino acids, K41, D138, K175, and E207, are essential for catalysis [68, 70]. By comparing the amino acid se‐ quence of the RrmJ and VP39 with CR VI of the L protein of NNS RNA viruses, it was sug‐ gested that this region of L protein might function as a 2′-O MTase. Sequence alignments suggest that residues K1651, D1762, K1795, and E1833 of the VSV L protein correspond to a catalytic KDKE tetrad (Fig.5). In fact, this KDKE motif is conserved in CR VI of L proteins of all NNS RNA viruses with the exception of Borna disease virus. Li et al., (2005) performed an extensive mutagenesis analysis in this predicted MTase catalytic KDKE tetrad in VSV L protein [14]. Recombinant VSVs carrying individual substitutions to K1651, D1762, K1795, and E1833 were recovered from an infectious cDNA clone of VSV. Analysis of the cap struc‐ ture of mRNA synthesized *in vitro* revealed that alterations to the predicted active site resi‐ dues abolished both G-N-7- and ribose 2′-O MTase activities. This result demonstrated that a single KDKE tetrad in CR-VI of the VSV L protein is essential for mRNA cap G-N-7- and

each containing its own binding site for the methyl donor, SAM.

including flaviviruses.

**methylation**

It has been a challenge to locate the active site for the novel PRNTase in the 241 kDa L protein. The only suggestive information regarding the location of the capping enzyme in L protein has come from the study of a novel inhibitor of the RSV polymerase which resulted in the synthesis of short uncapped viral RNAs *in vitro* [63]. Viral mutants resistant to this inhibitor were select‐ ed, and the resistance mutations were mapped to CR V, suggesting that CR V of L plays a role in mRNA cap formation. Sequence alignments of this region of L protein identified a total of 17 residues that were conserved among the NNS RNA viruses [16]. Guided by this information, an extensive mutagenesis analysis was performed within this region which led to the discov‐ ery of a new motif GxxT[n]HR composed of four amino acid residues (G1154A, T1157A, H1227A, and R1228A) in VSV L protein which are essential for mRNA cap formation [16]. *In vi‐ tro* RNA reconstitution assays have shown that these cap defective polymerases synthesized uncapped mRNAs that terminated prematurely. The size of these abortive transcripts ranged from 100 nt up to the full-length N mRNA, although the majority were less than 400 nt. Consis‐ tent with their inability to generate capped RNA during *in vitro* transcription reactions, G1154A, T1157A, H1227A, and R1228A were defective in *trans* capping of the 5-nt VSV gene start sequence, demonstrating that these amino acids in CR V of L protein are required for mRNA cap addition [16]. Importantly, GxxT[n]HR is highly conserved in the CR V of L pro‐ teins of all NNS RNA viruses, including Borna disease virus which replicates in nucleus. Fur‐ ther biochemical and mass spectrometric analyses found that H1227 in the conserved GxxT[n]HR motif of the VSV L protein is covalently linked to the 5′-monophosphate end of the RNA through a phosphoamide bond [13]. Therefore, amino acid residue H1227 is the active site of the PRNTase activity. Mutagenesis analysis also found that R1228A and R1228K muta‐ tions significantly decreased L-pRNA complex formation activities, suggesting that mutation in R1228 may affect the H1227-RNA intermediate formation [13]. Interestingly, this PRNTase activity was also found in L protein of Chandipura virus (CHPV), a rhabdovirus that is closely related to VSV [64]. Furthermore, mutations to HR motif in L protein of CHPV significantly re‐ duced the formation of the L-pRNA covalent intermediates in the PRNTase reaction. These re‐ sults demonstrate that this unconventional capping mechanism is conserved in the *Rhabdoviridae* family. Given the fact the HR motif is highly conserved in L proteins of NNS RNA viruses, it is likely that this novel capping mechanism is not only unique to rhadoviruses, but also may be utilized by other NNS RNA viruses.

#### **5. An unusual mechanism of mRNA cap methylation in VSV**

#### **5.1. Conventional mRNA cap methylation in eukaryotic cells**

In eukaryotic cells, the capped mRNA (GpppN-RNA) is typically methylated by two steps (Fig.4A) [65-68]. First, the capping guanylate is methylated by a G-N-7 methyltransferase (MTase) to yield 7mGpppN-RNA (cap 0). Second, the G-N-7 methylated cap structure can then be further methylated by a ribose-2'-O (2'-O) MTase to yield 7mGpppNm-RNA (cap 1). During mRNA cap methylation, S-adenosyl-L-methionine (SAM) serves as the methyl do‐ nor, and the by-product S-adenosyl-homocysteine (SAH) is the competitive inhibitor of the SAM-dependent MTase. These mRNA cap methylation reactions are conserved among all eukaryotes. In this conventional methylation reaction, G-N-7 methylation occurs prior to 2'- O methylation and the two methylase activities are carried out by two separate enzymes, each containing its own binding site for the methyl donor, SAM.

7mGpppNmNNN-RNA. **(C) VSV mRNA cap formation.** First, a GTPase (CR V of L protein) removes the γ-phosphate group of GTP to generate GDP. Second, a polyribonucelotidyltransferase (PRNTase) (CR V of L protein) transfers a monophosphate RNA onto a GDP acceptor through a covalent L-RNA intermediate to GpppAACAG-RNA. Third, the cap structure is methylated a 2'-O MTase (CR VI of L protein) to yield GpppAmACAG-RNA. Fourth, GpppAmACAG-RNA

It has been a challenge to locate the active site for the novel PRNTase in the 241 kDa L protein. The only suggestive information regarding the location of the capping enzyme in L protein has come from the study of a novel inhibitor of the RSV polymerase which resulted in the synthesis of short uncapped viral RNAs *in vitro* [63]. Viral mutants resistant to this inhibitor were select‐ ed, and the resistance mutations were mapped to CR V, suggesting that CR V of L plays a role in mRNA cap formation. Sequence alignments of this region of L protein identified a total of 17 residues that were conserved among the NNS RNA viruses [16]. Guided by this information, an extensive mutagenesis analysis was performed within this region which led to the discov‐ ery of a new motif GxxT[n]HR composed of four amino acid residues (G1154A, T1157A, H1227A, and R1228A) in VSV L protein which are essential for mRNA cap formation [16]. *In vi‐ tro* RNA reconstitution assays have shown that these cap defective polymerases synthesized uncapped mRNAs that terminated prematurely. The size of these abortive transcripts ranged from 100 nt up to the full-length N mRNA, although the majority were less than 400 nt. Consis‐ tent with their inability to generate capped RNA during *in vitro* transcription reactions, G1154A, T1157A, H1227A, and R1228A were defective in *trans* capping of the 5-nt VSV gene start sequence, demonstrating that these amino acids in CR V of L protein are required for mRNA cap addition [16]. Importantly, GxxT[n]HR is highly conserved in the CR V of L pro‐ teins of all NNS RNA viruses, including Borna disease virus which replicates in nucleus. Fur‐ ther biochemical and mass spectrometric analyses found that H1227 in the conserved GxxT[n]HR motif of the VSV L protein is covalently linked to the 5′-monophosphate end of the RNA through a phosphoamide bond [13]. Therefore, amino acid residue H1227 is the active site of the PRNTase activity. Mutagenesis analysis also found that R1228A and R1228K muta‐ tions significantly decreased L-pRNA complex formation activities, suggesting that mutation in R1228 may affect the H1227-RNA intermediate formation [13]. Interestingly, this PRNTase activity was also found in L protein of Chandipura virus (CHPV), a rhabdovirus that is closely related to VSV [64]. Furthermore, mutations to HR motif in L protein of CHPV significantly re‐ duced the formation of the L-pRNA covalent intermediates in the PRNTase reaction. These re‐ sults demonstrate that this unconventional capping mechanism is conserved in the *Rhabdoviridae* family. Given the fact the HR motif is highly conserved in L proteins of NNS RNA viruses, it is likely that this novel capping mechanism is not only unique to rhadoviruses, but

is further methylated a G-N-7 MTase (CR VI of L protein) to yield 7mGpppAmACAG-RNA.

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also may be utilized by other NNS RNA viruses.

**5. An unusual mechanism of mRNA cap methylation in VSV**

In eukaryotic cells, the capped mRNA (GpppN-RNA) is typically methylated by two steps (Fig.4A) [65-68]. First, the capping guanylate is methylated by a G-N-7 methyltransferase

**5.1. Conventional mRNA cap methylation in eukaryotic cells**

Many viruses encode their own mRNA cap methylaion machinery, the best-studied exam‐ ple of which is the poxvirus vaccinia virus. For vaccinia virus, the G-N-7 and 2'-O MTase activities are encoded by two separate viral proteins, D12L and VP39 [65, 68-70]. In the case of reovirus, G-N-7 and 2'-O MTases are catalyzed by two separate domains of the same viral polymerase protein [55, 71]. For VSV, G-N-7 and 2'-O MTases are accomplished by a single region (CR VI) located in the C terminus of viral polymerase protein, L (Fig.4C) [14, 17, 19]. Soon after the discovery of the dual MTase activities of VSV, the N terminus of flaviviruses polymerase protein (NS5) was found to encode both G-N-7 and 2'-O MTases (Fig.4B) [72-74]. In addition to this unusual dual MTase activity of CR VI, the order of mRNA cap methylation in VSV is unconventional in which 2'-O methylation precedes and facilitates the G-N-7 methylation [17, 18]. This is contrast to all known mRNA cap methylation reactions including flaviviruses.

#### **5.2. A single MTase catalytic site in CR-VI of L protein essential for both G-N-7 and 2'-O methylation**

The SAM-dependent MTase superfamily contains a series of conserved motifs (X and I to VIII) [75]. The crystal structure of several known 2'-O MTases including *E. coli* heat shockinduced methyltransferase RrmJ/FtsJ and vaccinia virus VP39 have been solved[67, 68, 70, 76]. In RrmJ, a catalytic tetrad of residues: K38, D124, K164, and E199 formed the active site of 2'-O MTase [67, 76]. Site-directed mutagenesis of RrmJ found that a catalytic triad of resi‐ dues K38, D124, and K164 are essential for 2'-O MTase whereas E199 plays only a minor role in the methyltransferase reaction *in vitro*. In vaccinia virus VP39, four amino acids, K41, D138, K175, and E207, are essential for catalysis [68, 70]. By comparing the amino acid se‐ quence of the RrmJ and VP39 with CR VI of the L protein of NNS RNA viruses, it was sug‐ gested that this region of L protein might function as a 2′-O MTase. Sequence alignments suggest that residues K1651, D1762, K1795, and E1833 of the VSV L protein correspond to a catalytic KDKE tetrad (Fig.5). In fact, this KDKE motif is conserved in CR VI of L proteins of all NNS RNA viruses with the exception of Borna disease virus. Li et al., (2005) performed an extensive mutagenesis analysis in this predicted MTase catalytic KDKE tetrad in VSV L protein [14]. Recombinant VSVs carrying individual substitutions to K1651, D1762, K1795, and E1833 were recovered from an infectious cDNA clone of VSV. Analysis of the cap struc‐ ture of mRNA synthesized *in vitro* revealed that alterations to the predicted active site resi‐ dues abolished both G-N-7- and ribose 2′-O MTase activities. This result demonstrated that a single KDKE tetrad in CR-VI of the VSV L protein is essential for mRNA cap G-N-7- and ribose 2′-O methylation [14]. Two models have been proposed to explain this result. One possibility is that CR VI functions as both G-N-7 and 2'-O MTases. However, this conflict the fact that all known G-N-7 and 2'-O MTases have distinct biochemistry during RNA methyl‐ ation reactions. An alternative explanation is that there is a sequential model for VSV mRNA cap methylation in which the product of one MTase acts as the substrate for the sec‐ ond (discussed below).

**5.3. A single SAM binding site in L protein essential for both G-N-7 and 2'-O methylation**

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The SAM-dependent MTase superfamily usually contains a G-rich motif (GxGxG) and an acidic residue (D/E) that is involved in SAM binding [75]. Sequence alignments between CR VI of NNS RNA virus L proteins and known MTases suggest that the SAM-binding residues of VSV L include G1670, G1672, G1674, G1675, and D1735 (Fig.5). Site-directed mutagenesis has been performed to define the roles of these amino acids in VSV mRNA methylation [17]. Each of these residues was individually substituted for alanine (A); or, for G4A, all four G residues were replaced with A; for G4AD, residue D1735 was also replaced with A. In addition, the flanking amino acid residues D1671 and S1673 within GDGSG motif were also substituted. Recombinant viruses were recovered from each of the *L* gene mutations. It was found that mutations to G1670, G1672, and S1673 specifical‐ ly diminished G-N-7, but not 2'-O methylation, suggesting that 2'-O methylation occurs prior to G-N-7 methylation in VSV [17]. In contrast, mutants D1671, G4A, G4AD, G1675A, and D1735 were defective in both 2'-O and G-N-7 methylations. Interestingly, mutant G1674A requires a higher concentration of SAM to achieve full methylation com‐ pared with wild type VSV and methylation is more sensitive to SAH inhibition. There‐ fore, amino acid substitutions to the predicted SAM binding site disrupted methylation at the G-N-7 position or at both the G-N-7 and ribose 2′-O positions of the mRNA cap. However, none of these mutants are specifically defective in 2'-O methylation alone. These studies provide genetic evidence that the two methylase activities share one single SAM binding site and, in contrast to other cap methylation reactions, methylation of the

**5.4. Mapping the potential RNA binding site that required for mRNA cap methylation**

To acquire methylation, the MTase usually directly or indirectly contacts an RNA sub‐ strate. This putative substrate binding site is poorly understood in NNS RNA viruses. However, this substrate binding site has been identified in several cellular and viral mRNA 2'-O MTases [67, 68, 70, 74, 77, 78]. To achieve 2'-O methylation, the RNA sub‐ strate interacts with the cap recognition site which requires stacking between the base of the cap and aromatic rings from a MTase [76, 79, 80]. Vaccinia VP39 is one of the best characterized 2'-O MTases. In VP39, it was found that the recognition of a methylated base is achieved by stacking between two aromatic residues (Y22 and F180) and the methyl group is in contact with residue Y204 (Fig.6A) [68, 70, 79]. In addition, the car‐ boxyl groups of residues D182 and E233 form hydrogen bonds with the NH and NH2 of the guanosine in VP39 (Fig.6A). Based on structure modeling and mutagenesis analysis, it was shown that residue F24 in West Nile virus (WNV) methylase (NS5) [81, 82] and Y29 and F173 in feline coronavirus 2'-O MTase (nsp16) [80] may play an equivalent role to residue Y22 in VP39 of vaccinia virus. The cellular cap binding protein-eukaryotic translation initiation factor 4E (eIF-4E) recognizes the cap by stacking between W56 and W102 [83]. In all known cases, aromatic residues are involved in cap binding and sub‐

G-N-7 position is not required for 2′-O methylation.

strate recognition.

**Figure 5.** Structure-based amino acid sequence alignments of conserved domain VI of representative NNS RNA virus L proteins with known 2'-O methyltransferase, the *E. coli* RrmJ and vaccinia virus VP39. The conserved motifs (X and I to VIII) correspond to the SAM-dependent MTase superfamily. MTase catalytic site is shown by pink color. SAM binding site are shown by yellow color. Conserved aromatic residues are shown by blue color. Predicted alpha-helical regions are shown by the cylinders and the ß-sheet regions by the arrows. STR, structure of RrmJ and predicted structure for the NNS RNA viruses; EBOM, Ebola virus; VSIV, VSV Indiana type; HRSV, human respiratory syncytial virus; RRMJ, *E. coli* heat shock 2'-O MTase; VP39, vaccinia virus 2'-O MTase VP39.

#### **5.3. A single SAM binding site in L protein essential for both G-N-7 and 2'-O methylation**

ribose 2′-O methylation [14]. Two models have been proposed to explain this result. One possibility is that CR VI functions as both G-N-7 and 2'-O MTases. However, this conflict the fact that all known G-N-7 and 2'-O MTases have distinct biochemistry during RNA methyl‐ ation reactions. An alternative explanation is that there is a sequential model for VSV mRNA cap methylation in which the product of one MTase acts as the substrate for the sec‐

**Figure 5.** Structure-based amino acid sequence alignments of conserved domain VI of representative NNS RNA virus L proteins with known 2'-O methyltransferase, the *E. coli* RrmJ and vaccinia virus VP39. The conserved motifs (X and I to VIII) correspond to the SAM-dependent MTase superfamily. MTase catalytic site is shown by pink color. SAM binding site are shown by yellow color. Conserved aromatic residues are shown by blue color. Predicted alpha-helical regions are shown by the cylinders and the ß-sheet regions by the arrows. STR, structure of RrmJ and predicted structure for the NNS RNA viruses; EBOM, Ebola virus; VSIV, VSV Indiana type; HRSV, human respiratory syncytial virus; RRMJ, *E. coli*

heat shock 2'-O MTase; VP39, vaccinia virus 2'-O MTase VP39.

ond (discussed below).

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The SAM-dependent MTase superfamily usually contains a G-rich motif (GxGxG) and an acidic residue (D/E) that is involved in SAM binding [75]. Sequence alignments between CR VI of NNS RNA virus L proteins and known MTases suggest that the SAM-binding residues of VSV L include G1670, G1672, G1674, G1675, and D1735 (Fig.5). Site-directed mutagenesis has been performed to define the roles of these amino acids in VSV mRNA methylation [17]. Each of these residues was individually substituted for alanine (A); or, for G4A, all four G residues were replaced with A; for G4AD, residue D1735 was also replaced with A. In addition, the flanking amino acid residues D1671 and S1673 within GDGSG motif were also substituted. Recombinant viruses were recovered from each of the *L* gene mutations. It was found that mutations to G1670, G1672, and S1673 specifical‐ ly diminished G-N-7, but not 2'-O methylation, suggesting that 2'-O methylation occurs prior to G-N-7 methylation in VSV [17]. In contrast, mutants D1671, G4A, G4AD, G1675A, and D1735 were defective in both 2'-O and G-N-7 methylations. Interestingly, mutant G1674A requires a higher concentration of SAM to achieve full methylation com‐ pared with wild type VSV and methylation is more sensitive to SAH inhibition. There‐ fore, amino acid substitutions to the predicted SAM binding site disrupted methylation at the G-N-7 position or at both the G-N-7 and ribose 2′-O positions of the mRNA cap. However, none of these mutants are specifically defective in 2'-O methylation alone. These studies provide genetic evidence that the two methylase activities share one single SAM binding site and, in contrast to other cap methylation reactions, methylation of the G-N-7 position is not required for 2′-O methylation.

#### **5.4. Mapping the potential RNA binding site that required for mRNA cap methylation**

To acquire methylation, the MTase usually directly or indirectly contacts an RNA sub‐ strate. This putative substrate binding site is poorly understood in NNS RNA viruses. However, this substrate binding site has been identified in several cellular and viral mRNA 2'-O MTases [67, 68, 70, 74, 77, 78]. To achieve 2'-O methylation, the RNA sub‐ strate interacts with the cap recognition site which requires stacking between the base of the cap and aromatic rings from a MTase [76, 79, 80]. Vaccinia VP39 is one of the best characterized 2'-O MTases. In VP39, it was found that the recognition of a methylated base is achieved by stacking between two aromatic residues (Y22 and F180) and the methyl group is in contact with residue Y204 (Fig.6A) [68, 70, 79]. In addition, the car‐ boxyl groups of residues D182 and E233 form hydrogen bonds with the NH and NH2 of the guanosine in VP39 (Fig.6A). Based on structure modeling and mutagenesis analysis, it was shown that residue F24 in West Nile virus (WNV) methylase (NS5) [81, 82] and Y29 and F173 in feline coronavirus 2'-O MTase (nsp16) [80] may play an equivalent role to residue Y22 in VP39 of vaccinia virus. The cellular cap binding protein-eukaryotic translation initiation factor 4E (eIF-4E) recognizes the cap by stacking between W56 and W102 [83]. In all known cases, aromatic residues are involved in cap binding and sub‐ strate recognition.

tion and completely blocked both G-N-7 and 2'-O methylation [84]. Based on the predicted structural model for the VSV MTase (amino acid residues from 1644 to 1842 in the L protein) (Fig.6B), the residues Y1650 and F1691 are located far from each other with a distance of 17.3 Å between their alpha carbon atoms. Y1650 is located in the middle of the first helix, and the F1691 is at the very C-terminal of the second helix. Perhaps, a stacking interaction with one aromatic residue causes a conformational and structural change in the VSV methylase, which results in the interaction with another aromatic residue. Changing of residue E1764 to D (maintenance of charge), Q (maintenance of size), or K (changing charge), even the very conservative change to D, dramatically inhibited both G-N-7 and 2'-O methylation [84]. The predicted structure of VSV MTase also shows that E1764, the residue adjacent to the catalyt‐ ic residue D1762, is exposed to the putative SAM binding site (Fig.6B). The side chain of E1764 shows close contact to the adenyl group of SAM (3.1 Å). In addition, it was found that Y1835A was found to require a higher SAM concentration to achieve full methylation and it

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To date, this work is the first attempt toward elucidation of the putative RNA substrate recogni‐ tion site in the L protein of NNS RNA viruses, which has shed light on the possible role of several conserved aromatic amino acids, including Y1650 and F1691, in RNA binding during cap meth‐ ylation. It would provide much more direct evidence for the role of these key amino acids in me‐ diating RNA binding if the RNA binding efficiency could be measured directly. Attempts to use a gel shift assay have failed to this end [84], as the existence of multiple RNA binding sites in L protein with a size as large as 241-kDa posed a tremendous challenge in discerning the effect of single point mutation. The use of a truncated CR VI of VSV L for in vitro RNA binding assays

For conventional mRNA cap methylation, two separate MTases sequentially methylated the cap structure, first at the G-N-7 position and subsequently at the ribose 2′-O position [65, 66]. Analysis of the cap methylation of mRNA synthesized *in vitro* suggests that mRNA cap methyla‐ tion in VSV is unusual, with methylation of ribose 2′-O occurring prior to G-N-7 methylation. First, early studies showed that at low concentrations of SAM, VSV mRNA was methylated at the 2'-O position only [85]. However, it could be chased into a doubly methylated cap structure at high SAM concentrations *in vitro*. Second, when *in vitro* mRNA synthesis was performed in the presence of MTase inhibitors such as SAH and sinefungin, G-N-7 methylation was inhibited prior to 2'-O methylation [86]. Third, a host range mutant of VSV, *hr*8, was shown to synthesize mRNA cap structures that lacked G-N-7 but were partially 2′-O-methylated [46, 87]. Finally, VSV mutants carrying mutations in the SAM binding site (such as G1670A, and G1672A) are spe‐

This unusual order of VSV mRNA cap methylation was also biochemically demonstrated by a *trans*-methylation assay in which both ribose 2′-O and G-N-7 MTases were recapitulated by using purified recombinant L and *in vitro*-synthesized RNA [18]. It was found that VSV L modifies the 2′-O position of the cap prior to the G-N-7 position and that G-N-7 methylation is diminished by pre-2′-O methylation of the substrate RNA [18], providing compelling evi‐

is more sensitive to MTase inhibitor [84].

might be a useful alternative strategy for future studies.

**5.5. An unusual order for mRNA cap methylation in VSV**

cifically defective in G-N-7, but not 2'-O methylation [17].

**Figure 6. The predicted structure of CR VI of VSV L protein**. **(A) Cap binding site of vaccinia virus 2'-O MTase, VP39**. The model was generated from the crystal structure of VP39 (PDB code: 1AV6) using PyMOL. The side chains of amino acid residues (Y22, F180, E233 and D182) involved in binding of 7mGp cap are shown as sticks. **(B) The predict‐ ed structure of CR VI of VSV L protein**. The model was generated based on the previous predicted structure of VSV MTase (amino acid residue from 1644 to 1842 in L protein) using PyMOL software. Alpha helices are shown in blue and beta strands are shown in pink. SAM and the side chains of critical residues are shown in sticks, and the oxygen atoms and nitrogen atoms are shown in red and blue, respectively. For SAM molecule, the carbon atoms are green. For the three important amino acids (Y1650, F1691 and E1764) that may be involved in RNA substrate binding, their side-chain carbon atoms are highlighted in orange. For the predicted catalytic residues (K1651, D1762, K1795 and E1833), the carbon atoms are shown in purple.

Guided by this information, the putative RNA binding site in VSV L protein was searched through mutagenesis analysis of selected conserved residues in region VI of VSV L protein that were physiochemically similar to those involved in substrate recognition in VP39. Se‐ quence alignment showed that there are a number of aromatic residues that are highly con‐ served in the MTase domain of L proteins of NNS RNA viruses (Fig. 5). Aromatic residues at positions 1650 (Y), 1691(F or Y), and 1835 (Y) are highly conserved in L proteins of NNS RNA viruses. Aromatic residues at positions 1742 (W), 1744 (Y), 1745 (F), and 1816 (F) are conserved in the L proteins of *Rhabdoviridae* and some *Paramyxoviridae* and *Filoviridae*. There‐ fore, these aromatic residues were selected as putative equivalents of Y22, F180, and Y204 in VP39. However, there is no amino acid precisely aligned with D182 and E233 in VP39. With the exception of two acidic amino acids in the catalytic site (K1651-D1762-K1795-E1833), po‐ sition E1764 is also conserved in all L proteins. Thus, E1764 was selected as a candidate for mimicking the role of VP39 residues D182 and E233. In addition, two serine mutations at the two most conserved positions at 1693 and 1827 of VSV L protein was also examined, based on the fact that it has been shown that a serine residue was involved in RNA-protein interac‐ tion in *E. coli* 2'-O MTase, RRMJ [67, 77]. To determine the role of these amino acid residues in mRNA cap methylation, a single point mutation was introduced to an infectious clone of VSV and recombinant VSVs harboring these mutations were recovered [84]. The importance of the maintenance of the aromatic ring at amino acids Y1650 and F1691 was revealed by the observation that the substitution of Y1650 and F1691 with two other possible aromatic resi‐ dues in the VSV infectious clone still produced viable recombinant viruses and produced a fully methylated mRNA cap, but alanine substitutions dramatically inhibited viral replica‐ tion and completely blocked both G-N-7 and 2'-O methylation [84]. Based on the predicted structural model for the VSV MTase (amino acid residues from 1644 to 1842 in the L protein) (Fig.6B), the residues Y1650 and F1691 are located far from each other with a distance of 17.3 Å between their alpha carbon atoms. Y1650 is located in the middle of the first helix, and the F1691 is at the very C-terminal of the second helix. Perhaps, a stacking interaction with one aromatic residue causes a conformational and structural change in the VSV methylase, which results in the interaction with another aromatic residue. Changing of residue E1764 to D (maintenance of charge), Q (maintenance of size), or K (changing charge), even the very conservative change to D, dramatically inhibited both G-N-7 and 2'-O methylation [84]. The predicted structure of VSV MTase also shows that E1764, the residue adjacent to the catalyt‐ ic residue D1762, is exposed to the putative SAM binding site (Fig.6B). The side chain of E1764 shows close contact to the adenyl group of SAM (3.1 Å). In addition, it was found that Y1835A was found to require a higher SAM concentration to achieve full methylation and it is more sensitive to MTase inhibitor [84].

To date, this work is the first attempt toward elucidation of the putative RNA substrate recogni‐ tion site in the L protein of NNS RNA viruses, which has shed light on the possible role of several conserved aromatic amino acids, including Y1650 and F1691, in RNA binding during cap meth‐ ylation. It would provide much more direct evidence for the role of these key amino acids in me‐ diating RNA binding if the RNA binding efficiency could be measured directly. Attempts to use a gel shift assay have failed to this end [84], as the existence of multiple RNA binding sites in L protein with a size as large as 241-kDa posed a tremendous challenge in discerning the effect of single point mutation. The use of a truncated CR VI of VSV L for in vitro RNA binding assays might be a useful alternative strategy for future studies.

#### **5.5. An unusual order for mRNA cap methylation in VSV**

**Figure 6. The predicted structure of CR VI of VSV L protein**. **(A) Cap binding site of vaccinia virus 2'-O MTase, VP39**. The model was generated from the crystal structure of VP39 (PDB code: 1AV6) using PyMOL. The side chains of amino acid residues (Y22, F180, E233 and D182) involved in binding of 7mGp cap are shown as sticks. **(B) The predict‐ ed structure of CR VI of VSV L protein**. The model was generated based on the previous predicted structure of VSV MTase (amino acid residue from 1644 to 1842 in L protein) using PyMOL software. Alpha helices are shown in blue and beta strands are shown in pink. SAM and the side chains of critical residues are shown in sticks, and the oxygen atoms and nitrogen atoms are shown in red and blue, respectively. For SAM molecule, the carbon atoms are green. For the three important amino acids (Y1650, F1691 and E1764) that may be involved in RNA substrate binding, their side-chain carbon atoms are highlighted in orange. For the predicted catalytic residues (K1651, D1762, K1795 and

Guided by this information, the putative RNA binding site in VSV L protein was searched through mutagenesis analysis of selected conserved residues in region VI of VSV L protein that were physiochemically similar to those involved in substrate recognition in VP39. Se‐ quence alignment showed that there are a number of aromatic residues that are highly con‐ served in the MTase domain of L proteins of NNS RNA viruses (Fig. 5). Aromatic residues at positions 1650 (Y), 1691(F or Y), and 1835 (Y) are highly conserved in L proteins of NNS RNA viruses. Aromatic residues at positions 1742 (W), 1744 (Y), 1745 (F), and 1816 (F) are conserved in the L proteins of *Rhabdoviridae* and some *Paramyxoviridae* and *Filoviridae*. There‐ fore, these aromatic residues were selected as putative equivalents of Y22, F180, and Y204 in VP39. However, there is no amino acid precisely aligned with D182 and E233 in VP39. With the exception of two acidic amino acids in the catalytic site (K1651-D1762-K1795-E1833), po‐ sition E1764 is also conserved in all L proteins. Thus, E1764 was selected as a candidate for mimicking the role of VP39 residues D182 and E233. In addition, two serine mutations at the two most conserved positions at 1693 and 1827 of VSV L protein was also examined, based on the fact that it has been shown that a serine residue was involved in RNA-protein interac‐ tion in *E. coli* 2'-O MTase, RRMJ [67, 77]. To determine the role of these amino acid residues in mRNA cap methylation, a single point mutation was introduced to an infectious clone of VSV and recombinant VSVs harboring these mutations were recovered [84]. The importance of the maintenance of the aromatic ring at amino acids Y1650 and F1691 was revealed by the observation that the substitution of Y1650 and F1691 with two other possible aromatic resi‐ dues in the VSV infectious clone still produced viable recombinant viruses and produced a fully methylated mRNA cap, but alanine substitutions dramatically inhibited viral replica‐

E1833), the carbon atoms are shown in purple.

248 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

For conventional mRNA cap methylation, two separate MTases sequentially methylated the cap structure, first at the G-N-7 position and subsequently at the ribose 2′-O position [65, 66]. Analysis of the cap methylation of mRNA synthesized *in vitro* suggests that mRNA cap methyla‐ tion in VSV is unusual, with methylation of ribose 2′-O occurring prior to G-N-7 methylation. First, early studies showed that at low concentrations of SAM, VSV mRNA was methylated at the 2'-O position only [85]. However, it could be chased into a doubly methylated cap structure at high SAM concentrations *in vitro*. Second, when *in vitro* mRNA synthesis was performed in the presence of MTase inhibitors such as SAH and sinefungin, G-N-7 methylation was inhibited prior to 2'-O methylation [86]. Third, a host range mutant of VSV, *hr*8, was shown to synthesize mRNA cap structures that lacked G-N-7 but were partially 2′-O-methylated [46, 87]. Finally, VSV mutants carrying mutations in the SAM binding site (such as G1670A, and G1672A) are spe‐ cifically defective in G-N-7, but not 2'-O methylation [17].

This unusual order of VSV mRNA cap methylation was also biochemically demonstrated by a *trans*-methylation assay in which both ribose 2′-O and G-N-7 MTases were recapitulated by using purified recombinant L and *in vitro*-synthesized RNA [18]. It was found that VSV L modifies the 2′-O position of the cap prior to the G-N-7 position and that G-N-7 methylation is diminished by pre-2′-O methylation of the substrate RNA [18], providing compelling evi‐ dence that 2′-O methylation precedes and facilitates G-N-7 methylation. In light that both two MTase activities appear to reside in the same domain of L protein with the same SAM binding site, it is conceivable that mRNA cap and/or the SAM binding site might need to be repositioned at the end of the first methylation reaction to facilitate the second round of methylation. How this coordination happens *in vitro* is still a mystery. Bearing in mind that G-N-7 position is upstream of the ribose 2'-O position in the mRNA strand, reorientation is thus less likely to have resulted from forward movement of capped RNA through CR VI during transcription, but rather it might entail a fine spatial rearrangement. Collectively, these experiments have shown that the order of VSV mRNA cap methylation is distinct from all other known mRNA cap methylation mechanisms.

**5.8. Model for mRNA cap methylation**

the binding of a subsequent molecule of SAM.

**5.9. Comparison of mRNA cap methylation in VSV and WNV**

The process of VSV L protein-mediated cap methylation can be best summarized with the following model (Fig.7). Initially in response to a specific *cis-*acting element in the VSV gene start sequence, CR VI of L protein methylates the cap structure first at the 2'-O position to produce GpppAmACAG-RNA. The by-product of this reaction, SAH, is released from this reaction. Following 2'-O methylation, a second molecule of SAM binds to CR VI of L protein that may facilitate a subsequent methylation of the RNA at the G-N-7 position. Methylation at the 2'-O position favors G-N-7 methylation in the cap structure through a currently un‐ known mechanism. G-N-7 methlyation may be facilitated by the contact with the RNA mol‐ ecule remains bound to the L protein at the end of the initial methylation at the 2'-O position. Another possibility is that CR VI of L protein exists in two different conformations. An initial conformation may favor binding of GpppRNA and SAM. Methylation of RNA at the 2'-O position perhaps induces a conformational change that facilitates the repositioning of the RNA for subsequent G-N-7 methylation, and/or favors the release of SAH as well as

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To date, the rhabdovirus, VSV, and the flavivirus, WNV, are the two best characterized vi‐ ruses that utilize a single region in the polymerase protein for both G-N-7 and 2'-O methyla‐ tions. However, the mechanism of VSV mRNA methylation is distinct from that of the WNV system (Fig.4B and C). In VSV, 2'-O methylation precedes and facilitates subsequent G-N-7 methylation [17, 18]. However, WNV MTases modify the cap structure, first at the G-N-7 position and subsequently at the ribose 2'-O position [72, 73, 78]. In VSV, the G-N-7 and 2'-O MTases require similar conditions for methylation with an optimal pH at 7.0 [18]. In con‐ trast, the G-N-7 and 2'-O MTases of WNV require an optimal pH at 6.5 and 10, respectively [72]. Both VSV and WNV MTases modify the RNA in a sequence-specific manner, but re‐ quire different elements in the RNA substrate. VSV G-N-7 and 2'-O MTases require specific gene start sequences with a minimum mRNA length of 10 nucleotides [18]. In the WNV model, N-7 cap methylation requires the presence of specific nucleotides at the second and third positions and a 5' stem-loop structure within the 74-nucleotide viral RNA; in contrast, 2'-O ribose methylation requires specific nucleotides at the first and second positions, with a minimum 5' viral RNA of 20 nucleotides in length [81]. In addition, there is striking differ‐ ence in the cap recognition site between the VSV and WNV MTases. For the WNV MTase, the cap recognition site is essential for 2'-O, but not G-N-7 methylation [73, 82]. Consistent with this finding, it was found that GTP and cap analogs specifically inhibited 2'-O, but not G-N-7 methylation [73, 82]. However, mutations to the putative RNA binding site in VSV L protein affected both G-N-7 and 2'-O methylations. GTP and cap analogs did not affect VSV mRNA cap methylation *in vitro*. Overall, the mechanism of VSV mRNA cap methylation is significantly different from that of WNV. Most recently, it was found that capping of flavivi‐ rus RNA is catalyzed by conventional RNA guanylyltransferase via a covalent GMP-en‐ zyme intermediate [88]. However, VSV capping is catalyzed by a novel PRNTase [12, 13, 16, 89]. These studies suggest that VSV and perhaps other NNS RNA viruses have evolved a

unique mechanism to add the cap to their mRNA and to methylate the cap structure.

#### **5.6. VSV methylases require** *cis***-element in RNA**

During mRNA synthesis, the VSV polymerase initiates synthesis at the first gene-start (GS) sequence (3′ UUGUCNNUAC 5′), and the nascent mRNA chain is capped and methylated, and recognizes a specific gene-end (GE) sequence (3′-AUACUUUUUUU-5′), the polymerase polyadenylates and terminates. It has been well demonstrated that the GS sequence contains a key *cis*-acting regulatory element for the initiation of mRNA synthesis [31, 32]. Specifically, the first three positions of the GS sequence have been found to be critical for mRNA synthe‐ sis. Recently, both *trans* capping assays with 5-nt oligo RNA substrates and detergent-acti‐ vated virus transcription reactions pointed out the importance of positions 1, 2, 3, and 5 in mRNA cap addition, although position 5 substitutions were more tolerated [12, 31, 32]. Us‐ ing a *trans* methylation assay, it was found that similar signals were required for mRNA cap methylation [18]. As expected, VSV L protein efficiently methylated a 110 nt of RNA with an authentic gene start sequence at position 2'-O. However, when the gene start sequence of this 110 nt was replaced with non-viral sequence (5′ GpppGGACGAAGAC-RNA), the effi‐ ciency of 2'-O methylation was reduced approximately 9 times. Similarly, VSV L protein ef‐ ficiently methylated a pre-2′-O-methylated VSV mRNA at position G-N-7. In contrast, the efficiency of G-N-7 methylation decreased nearly 7 times when incubated with a substrate with non-VSV mRNA start RNA. Therefore, the gene start sequence of VSV mRNA contains a signal for initiation of mRNA synthesis, mRNA cap addition, and cap methylation.

#### **5.7. The length of mRNA in cap methylation**

In the *trans* mRNA capping assay, the VSV L protein efficiently caps the 5-nt gene start se‐ quence [12, 16], demonstrating that a 5-nt RNA substrate is sufficient for mRNA cap addi‐ tion. In order to determine the minimum length of RNA required for mRNA cap methylation, 5-,10-, 51-, and 110-nt RNAs were used as substrates for a *trans* methylation as‐ say *in vitro* [18]. Interestingly, the 10-, 51-, and 110-nt RNAs were able to serve as substrates for both G-N-7 and ribose 2'-O methylations, whereas the 5-nt RNA was not methylated by the VSV L protein at either the G-N-7 or the ribose 2'-O position [18]. Therefore, in contrast to *trans* capping, a 5-nt substrate is not sufficient for trans methylation and likely the con‐ served positions 8, 9, and 10 in VSV gene start sequence are required for mRNA cap methyl‐ ation. Clearly, the length of RNA required for methylation is longer than that required for capping by the VSV L protein.

#### **5.8. Model for mRNA cap methylation**

dence that 2′-O methylation precedes and facilitates G-N-7 methylation. In light that both two MTase activities appear to reside in the same domain of L protein with the same SAM binding site, it is conceivable that mRNA cap and/or the SAM binding site might need to be repositioned at the end of the first methylation reaction to facilitate the second round of methylation. How this coordination happens *in vitro* is still a mystery. Bearing in mind that G-N-7 position is upstream of the ribose 2'-O position in the mRNA strand, reorientation is thus less likely to have resulted from forward movement of capped RNA through CR VI during transcription, but rather it might entail a fine spatial rearrangement. Collectively, these experiments have shown that the order of VSV mRNA cap methylation is distinct

During mRNA synthesis, the VSV polymerase initiates synthesis at the first gene-start (GS) sequence (3′ UUGUCNNUAC 5′), and the nascent mRNA chain is capped and methylated, and recognizes a specific gene-end (GE) sequence (3′-AUACUUUUUUU-5′), the polymerase polyadenylates and terminates. It has been well demonstrated that the GS sequence contains a key *cis*-acting regulatory element for the initiation of mRNA synthesis [31, 32]. Specifically, the first three positions of the GS sequence have been found to be critical for mRNA synthe‐ sis. Recently, both *trans* capping assays with 5-nt oligo RNA substrates and detergent-acti‐ vated virus transcription reactions pointed out the importance of positions 1, 2, 3, and 5 in mRNA cap addition, although position 5 substitutions were more tolerated [12, 31, 32]. Us‐ ing a *trans* methylation assay, it was found that similar signals were required for mRNA cap methylation [18]. As expected, VSV L protein efficiently methylated a 110 nt of RNA with an authentic gene start sequence at position 2'-O. However, when the gene start sequence of this 110 nt was replaced with non-viral sequence (5′ GpppGGACGAAGAC-RNA), the effi‐ ciency of 2'-O methylation was reduced approximately 9 times. Similarly, VSV L protein ef‐ ficiently methylated a pre-2′-O-methylated VSV mRNA at position G-N-7. In contrast, the efficiency of G-N-7 methylation decreased nearly 7 times when incubated with a substrate with non-VSV mRNA start RNA. Therefore, the gene start sequence of VSV mRNA contains

a signal for initiation of mRNA synthesis, mRNA cap addition, and cap methylation.

In the *trans* mRNA capping assay, the VSV L protein efficiently caps the 5-nt gene start se‐ quence [12, 16], demonstrating that a 5-nt RNA substrate is sufficient for mRNA cap addi‐ tion. In order to determine the minimum length of RNA required for mRNA cap methylation, 5-,10-, 51-, and 110-nt RNAs were used as substrates for a *trans* methylation as‐ say *in vitro* [18]. Interestingly, the 10-, 51-, and 110-nt RNAs were able to serve as substrates for both G-N-7 and ribose 2'-O methylations, whereas the 5-nt RNA was not methylated by the VSV L protein at either the G-N-7 or the ribose 2'-O position [18]. Therefore, in contrast to *trans* capping, a 5-nt substrate is not sufficient for trans methylation and likely the con‐ served positions 8, 9, and 10 in VSV gene start sequence are required for mRNA cap methyl‐ ation. Clearly, the length of RNA required for methylation is longer than that required for

from all other known mRNA cap methylation mechanisms.

**5.6. VSV methylases require** *cis***-element in RNA**

250 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**5.7. The length of mRNA in cap methylation**

capping by the VSV L protein.

The process of VSV L protein-mediated cap methylation can be best summarized with the following model (Fig.7). Initially in response to a specific *cis-*acting element in the VSV gene start sequence, CR VI of L protein methylates the cap structure first at the 2'-O position to produce GpppAmACAG-RNA. The by-product of this reaction, SAH, is released from this reaction. Following 2'-O methylation, a second molecule of SAM binds to CR VI of L protein that may facilitate a subsequent methylation of the RNA at the G-N-7 position. Methylation at the 2'-O position favors G-N-7 methylation in the cap structure through a currently un‐ known mechanism. G-N-7 methlyation may be facilitated by the contact with the RNA mol‐ ecule remains bound to the L protein at the end of the initial methylation at the 2'-O position. Another possibility is that CR VI of L protein exists in two different conformations. An initial conformation may favor binding of GpppRNA and SAM. Methylation of RNA at the 2'-O position perhaps induces a conformational change that facilitates the repositioning of the RNA for subsequent G-N-7 methylation, and/or favors the release of SAH as well as the binding of a subsequent molecule of SAM.

#### **5.9. Comparison of mRNA cap methylation in VSV and WNV**

To date, the rhabdovirus, VSV, and the flavivirus, WNV, are the two best characterized vi‐ ruses that utilize a single region in the polymerase protein for both G-N-7 and 2'-O methyla‐ tions. However, the mechanism of VSV mRNA methylation is distinct from that of the WNV system (Fig.4B and C). In VSV, 2'-O methylation precedes and facilitates subsequent G-N-7 methylation [17, 18]. However, WNV MTases modify the cap structure, first at the G-N-7 position and subsequently at the ribose 2'-O position [72, 73, 78]. In VSV, the G-N-7 and 2'-O MTases require similar conditions for methylation with an optimal pH at 7.0 [18]. In con‐ trast, the G-N-7 and 2'-O MTases of WNV require an optimal pH at 6.5 and 10, respectively [72]. Both VSV and WNV MTases modify the RNA in a sequence-specific manner, but re‐ quire different elements in the RNA substrate. VSV G-N-7 and 2'-O MTases require specific gene start sequences with a minimum mRNA length of 10 nucleotides [18]. In the WNV model, N-7 cap methylation requires the presence of specific nucleotides at the second and third positions and a 5' stem-loop structure within the 74-nucleotide viral RNA; in contrast, 2'-O ribose methylation requires specific nucleotides at the first and second positions, with a minimum 5' viral RNA of 20 nucleotides in length [81]. In addition, there is striking differ‐ ence in the cap recognition site between the VSV and WNV MTases. For the WNV MTase, the cap recognition site is essential for 2'-O, but not G-N-7 methylation [73, 82]. Consistent with this finding, it was found that GTP and cap analogs specifically inhibited 2'-O, but not G-N-7 methylation [73, 82]. However, mutations to the putative RNA binding site in VSV L protein affected both G-N-7 and 2'-O methylations. GTP and cap analogs did not affect VSV mRNA cap methylation *in vitro*. Overall, the mechanism of VSV mRNA cap methylation is significantly different from that of WNV. Most recently, it was found that capping of flavivi‐ rus RNA is catalyzed by conventional RNA guanylyltransferase via a covalent GMP-en‐ zyme intermediate [88]. However, VSV capping is catalyzed by a novel PRNTase [12, 13, 16, 89]. These studies suggest that VSV and perhaps other NNS RNA viruses have evolved a unique mechanism to add the cap to their mRNA and to methylate the cap structure.

G-N-7 position [43]. However, the "*trans*-methylation" assay used in their study does not allow detection of 2'-O methylation although it is known that Sendai virus encodes two MTases. More recently, recombinant Sendai virus carrying mutations in catalytic KDKE tetrad and SAM binding site were recovered from infectious clones [91, 92]. It was found that these muta‐ tions affected mRNA cap methylation. However, whether they are specifically defective in G-N-7 and/or 2'-O methylation is not known because of the assay did not have the ability to distinguish the two methyl groups. In addition, the order of methylation may be different among NNS RNA viruses. In contrast to VSV, methylation of RSV mRNA at low SAM concen‐ trations was found at only the G-N-7 position; however, it was found to be doubly methylated at high SAM concentrations [63]. This study provided evidence that the order of mRNA cap methylation in RSV may be different with VSV. Clearly, more studies are needed to under‐

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**Figure 8. SAM binding motifs in NNS RNA virus MTases.** The proposed SAM binding region for the *Filoviridae*, Ru‐ *bulavirus*, and *Avulavirus* genera of the *Paramyxoviridae* contains a conserved AxGxG sequence rather than GxGxG. NNS RNA viruses include VSVI, VSV Indiana; VSVN, VSV New Jersey; BEFV, bovine emphemeral fever virus; RABV, rabies virus; Marb, Marburg virus; MeV: measles; HV, Hendra virus; NPV, Nipah virus; AMPV, avian metapneuomovirus; HMPV, human metapneuomovirus; APMV6, avian paramyxovirus 6; MuV, mumps virus; HPIV2, human parainfluenza

SS(+) RNA: single strand positive RNA viruses include DNV, dengue virus; WNV, West

stand the mRNA cap methylation in other NNS RNA viruses.

virus 2; SV5, simian virus 5; SV41, simian virus 41;

Nile virus; YFV, yellow fever virus.

**Figure 7. Proposed model for mRNA cap methylation in VSV and eukaryotic cells. (A) VSV MTase model.** VSV mRNA cap structure is first methylated by CR VI of L protein at the 2'-O position to produce GpppAmACAG-RNA. The by-product, SAH, is released before the binding of a second molecule of SAM. G-N-7 methlyation may be facilitated by the contact with the RNA molecule remains bound to the polymerase at the end of the initial methylation at the 2'-O position. Or, methylation of RNA at the 2'-O position may induce a conformational change that facilitates the reposi‐ tioning of the RNA for subsequent G-N-7 methylation. **(B) Cellular MTase model.** GpppNNNN-RNA is first methylated by a G-N-7 MTase to yield 7mGpppNNNN-RNA. Following G-N-7 methylation, 7mGpppNNNN-RNA dissociates with G-N-7 MTase, and re-associates with a separate 2'-O MTase to yield 7mGpppNmNNN-RNA. In this model, the two methyl‐ ase activities are carried out by two separate enzymes, each containing its own SAM binding site.

#### **5.10. mRNA cap methylation in other NNS RNA viruses**

Limited accomplishments have been made in understanding the mechanism of mRNA cap methylation in other NNS RNA viruses, due to the lack of a robust *in vitro* mRNA synthesis for most of NNS RNA viruses, and the technical challenge of expression and purification of a func‐ tional polymerase protein or fragment. In early 1970, Colonno and Stone showed that the NDV mRNA cap structure was methylated only at G-N-7 [90]. This is distinct from the cap structures of other NNS RNA viruses which typically contain two methyl groups, at positions G-N-7 and ribose 2'-O. However, detailed characterization of the NDV methylase activities and the mech‐ anism involved in this unique methylation is not understood. The mechanism underlying this difference and the biological significance of the lack of 2'-O MTase is not known. Sequence analysis has revealed that the proposed SAM binding region for the *Filoviridae*, *Rubulavirus*, and *Avulavirus* genera of the *Paramyxoviridae* contains a conserved AxGxG sequence rather than GxGxG within motif I of the SAM-dependent MTase superfamily (Fig.8). It will be inter‐ esting to determine if there is a link between this differential SAM binding sites and the lack of 2'-O methylation in NDV. Recently, it was shown that a fragment of Sendai virus L protein that includes CR VI was able to methylate short Sendai virus-specific RNA sequences *in trans* at the G-N-7 position [43]. However, the "*trans*-methylation" assay used in their study does not allow detection of 2'-O methylation although it is known that Sendai virus encodes two MTases. More recently, recombinant Sendai virus carrying mutations in catalytic KDKE tetrad and SAM binding site were recovered from infectious clones [91, 92]. It was found that these muta‐ tions affected mRNA cap methylation. However, whether they are specifically defective in G-N-7 and/or 2'-O methylation is not known because of the assay did not have the ability to distinguish the two methyl groups. In addition, the order of methylation may be different among NNS RNA viruses. In contrast to VSV, methylation of RSV mRNA at low SAM concen‐ trations was found at only the G-N-7 position; however, it was found to be doubly methylated at high SAM concentrations [63]. This study provided evidence that the order of mRNA cap methylation in RSV may be different with VSV. Clearly, more studies are needed to under‐ stand the mRNA cap methylation in other NNS RNA viruses.

**Figure 7. Proposed model for mRNA cap methylation in VSV and eukaryotic cells. (A) VSV MTase model.** VSV mRNA cap structure is first methylated by CR VI of L protein at the 2'-O position to produce GpppAmACAG-RNA. The by-product, SAH, is released before the binding of a second molecule of SAM. G-N-7 methlyation may be facilitated by the contact with the RNA molecule remains bound to the polymerase at the end of the initial methylation at the 2'-O position. Or, methylation of RNA at the 2'-O position may induce a conformational change that facilitates the reposi‐ tioning of the RNA for subsequent G-N-7 methylation. **(B) Cellular MTase model.** GpppNNNN-RNA is first methylated by a G-N-7 MTase to yield 7mGpppNNNN-RNA. Following G-N-7 methylation, 7mGpppNNNN-RNA dissociates with G-N-7 MTase, and re-associates with a separate 2'-O MTase to yield 7mGpppNmNNN-RNA. In this model, the two methyl‐

Limited accomplishments have been made in understanding the mechanism of mRNA cap methylation in other NNS RNA viruses, due to the lack of a robust *in vitro* mRNA synthesis for most of NNS RNA viruses, and the technical challenge of expression and purification of a func‐ tional polymerase protein or fragment. In early 1970, Colonno and Stone showed that the NDV mRNA cap structure was methylated only at G-N-7 [90]. This is distinct from the cap structures of other NNS RNA viruses which typically contain two methyl groups, at positions G-N-7 and ribose 2'-O. However, detailed characterization of the NDV methylase activities and the mech‐ anism involved in this unique methylation is not understood. The mechanism underlying this difference and the biological significance of the lack of 2'-O MTase is not known. Sequence analysis has revealed that the proposed SAM binding region for the *Filoviridae*, *Rubulavirus*, and *Avulavirus* genera of the *Paramyxoviridae* contains a conserved AxGxG sequence rather than GxGxG within motif I of the SAM-dependent MTase superfamily (Fig.8). It will be inter‐ esting to determine if there is a link between this differential SAM binding sites and the lack of 2'-O methylation in NDV. Recently, it was shown that a fragment of Sendai virus L protein that includes CR VI was able to methylate short Sendai virus-specific RNA sequences *in trans* at the

ase activities are carried out by two separate enzymes, each containing its own SAM binding site.

**5.10. mRNA cap methylation in other NNS RNA viruses**

252 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 8. SAM binding motifs in NNS RNA virus MTases.** The proposed SAM binding region for the *Filoviridae*, Ru‐ *bulavirus*, and *Avulavirus* genera of the *Paramyxoviridae* contains a conserved AxGxG sequence rather than GxGxG. NNS RNA viruses include VSVI, VSV Indiana; VSVN, VSV New Jersey; BEFV, bovine emphemeral fever virus; RABV, rabies virus; Marb, Marburg virus; MeV: measles; HV, Hendra virus; NPV, Nipah virus; AMPV, avian metapneuomovirus; HMPV, human metapneuomovirus; APMV6, avian paramyxovirus 6; MuV, mumps virus; HPIV2, human parainfluenza virus 2; SV5, simian virus 5; SV41, simian virus 41;

SS(+) RNA: single strand positive RNA viruses include DNV, dengue virus; WNV, West Nile virus; YFV, yellow fever virus.

#### **6. The effects of 5' mRNA cap addition and cap methylation on 3' mRNA polyadenylation**

Perhaps, K 1651A and D1762E substitutions might favor the binding of SAH at the SAM binding site in CR VI, resulting in hyperpolyadenylation without the need for supplemental SAH. Clearly, further studies are needed to understand the relationship between 5' mRNA

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However, it appears clear that 5' cap addition is required for 3' polyadenylation, as evi‐ denced by the polymerase mutants (G1154, T1157, H1227, and R1228) within CR V of L that inhibited cap addition also inhibit polyadenylation [15]. These cap-defective polymerases synthesized truncated transcripts that predominantly terminated within the first 500 nt of the N gene and contained short A-rich sequences at their 3′ termini. To examine how the cap-defective polymerases respond to an authentic VSV termination and re-initiation signal present at each gene junction, a 382 nt gene was inserted at the leader-*N* gene junction in the VSV genome. Using this N-RNA as the template, the cap-defective polymerases were able to synthesize full-length 382-nt transcripts that were not capped at 5'end. Interestingly, these uncapped transcripts lacked an authentic polyadenylate tail and instead contained 0 to 24 A residues [15]. In addition, the cap-defective polymerases were also unable to efficiently copy the downstream genes [15]. This finding strongly supports that 5′ mRNA cap addition and

cap methyaltion and 3' polyadenylation.

**expression**

3′ polyadenylation are mechanistically and functionally linked.

**7. Impact of mRNA cap methylation on viral replication and gene**

In eukaryotic cells, it is well established that G-N-7 methylation of the mRNA cap structure is essential for mRNA stability and efficient translation [11, 48, 66]. Specifically, G-N-7 meth‐ ylation of the mRNA cap structure is required for recognition of the cap by the rate limiting factor for translation initiation, eIF-4E [100, 101]. The mRNA cap structures of NNS RNA vi‐ ruses are typically G-N-7 and 2'-O methylated. Although the precise mechanism by which VSV mRNAs are translated is unclear, they are broadly thought to utilize a variation of the canonical cap-dependent translational pathway [102-104]. In VSV-infected cells, host mRNA translation is rapidly inhibited through the suppression of the intracellular pools of eIF-4E by a manipulation of the phosphorylation status of the 4E binding protein (4E-BP1) [104]. Nevertheless, *in vitro* experiments have shown that G-N-7 cap methylation facilitates trans‐ lation of VSV proteins. MTase-defective VSV would provide a tool to study the role of mRNA cap methylation in viral protein synthesis. Ultimately, it will affect viral genome rep‐ lication and gene expression since viral replication requires ongoing protein synthesis.

Based on the status of mRNA methylation, MTase-defective VSVs can be classified into three groups [14, 17, 84]. Viruses in the first group are completely defective in both G-N-7 and 2'-O methylation, including mutations in MTase active site (rVSV-K1651A, D1762A, K1795A, E1833Q, and E1833A), SAM binding site (rVSV-D1671V, G1675A, G4A, and G4AD), and putative RNA binding site (rVSV-Y1650A, F1691A, and E1764A). Viruses in the second group are specifically defective in G-N-7, but not 2'-O MTase, including mutants in SAM binding site (rVSV-G1670A, G1672A, and S1673A). Viruses in third group that require

VSV mRNA is capped and methylated at the 5′ end and polyadenylated at the 3′ end. Cap addition, cap methylation, and polyadenylation are carried out by three different re‐ gions (CR V, CR VI, and CR III) in L protein. During VSV mRNA synthesis, modifica‐ tions of the 5′ and 3′ ends of the mRNAs are tightly coupled to transcription [21-23]. Although the detailed mechanism by which the polymerase coordinates these modifica‐ tion events is poorly understand, available evidence suggests there is a link between au‐ thentic 5′-end formation and 3′-end formation during VSV mRNA synthesis. Early studies demonstrated that the length of poly (A) tails on VSV mRNAs is affected by the presence of SAH, the by-product and competitive inhibitor of SAM-mediated methyl‐ transferases [93-95]. The fact that the polymerase can synthesize full-length mRNAs *in vi‐ tro* in the absence of SAM or in the presence of SAH, suggests that transcription is not dependent on cap methylation. When *in vitro* transcription reactions are performed in the presence of SAM, the RdRp synthesizes mRNA with poly (A) tail of 100 to 200 nt in length, similar to those synthesized in VSV-infected cells. However, when VSV mRNA cap methylation was inhibited during *in vitro* transcription reactions by supplementing with 1 mM SAH, the synthesized mRNA was heterogeneous in length due to having ex‐ tremely long poly (A) tails, from 700 to 2,400 nucleotides (nt) in length [93, 96]. Recent work demonstrated that SAH-induced hyperpolyadenylation also occurs in cells infected by wild-type VSV in the presence of adenosine dialdehyde (AdOX), a compound that in‐ hibits the activity of SAH hydrolase [97]. These results indicated that chemical inhibition of VSV mRNA cap methylation by SAH resulted in hyperpolyadenylation of viral mRNA. Interestingly, this hyperpolyadenylation of VSV mRNAs has been observed in *ts*(G)16, a VSV mutant identified in 1970 based on its ability to grow at 31°C but not at 39°C [93, 98, 99]. In the absence of SAH, the mRNAs synthesized by VSV mutant *ts*(G)16 were hyperpolyadenylated at the 3′ end. Genomic sequence analysis found that the L protein of *ts*(G)16 contains two amino acid changes, C1291Y and F1488S, compared to wild type. Combined with the analysis of revertants of *ts*(G)16, it was found that F1488S, located in the variable region of L between CR V and CR VI, is responsible for the hy‐ perpolyadenylating phenotype.

The characterization of a panel of MTase-defective VSVs may serve as a tool to understand the mechanism by which SAH or the failure to methylate the cap structure results in hyper‐ polyadenylation. It was found that rVSV-K1651A, a mutation in MTase active site and com‐ pletely defective in G-N-7 and 2'-O methylation, synthesized excessively long poly(A) tails, similar to those produced by wild-type L in the presence of SAH [15]. Similarly, the substi‐ tution D1762E at the MTase active site, which inhibits both G-N-7 and 2'-O methylation, produces large polyadenylate in the presence or absence of SAH [97]. This data confirms the earlier work demonstrating that the inhibition of cap methylation results in large polyade‐ nylate. In contrast, several other substitutions that inhibit cap methylation, including D1762G, D1762N, G1672P, and G1675P, did not produce hyperpolyadenylated mRNA [97]. Perhaps, K 1651A and D1762E substitutions might favor the binding of SAH at the SAM binding site in CR VI, resulting in hyperpolyadenylation without the need for supplemental SAH. Clearly, further studies are needed to understand the relationship between 5' mRNA cap methyaltion and 3' polyadenylation.

**6. The effects of 5' mRNA cap addition and cap methylation on 3' mRNA**

VSV mRNA is capped and methylated at the 5′ end and polyadenylated at the 3′ end. Cap addition, cap methylation, and polyadenylation are carried out by three different re‐ gions (CR V, CR VI, and CR III) in L protein. During VSV mRNA synthesis, modifica‐ tions of the 5′ and 3′ ends of the mRNAs are tightly coupled to transcription [21-23]. Although the detailed mechanism by which the polymerase coordinates these modifica‐ tion events is poorly understand, available evidence suggests there is a link between au‐ thentic 5′-end formation and 3′-end formation during VSV mRNA synthesis. Early studies demonstrated that the length of poly (A) tails on VSV mRNAs is affected by the presence of SAH, the by-product and competitive inhibitor of SAM-mediated methyl‐ transferases [93-95]. The fact that the polymerase can synthesize full-length mRNAs *in vi‐ tro* in the absence of SAM or in the presence of SAH, suggests that transcription is not dependent on cap methylation. When *in vitro* transcription reactions are performed in the presence of SAM, the RdRp synthesizes mRNA with poly (A) tail of 100 to 200 nt in length, similar to those synthesized in VSV-infected cells. However, when VSV mRNA cap methylation was inhibited during *in vitro* transcription reactions by supplementing with 1 mM SAH, the synthesized mRNA was heterogeneous in length due to having ex‐ tremely long poly (A) tails, from 700 to 2,400 nucleotides (nt) in length [93, 96]. Recent work demonstrated that SAH-induced hyperpolyadenylation also occurs in cells infected by wild-type VSV in the presence of adenosine dialdehyde (AdOX), a compound that in‐ hibits the activity of SAH hydrolase [97]. These results indicated that chemical inhibition of VSV mRNA cap methylation by SAH resulted in hyperpolyadenylation of viral mRNA. Interestingly, this hyperpolyadenylation of VSV mRNAs has been observed in *ts*(G)16, a VSV mutant identified in 1970 based on its ability to grow at 31°C but not at 39°C [93, 98, 99]. In the absence of SAH, the mRNAs synthesized by VSV mutant *ts*(G)16 were hyperpolyadenylated at the 3′ end. Genomic sequence analysis found that the L protein of *ts*(G)16 contains two amino acid changes, C1291Y and F1488S, compared to wild type. Combined with the analysis of revertants of *ts*(G)16, it was found that F1488S, located in the variable region of L between CR V and CR VI, is responsible for the hy‐

The characterization of a panel of MTase-defective VSVs may serve as a tool to understand the mechanism by which SAH or the failure to methylate the cap structure results in hyper‐ polyadenylation. It was found that rVSV-K1651A, a mutation in MTase active site and com‐ pletely defective in G-N-7 and 2'-O methylation, synthesized excessively long poly(A) tails, similar to those produced by wild-type L in the presence of SAH [15]. Similarly, the substi‐ tution D1762E at the MTase active site, which inhibits both G-N-7 and 2'-O methylation, produces large polyadenylate in the presence or absence of SAH [97]. This data confirms the earlier work demonstrating that the inhibition of cap methylation results in large polyade‐ nylate. In contrast, several other substitutions that inhibit cap methylation, including D1762G, D1762N, G1672P, and G1675P, did not produce hyperpolyadenylated mRNA [97].

**polyadenylation**

254 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

perpolyadenylating phenotype.

However, it appears clear that 5' cap addition is required for 3' polyadenylation, as evi‐ denced by the polymerase mutants (G1154, T1157, H1227, and R1228) within CR V of L that inhibited cap addition also inhibit polyadenylation [15]. These cap-defective polymerases synthesized truncated transcripts that predominantly terminated within the first 500 nt of the N gene and contained short A-rich sequences at their 3′ termini. To examine how the cap-defective polymerases respond to an authentic VSV termination and re-initiation signal present at each gene junction, a 382 nt gene was inserted at the leader-*N* gene junction in the VSV genome. Using this N-RNA as the template, the cap-defective polymerases were able to synthesize full-length 382-nt transcripts that were not capped at 5'end. Interestingly, these uncapped transcripts lacked an authentic polyadenylate tail and instead contained 0 to 24 A residues [15]. In addition, the cap-defective polymerases were also unable to efficiently copy the downstream genes [15]. This finding strongly supports that 5′ mRNA cap addition and 3′ polyadenylation are mechanistically and functionally linked.

### **7. Impact of mRNA cap methylation on viral replication and gene expression**

In eukaryotic cells, it is well established that G-N-7 methylation of the mRNA cap structure is essential for mRNA stability and efficient translation [11, 48, 66]. Specifically, G-N-7 meth‐ ylation of the mRNA cap structure is required for recognition of the cap by the rate limiting factor for translation initiation, eIF-4E [100, 101]. The mRNA cap structures of NNS RNA vi‐ ruses are typically G-N-7 and 2'-O methylated. Although the precise mechanism by which VSV mRNAs are translated is unclear, they are broadly thought to utilize a variation of the canonical cap-dependent translational pathway [102-104]. In VSV-infected cells, host mRNA translation is rapidly inhibited through the suppression of the intracellular pools of eIF-4E by a manipulation of the phosphorylation status of the 4E binding protein (4E-BP1) [104]. Nevertheless, *in vitro* experiments have shown that G-N-7 cap methylation facilitates trans‐ lation of VSV proteins. MTase-defective VSV would provide a tool to study the role of mRNA cap methylation in viral protein synthesis. Ultimately, it will affect viral genome rep‐ lication and gene expression since viral replication requires ongoing protein synthesis.

Based on the status of mRNA methylation, MTase-defective VSVs can be classified into three groups [14, 17, 84]. Viruses in the first group are completely defective in both G-N-7 and 2'-O methylation, including mutations in MTase active site (rVSV-K1651A, D1762A, K1795A, E1833Q, and E1833A), SAM binding site (rVSV-D1671V, G1675A, G4A, and G4AD), and putative RNA binding site (rVSV-Y1650A, F1691A, and E1764A). Viruses in the second group are specifically defective in G-N-7, but not 2'-O MTase, including mutants in SAM binding site (rVSV-G1670A, G1672A, and S1673A). Viruses in third group that require elevated SAM concentrations to permit full methylation including a mutant in SAM binding site (rVSV-G1674A) and putative RNA binding site (rVSV-Y1835A). With the exception of rVSV-G1674A and Y1835A, all MTase-defective VSVs were attenuated in cell culture as judged by diminished viral plaque size, reduced infectious viral progeny release (in singlestep growth curves), and decreased levels of viral genomic RNA, mRNA, and protein syn‐ thesis. It appears that the degree of attenuation is consistent with the defects of the methylation. For example, viruses defective in both G-N-7 and 2'-O methylation had 2-5 log reductions in growth whereas viruses only defective in G-N-7 had 1-2 log declines in repli‐ cation [14]. Recombinant rVSV-G1674A and Y1835A replicated as efficiently as wild type rVSV [14]. A remarkable finding is that some of the mutants in the SAM binding site (rVSV-G1675A, G4A, and G4AD) affected transcription and replication differently [17]. For these mutants, replication was enhanced 2.5- to 4-fold, and transcription decreased up to 8-fold compared with rVSV. One feature of the gene expression strategy of NNS RNA viruses is that the polymerase complex controls two distinct RNA synthetic events: genomic RNA rep‐ lication and mRNA transcription [20, 105, 106]. It is possible that SAM binding influences the switch of polymerase between replicase and transcriptase. Perhaps, L protein with SAM binding favors to function as transcriptase, whereas L protein that lacks SAM binding favors replicase function.

days but recovered quickly. But, none of mice in rVSV-G1670A and G1672A had neurologi‐ cal symptoms. These two recombinants had moderate defects in replication in lungs and brain, and caused moderate histopathological changes in lungs. Interestingly, recombinant rVSV-G4A, which carries four mutations (G1670A, G1672A, G1674A, and G1675A) in SAM binding site and is completely defective in both G-N-7 and 2'-O methylation, exhibited an interesting pathotype [107]. Similar to VSV mutants at the MTase catalytic site, rVSV-G4A had dramatically impaired viral replication in the lungs and brain of the mice. However, the rVSV-G4A group exhibited body weight losses that were comparable to G1670A and G1672A, and clinical signs that were more severe than G1670A and G1672A. These results suggest rVSV-G4A retained low to moderate virulence despite the fact that it was attenuat‐ ed for replication and or spread *in vitro*. As predicted, recombinant rVSV-G1674A, which contains a point mutation in the SAM binding site and requires elevated SAM concentra‐ tions to permit full methylation, was highly virulent to mice. These results suggest that (i) the relationship between mRNA cap methylation and viral pathogenesis is not clear cut in VSV infections; and (ii) inactivation of the predicted catalytic residues attenuates the virus to

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a greater extent *in vitro*, than does inactivation of the predicted SAM binding site.

While it is firmly established that G-N-7 methylation is essential for mRNA stability as well as efficient translation, the role(s) of ribose 2'-O methylation have proven more elusive. Re‐ cent studies on West Nile virus (WNV) suggest that the 2′-O methylation of the 5′ cap of viral RNA functions to evade innate host antiviral responses through escape of the suppres‐ sion of interferon-stimulated genes, tetratricopeptide repeats (IFIT)[111]. Specifically, mu‐ tant WNV (E218A) defective in 2′-O MTase activity was attenuated in wild-type C57BL/6 mice, but remained pathogenic in knockout mice that lacked the type I interferon (IFN) sig‐ naling pathway. In addition, a vaccinia virus mutant (J3-K175R) and mouse hepatitis virus (MHV) mutant D130A, both of which lacked 2′-O MTase activity, exhibited enhanced sensi‐ tivities to the antiviral actions of IFN mediated by IFIT proteins. Interestingly, it was also reported that 2'-O methylation of mouse and human coronavirus RNA facilitates evasion from detection by the cytoplasmic RNA sensor Mda5 [112]. Taken together, these studies suggest that 2'-O methylation of viral RNA provides a molecular signature for the discrimi‐ nation of self and non-self mRNA. It is known that mRNAs of most NNS RNA viruses con‐ tain G-N-7 and 2'-O methylation. However, whether 2'-O methylation plays a similar role in

VSV is an excellent model to aid in the understanding the role of viral mRNA cap methyla‐ tion in innate immunity. The mechanism of VSV mRNA cap methylation is unique in that 2'-O methylation precedes and facilitates the G-N-7 methylation [17, 18]. It is unknown why the order of VSV mRNA methylation is reversed compared to all known mRNA cap methyl‐ ation reactions. One possibility is that the methylation of 2'-O allows VSV to successfully mimic cellular mRNA to avoid the detection by host innate immunity, which in turn pro‐ motes efficient viral replication in hosts. In fact, VSV is one of only a few viruses that repli‐

**9. 2'-O methylation and innate immunity**

all NNS RNA viruses is not known.

#### **8. Impact of mRNA cap methylation on viral pathogenesis** *in vitro*

Although it is well studied that MTase-defective viruses were attenuated in cell culture, the impact of mRNA cap methylation on viral pathogenesis *in vitro* is poorly understood. Re‐ cently, Ma et al., (2012) examined the pathogenicity of MTase-defective VSVs in mice [107]. VSV infects a wide range of wild and domestic animals such as cattle, horses, deer, and pigs, characterized by vesicular lesions in the mouth, tongue, lips, gums, teats, and feet. Although the mouse is not the natural host of VSV, it represents an excellent small animal model to understand VSV pathogenesis because VSV causes systemic infection and fatal encephalitis [108-110]. After intranasal inoculation, VSV infects olfactory neurons in the nasal mucosa and subsequently enters the central nervous system (CNS) through the olfactory nerves. The virus is then disseminated to other areas in the brain through retrograde and possibly ante‐ rograde trans-neuronal transport, ultimately causing an acute brain infection. It was found that VSV mutants, rVSV-K1651A, D1762A, and E1833Q, which have mutations in the MTase catalytic site and are defective in both G-N-7 and 2'O methylation, were highly attenuated in mice [107]. Mice inoculated with these recombinant viruses did not show any clinical signs of VSV infection such as weight loss, ruffled fur, hyperexcitability, tremors, circling, and pa‐ ralysis. Furthermore, these mutant viruses were not able to enter the brain, had dramatic de‐ fects in replication in lungs, and did not cause significant histopathological changes in lungs and brain. Recombinant rVSV-G1670A and G1672A, which have mutations in the SAM binding site and are defective in G-N-7 but not 2'-O methylation, retained low virulence in mice [107]. Mice inoculated these two recombinants exhibited weight loss of approximately 2-3 g during days 3-7 post-inoculation and showed mild illnesses such as ruffled coat for 2-3 days but recovered quickly. But, none of mice in rVSV-G1670A and G1672A had neurologi‐ cal symptoms. These two recombinants had moderate defects in replication in lungs and brain, and caused moderate histopathological changes in lungs. Interestingly, recombinant rVSV-G4A, which carries four mutations (G1670A, G1672A, G1674A, and G1675A) in SAM binding site and is completely defective in both G-N-7 and 2'-O methylation, exhibited an interesting pathotype [107]. Similar to VSV mutants at the MTase catalytic site, rVSV-G4A had dramatically impaired viral replication in the lungs and brain of the mice. However, the rVSV-G4A group exhibited body weight losses that were comparable to G1670A and G1672A, and clinical signs that were more severe than G1670A and G1672A. These results suggest rVSV-G4A retained low to moderate virulence despite the fact that it was attenuat‐ ed for replication and or spread *in vitro*. As predicted, recombinant rVSV-G1674A, which contains a point mutation in the SAM binding site and requires elevated SAM concentra‐ tions to permit full methylation, was highly virulent to mice. These results suggest that (i) the relationship between mRNA cap methylation and viral pathogenesis is not clear cut in VSV infections; and (ii) inactivation of the predicted catalytic residues attenuates the virus to a greater extent *in vitro*, than does inactivation of the predicted SAM binding site.

#### **9. 2'-O methylation and innate immunity**

elevated SAM concentrations to permit full methylation including a mutant in SAM binding site (rVSV-G1674A) and putative RNA binding site (rVSV-Y1835A). With the exception of rVSV-G1674A and Y1835A, all MTase-defective VSVs were attenuated in cell culture as judged by diminished viral plaque size, reduced infectious viral progeny release (in singlestep growth curves), and decreased levels of viral genomic RNA, mRNA, and protein syn‐ thesis. It appears that the degree of attenuation is consistent with the defects of the methylation. For example, viruses defective in both G-N-7 and 2'-O methylation had 2-5 log reductions in growth whereas viruses only defective in G-N-7 had 1-2 log declines in repli‐ cation [14]. Recombinant rVSV-G1674A and Y1835A replicated as efficiently as wild type rVSV [14]. A remarkable finding is that some of the mutants in the SAM binding site (rVSV-G1675A, G4A, and G4AD) affected transcription and replication differently [17]. For these mutants, replication was enhanced 2.5- to 4-fold, and transcription decreased up to 8-fold compared with rVSV. One feature of the gene expression strategy of NNS RNA viruses is that the polymerase complex controls two distinct RNA synthetic events: genomic RNA rep‐ lication and mRNA transcription [20, 105, 106]. It is possible that SAM binding influences the switch of polymerase between replicase and transcriptase. Perhaps, L protein with SAM binding favors to function as transcriptase, whereas L protein that lacks SAM binding favors

256 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**8. Impact of mRNA cap methylation on viral pathogenesis** *in vitro*

Although it is well studied that MTase-defective viruses were attenuated in cell culture, the impact of mRNA cap methylation on viral pathogenesis *in vitro* is poorly understood. Re‐ cently, Ma et al., (2012) examined the pathogenicity of MTase-defective VSVs in mice [107]. VSV infects a wide range of wild and domestic animals such as cattle, horses, deer, and pigs, characterized by vesicular lesions in the mouth, tongue, lips, gums, teats, and feet. Although the mouse is not the natural host of VSV, it represents an excellent small animal model to understand VSV pathogenesis because VSV causes systemic infection and fatal encephalitis [108-110]. After intranasal inoculation, VSV infects olfactory neurons in the nasal mucosa and subsequently enters the central nervous system (CNS) through the olfactory nerves. The virus is then disseminated to other areas in the brain through retrograde and possibly ante‐ rograde trans-neuronal transport, ultimately causing an acute brain infection. It was found that VSV mutants, rVSV-K1651A, D1762A, and E1833Q, which have mutations in the MTase catalytic site and are defective in both G-N-7 and 2'O methylation, were highly attenuated in mice [107]. Mice inoculated with these recombinant viruses did not show any clinical signs of VSV infection such as weight loss, ruffled fur, hyperexcitability, tremors, circling, and pa‐ ralysis. Furthermore, these mutant viruses were not able to enter the brain, had dramatic de‐ fects in replication in lungs, and did not cause significant histopathological changes in lungs and brain. Recombinant rVSV-G1670A and G1672A, which have mutations in the SAM binding site and are defective in G-N-7 but not 2'-O methylation, retained low virulence in mice [107]. Mice inoculated these two recombinants exhibited weight loss of approximately 2-3 g during days 3-7 post-inoculation and showed mild illnesses such as ruffled coat for 2-3

replicase function.

While it is firmly established that G-N-7 methylation is essential for mRNA stability as well as efficient translation, the role(s) of ribose 2'-O methylation have proven more elusive. Re‐ cent studies on West Nile virus (WNV) suggest that the 2′-O methylation of the 5′ cap of viral RNA functions to evade innate host antiviral responses through escape of the suppres‐ sion of interferon-stimulated genes, tetratricopeptide repeats (IFIT)[111]. Specifically, mu‐ tant WNV (E218A) defective in 2′-O MTase activity was attenuated in wild-type C57BL/6 mice, but remained pathogenic in knockout mice that lacked the type I interferon (IFN) sig‐ naling pathway. In addition, a vaccinia virus mutant (J3-K175R) and mouse hepatitis virus (MHV) mutant D130A, both of which lacked 2′-O MTase activity, exhibited enhanced sensi‐ tivities to the antiviral actions of IFN mediated by IFIT proteins. Interestingly, it was also reported that 2'-O methylation of mouse and human coronavirus RNA facilitates evasion from detection by the cytoplasmic RNA sensor Mda5 [112]. Taken together, these studies suggest that 2'-O methylation of viral RNA provides a molecular signature for the discrimi‐ nation of self and non-self mRNA. It is known that mRNAs of most NNS RNA viruses con‐ tain G-N-7 and 2'-O methylation. However, whether 2'-O methylation plays a similar role in all NNS RNA viruses is not known.

VSV is an excellent model to aid in the understanding the role of viral mRNA cap methyla‐ tion in innate immunity. The mechanism of VSV mRNA cap methylation is unique in that 2'-O methylation precedes and facilitates the G-N-7 methylation [17, 18]. It is unknown why the order of VSV mRNA methylation is reversed compared to all known mRNA cap methyl‐ ation reactions. One possibility is that the methylation of 2'-O allows VSV to successfully mimic cellular mRNA to avoid the detection by host innate immunity, which in turn pro‐ motes efficient viral replication in hosts. In fact, VSV is one of only a few viruses that repli‐ cates efficiently in a wide range of cell lines including mammalian cells, insect cells, and worms [113]. Unlike WNV, VSV that is specifically defective in G-N-7 methylation can also be successfully recovered [17]. VSV mutants defective in G-N-7 methylation or both G-N-7 and 2'-O methylations can serve as prototypes or controls to elucidate the role of methyla‐ tion in innate immunity. In addition, the VSV mutant (rVSV-G4A) that was attenuated in cell culture retained low to moderate virulence, suggesting the possible role of methylation in averting the innate immune response [107]. Notably, mRNAs of NDV, an avian paramyx‐ ovirus, are not 2'-O methylated [90]. A direct comparison to determine whether 2'-O meth‐ ylation of VSV has a similar biological function compared to WNV and MHV should prove very compelling.

viruses. It was reported that recombinant Sendai virus carrying point mutations in the MTase catalytic site (rSeV-K1782A) and the SAM binding site (rSeV-E1805A) were atte‐ nuated in cell culture [91]. It will be of interesting to determine whether these Sendai re‐

Messenger RNA Cap Methylation in Vesicular Stomatitis Virus, a Prototype of Non‐Segmented Negative‐Sense RNA

Virus

259

http://dx.doi.org/10.5772/54598

It appears that the entire mRNA capping and methylation machinery in NNS RNA viruses is different from that of their hosts. This difference, coupled with the fact that replication of NNS RNA viruses occurs in the cytoplasm, suggests that mRNA cap formation is an excel‐ lent target for anti-viral drug discovery. Inhibition of the viral mRNA cap formation would likely inhibit downstream events such as replication, gene expression, viral spread, and ulti‐ mately viral infection. Since the mRNA of all NNS RNA viruses contains a methylated cap structure, classes of broadly active anti-viral agents may be developed by targeting the viral cap formation. For human RSV, several compounds were shown to inhibit polymerase activ‐ ity which resulted in the synthesis of short uncapped transcripts [63]. RSV mutants resistant to these inhibitors were selected and sequenced. It was found that these resistant mutants contained substitutions in CR V of L, specifically at E1269D, I1381S, and L1421F, suggesting that the mechanism of the action of these compounds is the inhibition of viral mRNA cap addition. Interestingly, these compounds showed strong antiviral activity against RSV infec‐ tion in cell culture as well as in a mouse model, demonstrating that mRNA cap addition is an attractive antiviral target. It is known that SAH can inhibit viral mRNA cap methylation. Therefore, many adenosine analogues such as 3-deazaeplanocin-A are potent antiviral agents which can significantly inhibit VSV replication in cell culture [114, 115]. The mecha‐ nism of the action of these adenosine analogues is through the interference with the host en‐ zyme SAH hydrolase that catalyzes the hydrolysis of SAH to adenosine and Lhomocysteine. This reaction is reversible, and the products of this reaction are inhibitory to SAH hydrolase. Obviously, compounds that directly inhibit viral mRNA cap methylation are potent antiviral drugs. For example, sinefungin (SIN), a natural *S*-adenosyl-l-methionine analog produced by *Streptomyces griseolus*, is an inhibitor of methyltransferases. SIN is struc‐ turally related to SAM, with the exception that the methyl group that is donated from SAM is replaced by an amino group in SIN. Crystal structures of several MTases have been solved in complex with SIN, which binds to a region that overlaps the SAM binding site [116]. It was found that SIN inhibited VSV G-N-7 and 2'-O methylation with a 50% inhibitory con‐ centration (IC50) of 2.5 µM and 40 µM, respectively [86]. In cell culture, SIN efficiently inhib‐ ited VSV replication, gene expression, and diminished the size of viral plaques without having significant effect on cell viability. SIN was also shown to inhibit the MTases of other NNS RNA viruses such as NDV [90]. These examples demonstrate that mRNA capping and methylation is an excellent antiviral target for NNS RNA viruses. An important future direc‐ tion is to develop high-throughput screening (HTS) to systemically screen compounds that can inhibit mRNA capping and/or methylation of VSV and other NNS RNA viruses. These

**11. mRNA cap methylation as a target for anti-viral drug discovery**

inhibitors may be broadly active anti-viral agents against NNS RNA viruses.

combinants are attenuated *in vitro*.

#### **10. MTase-defective viruses as live vaccine candidates**

Recombinant viruses defective in MTase can be recovered from cloned full-length viral cDNA by a reverse genetics system. Viruses lacking MTase would likely be attenuated with‐ out affecting immunogenicity, since the MTase is located in L protein, which is not a neu‐ tralizing antibody target. Our group and others have identified a panel of MTase-defective VSV mutants which are attenuated in cell culture as well as in animal models [14, 17, 84, 107]. In addition, MTase-defective Sendai viruses also showed significant defects in viral growth in cell culture [91, 92]. By combining multiple substitutions within the methylation region, it should be possible to generate an attenuated virus that is genetically stable, as re‐ version to wild type at any single amino acid should not provide a fitness gain. Thus, ablat‐ ing viral mRNA cap methylation would provide a new avenue to rationally attenuate these viruses for the development of live attenuated vaccines and exploit their use as viral vectors for vaccines, oncolytic therapy, and gene delivery. Recently, Ma et al., (2012) showed that MTase-defective VSVs were able to induce high levels of VSV-specific antibodies in mice and thus provided full protection against a virulent challenge with the VSV Indiana sero‐ type [107]. Recombinants rVSV-K1651A, D1762A, and E1833Q which were defective in both G-N-7 and 2'-O methylation, are attractive vaccine candidates since they are not only highly attenuated but also retain high immunogenicity. Although recombinants rVSV-G1670A and G1672A retained low virulence to mice, their pathogenicity was significantly reduced com‐ pared to rVSV. The safety of using these two viruses as live vaccine candidates necessitates further investigation.

Our studies on MTase-defective VSVs also shed light on developing live vaccine candidates for other NNS RNA viruses, particularly paramyxoviruses. Within paramyxoviruses,

RSV, hMPV, and PIV3 account for the majority of respiratory diseases infants, children, and the elderly [1-3]. However, there is no vaccine available for these important viruses. Recent research found that live attenuated vaccines are the most promising vaccine can‐ didates for paramyxoviruses [1-3]. However, it has been technically challenging to isolate a virus with low virulence while retaining high immunogenicity. Introducing mutations in the MTase may provide a novel approach to generate live attenuated viruses for these viruses. It was reported that recombinant Sendai virus carrying point mutations in the MTase catalytic site (rSeV-K1782A) and the SAM binding site (rSeV-E1805A) were atte‐ nuated in cell culture [91]. It will be of interesting to determine whether these Sendai re‐ combinants are attenuated *in vitro*.

#### **11. mRNA cap methylation as a target for anti-viral drug discovery**

cates efficiently in a wide range of cell lines including mammalian cells, insect cells, and worms [113]. Unlike WNV, VSV that is specifically defective in G-N-7 methylation can also be successfully recovered [17]. VSV mutants defective in G-N-7 methylation or both G-N-7 and 2'-O methylations can serve as prototypes or controls to elucidate the role of methyla‐ tion in innate immunity. In addition, the VSV mutant (rVSV-G4A) that was attenuated in cell culture retained low to moderate virulence, suggesting the possible role of methylation in averting the innate immune response [107]. Notably, mRNAs of NDV, an avian paramyx‐ ovirus, are not 2'-O methylated [90]. A direct comparison to determine whether 2'-O meth‐ ylation of VSV has a similar biological function compared to WNV and MHV should prove

Recombinant viruses defective in MTase can be recovered from cloned full-length viral cDNA by a reverse genetics system. Viruses lacking MTase would likely be attenuated with‐ out affecting immunogenicity, since the MTase is located in L protein, which is not a neu‐ tralizing antibody target. Our group and others have identified a panel of MTase-defective VSV mutants which are attenuated in cell culture as well as in animal models [14, 17, 84, 107]. In addition, MTase-defective Sendai viruses also showed significant defects in viral growth in cell culture [91, 92]. By combining multiple substitutions within the methylation region, it should be possible to generate an attenuated virus that is genetically stable, as re‐ version to wild type at any single amino acid should not provide a fitness gain. Thus, ablat‐ ing viral mRNA cap methylation would provide a new avenue to rationally attenuate these viruses for the development of live attenuated vaccines and exploit their use as viral vectors for vaccines, oncolytic therapy, and gene delivery. Recently, Ma et al., (2012) showed that MTase-defective VSVs were able to induce high levels of VSV-specific antibodies in mice and thus provided full protection against a virulent challenge with the VSV Indiana sero‐ type [107]. Recombinants rVSV-K1651A, D1762A, and E1833Q which were defective in both G-N-7 and 2'-O methylation, are attractive vaccine candidates since they are not only highly attenuated but also retain high immunogenicity. Although recombinants rVSV-G1670A and G1672A retained low virulence to mice, their pathogenicity was significantly reduced com‐ pared to rVSV. The safety of using these two viruses as live vaccine candidates necessitates

Our studies on MTase-defective VSVs also shed light on developing live vaccine candidates

RSV, hMPV, and PIV3 account for the majority of respiratory diseases infants, children, and the elderly [1-3]. However, there is no vaccine available for these important viruses. Recent research found that live attenuated vaccines are the most promising vaccine can‐ didates for paramyxoviruses [1-3]. However, it has been technically challenging to isolate a virus with low virulence while retaining high immunogenicity. Introducing mutations in the MTase may provide a novel approach to generate live attenuated viruses for these

for other NNS RNA viruses, particularly paramyxoviruses. Within paramyxoviruses,

**10. MTase-defective viruses as live vaccine candidates**

258 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

very compelling.

further investigation.

It appears that the entire mRNA capping and methylation machinery in NNS RNA viruses is different from that of their hosts. This difference, coupled with the fact that replication of NNS RNA viruses occurs in the cytoplasm, suggests that mRNA cap formation is an excel‐ lent target for anti-viral drug discovery. Inhibition of the viral mRNA cap formation would likely inhibit downstream events such as replication, gene expression, viral spread, and ulti‐ mately viral infection. Since the mRNA of all NNS RNA viruses contains a methylated cap structure, classes of broadly active anti-viral agents may be developed by targeting the viral cap formation. For human RSV, several compounds were shown to inhibit polymerase activ‐ ity which resulted in the synthesis of short uncapped transcripts [63]. RSV mutants resistant to these inhibitors were selected and sequenced. It was found that these resistant mutants contained substitutions in CR V of L, specifically at E1269D, I1381S, and L1421F, suggesting that the mechanism of the action of these compounds is the inhibition of viral mRNA cap addition. Interestingly, these compounds showed strong antiviral activity against RSV infec‐ tion in cell culture as well as in a mouse model, demonstrating that mRNA cap addition is an attractive antiviral target. It is known that SAH can inhibit viral mRNA cap methylation. Therefore, many adenosine analogues such as 3-deazaeplanocin-A are potent antiviral agents which can significantly inhibit VSV replication in cell culture [114, 115]. The mecha‐ nism of the action of these adenosine analogues is through the interference with the host en‐ zyme SAH hydrolase that catalyzes the hydrolysis of SAH to adenosine and Lhomocysteine. This reaction is reversible, and the products of this reaction are inhibitory to SAH hydrolase. Obviously, compounds that directly inhibit viral mRNA cap methylation are potent antiviral drugs. For example, sinefungin (SIN), a natural *S*-adenosyl-l-methionine analog produced by *Streptomyces griseolus*, is an inhibitor of methyltransferases. SIN is struc‐ turally related to SAM, with the exception that the methyl group that is donated from SAM is replaced by an amino group in SIN. Crystal structures of several MTases have been solved in complex with SIN, which binds to a region that overlaps the SAM binding site [116]. It was found that SIN inhibited VSV G-N-7 and 2'-O methylation with a 50% inhibitory con‐ centration (IC50) of 2.5 µM and 40 µM, respectively [86]. In cell culture, SIN efficiently inhib‐ ited VSV replication, gene expression, and diminished the size of viral plaques without having significant effect on cell viability. SIN was also shown to inhibit the MTases of other NNS RNA viruses such as NDV [90]. These examples demonstrate that mRNA capping and methylation is an excellent antiviral target for NNS RNA viruses. An important future direc‐ tion is to develop high-throughput screening (HTS) to systemically screen compounds that can inhibit mRNA capping and/or methylation of VSV and other NNS RNA viruses. These inhibitors may be broadly active anti-viral agents against NNS RNA viruses.

#### **12. Concluding remarks**

In recent years, significant progress has been made in understanding the unusual mechanism of mRNA cap addition and methylation employed by VSV. First, VSV mRNA addition utilizes a novel PRNTase that transfers RNA to the GDP acceptor. Second, VSV mRNA cap methylation is catalyzed by a dual MTase that sequentially methylates the position 2'-O followed by G-N-7. Third, PRNTase and dual MTase have been mapped to single amino acid level in CR V and CR VI in the L protein, respectively. Finally, 5' mRNA cap addition and methylation and 3' polyadeny‐ lation are mechanistically and functionally linked. Apparently, the entire mRNA cap formation in VSV evolves a mechanism distinct to hosts. Thus, mRNA cap modification is an ideal target for vaccine and antiviral drug development. However, there are many questions need to be ad‐ dressed. It is not known how polymerase coordinates nucleotide polymerization, mRNA cap addition, cap methylation, and polyadenylation. Although the general mechanism of mRNA modifications is defined, the detailed step in each reaction is still poorly understood. The GTPase required for mRNA capping has not been mapped in L protein. Besides the HR motif, it is not known which step of mRNA capping was affected by other mutations (such as T1157A and G1154A) in CR V of L protein. During mRNA cap methylation the exact mechanism by which the dual MTase methylates the 2'-O and G-N-7 is not known. The crystal structure is not known for L protein or any portion of L protein. The concept of using mRNA cap formation as antiviral target has been experimentally demonstrated, however, high-throughput screening methods toward drug discovery have not been developed. Furthermore, there is urgent need to understand the mRNA cap modification in other NNS RNA virus although it is speculated that they may also be achieved in a similar mechanism.

2 Division of Environmental Health Sciences, College of Public Health, The Ohio State Uni‐

Messenger RNA Cap Methylation in Vesicular Stomatitis Virus, a Prototype of Non‐Segmented Negative‐Sense RNA

Virus

261

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#### **Acknowledgements**

Work in Dr. Jianrong Li's laboratory was supported by grants from NIH/NIAID (R01AI090060), USDA Agriculture and Food Research Initiative (2010-65119-20602), and OSU Center for Clinical and Translational Science (CCTS). Yu Zhang is a fellow of Center for RNA Biology at The Ohio State University. We thank Erin DiCaprio for critical review of this manuscript.

#### **Author details**

Jianrong Li1,2,3\* and Yu Zhang1

\*Address all correspondence to: li.926@osu.edu

1 Department of Food Science and Technology, College of Food, Agricultural and Environ‐ mental Sciences, The Ohio State University, USA

2 Division of Environmental Health Sciences, College of Public Health, The Ohio State Uni‐ versity, USA

3 Center for RNA Biology, The Ohio State University, Columbus, Ohio, USA

#### **References**

**12. Concluding remarks**

260 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

achieved in a similar mechanism.

**Acknowledgements**

this manuscript.

**Author details**

Jianrong Li1,2,3\* and Yu Zhang1

\*Address all correspondence to: li.926@osu.edu

mental Sciences, The Ohio State University, USA

In recent years, significant progress has been made in understanding the unusual mechanism of mRNA cap addition and methylation employed by VSV. First, VSV mRNA addition utilizes a novel PRNTase that transfers RNA to the GDP acceptor. Second, VSV mRNA cap methylation is catalyzed by a dual MTase that sequentially methylates the position 2'-O followed by G-N-7. Third, PRNTase and dual MTase have been mapped to single amino acid level in CR V and CR VI in the L protein, respectively. Finally, 5' mRNA cap addition and methylation and 3' polyadeny‐ lation are mechanistically and functionally linked. Apparently, the entire mRNA cap formation in VSV evolves a mechanism distinct to hosts. Thus, mRNA cap modification is an ideal target for vaccine and antiviral drug development. However, there are many questions need to be ad‐ dressed. It is not known how polymerase coordinates nucleotide polymerization, mRNA cap addition, cap methylation, and polyadenylation. Although the general mechanism of mRNA modifications is defined, the detailed step in each reaction is still poorly understood. The GTPase required for mRNA capping has not been mapped in L protein. Besides the HR motif, it is not known which step of mRNA capping was affected by other mutations (such as T1157A and G1154A) in CR V of L protein. During mRNA cap methylation the exact mechanism by which the dual MTase methylates the 2'-O and G-N-7 is not known. The crystal structure is not known for L protein or any portion of L protein. The concept of using mRNA cap formation as antiviral target has been experimentally demonstrated, however, high-throughput screening methods toward drug discovery have not been developed. Furthermore, there is urgent need to understand the mRNA cap modification in other NNS RNA virus although it is speculated that they may also be

Work in Dr. Jianrong Li's laboratory was supported by grants from NIH/NIAID (R01AI090060), USDA Agriculture and Food Research Initiative (2010-65119-20602), and OSU Center for Clinical and Translational Science (CCTS). Yu Zhang is a fellow of Center for RNA Biology at The Ohio State University. We thank Erin DiCaprio for critical review of

1 Department of Food Science and Technology, College of Food, Agricultural and Environ‐


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**Chapter 11**

**The Methylation of Metals and Metalloids in Aquatic**

This chapter will focus on the formation processes and fate of the more common methylated metals and metalloids in the aquatic environment, focusing on both the ocean and freshwa‐ ter ecosystems. In addition to the formation of the methylated compounds, the biotic and abiotic degradation of these compounds in the natural environment will also be discussed. The formation pathways and the microbes responsible for environmental methylation of dif‐ ferent elements will be examined in detail with the focus on the organometallic or organic metalloid compounds of Hg, As, Sb and Se. Methylated compounds of other metal(loids)s will also be discussed. Such compounds are defined here as those in which the attachment of the organic moiety to the metal/metalloid ion is directly through a carbon-metal bound. Most of these bonds are covalent, especially for the metals and metalloids which have filled d and f orbitals [1]. There is an ever growing field of organometallic chemistry related to the use of manufactured transition metal compounds as catalysts or in organic production syn‐ thesis, or for other uses (e.g. alkylated Pb and butylated Sn compounds). These compounds

Most of the compounds that will be discussed contain one or more methyl group attached to the metal or metalloid atom (Table 1). Methylated halogens are formed in the environment but their formation and fate will not be included in this chapter. Methylation of transition metals does not occur under environmental conditions. In terms of the Periodic Table this chapter will focus on Groups 12-16, but will not directly discuss the major elements of Groups 14-16 (C, N, O, Si, P and S). Organometallic compounds with other alkyl groups (ethylated, butylated or phenyl-metal compounds) in the environment are mostly added as a result of human activity [1]. Most of these methylated compounds are formed biotically in the environment by microorganisms but abiotic pathways of methylation by methyl donor

> © 2013 Mason; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

**Systems**

Robert P. Mason

**1. Introduction**

http://dx.doi.org/10.5772/51774

will not be discussed in detail.

Additional information is available at the end of the chapter

## **The Methylation of Metals and Metalloids in Aquatic Systems**

Robert P. Mason

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51774

#### **1. Introduction**

This chapter will focus on the formation processes and fate of the more common methylated metals and metalloids in the aquatic environment, focusing on both the ocean and freshwa‐ ter ecosystems. In addition to the formation of the methylated compounds, the biotic and abiotic degradation of these compounds in the natural environment will also be discussed. The formation pathways and the microbes responsible for environmental methylation of dif‐ ferent elements will be examined in detail with the focus on the organometallic or organic metalloid compounds of Hg, As, Sb and Se. Methylated compounds of other metal(loids)s will also be discussed. Such compounds are defined here as those in which the attachment of the organic moiety to the metal/metalloid ion is directly through a carbon-metal bound. Most of these bonds are covalent, especially for the metals and metalloids which have filled d and f orbitals [1]. There is an ever growing field of organometallic chemistry related to the use of manufactured transition metal compounds as catalysts or in organic production syn‐ thesis, or for other uses (e.g. alkylated Pb and butylated Sn compounds). These compounds will not be discussed in detail.

Most of the compounds that will be discussed contain one or more methyl group attached to the metal or metalloid atom (Table 1). Methylated halogens are formed in the environment but their formation and fate will not be included in this chapter. Methylation of transition metals does not occur under environmental conditions. In terms of the Periodic Table this chapter will focus on Groups 12-16, but will not directly discuss the major elements of Groups 14-16 (C, N, O, Si, P and S). Organometallic compounds with other alkyl groups (ethylated, butylated or phenyl-metal compounds) in the environment are mostly added as a result of human activity [1]. Most of these methylated compounds are formed biotically in the environment by microorganisms but abiotic pathways of methylation by methyl donor

© 2013 Mason; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

reactions within the aquatic system will also be discussed. Methylation within cells is a fun‐ damental biochemical process and can be carried out by a number of biochemical pathways. It appears, however, that the mechanisms of methylation of metals and metalloids are car‐ ried out by one of three pathways, involving either: S-adenosylmethionine, methylcobala‐ min or N-methyltetrahydrofolate (Figure 1) [1].

could likely be methylated by SAM, methylation of HgII by other pathways appears to be

Bi (CH3)3Bi, (CH3)2BiH, CH3BiH2, (CH3)2B+, CH3Bi2+ Se (CH3)2Se2, (CH3)2Se, (CH3)2SeS, (CH3)2SeH

**Table 1.** Known forms of the various methylated compounds in the environment, including those that are formed by

Therefore, the various pathways of methylation are entirely distinct and which pathway dominates is a function of the differences in the speciation and oxidation state of the metal and metalloid in aquatic systems. For example, there is some evidence for the methylation of As and other metalloids in anoxic sediments via the cobalamin pathway [5, 10, 11]. In con‐ trast to metal cations, the metalloids are present in environmental solutions mostly as an oxyanion of a weak acid, and therefore their form depends on pH. It has been proposed that Hg is taken up as a neutral Hg-sulfide complex via passive diffusion prior to methylation by sulfate-reducing bacteria (SRB), which are thought to be the most important methylating or‐ ganisms [12]. However, there is also the potential for active transport of metals, especially through their uptake via channels designed for major ion transport, such as the channels for phosphate assimilation, or for the acquisition of required metals, such as Fe and Zn, or when

Arsenic is a compound that appears to be methylated by organisms to reduce its toxicity [14]. In low phosphate environments, concentrations of arsenate can rival those of phos‐ phate and given their similar chemistry, organisms can take up AsV inadvertently. Methyla‐ tion appears to be a way to detoxify the As and allow for its secretion into the environment. This is more prevalent in marine waters where As concentrations are higher, as discussed further below. In contrast, methylation of Hg and other metals enhances their toxicity [15]. However, most of the methylated Se compounds are derived from the decomposition of larger organic Se biochemicals such as selenoproteins [16]. Throughout the chapter such

In summary, this chapter will specifically consider the methylation of elements in Groups 12-16 of the periodic table in aquatic systems, with a focus of those present as minor or trace elements. Most of these elements, especially those lower in the periodic table (i.e. higher atomic mass) form covalent bonds with carbon because of the shielding of the nuclear

Sb (CH3)3Sb,(CH3)3SbO, (CH3)2SbH, CH3SbH2

The Methylation of Metals and Metalloids in Aquatic Systems

http://dx.doi.org/10.5772/51774

273

Sn (CH3)4Sn,(CH3)3AsH, (CH3)2SnH2,CH3SnH3

**Element Methylated species Element Methylated species**

Te (CH3)2Te, Tl (CH3)2Te+

hydride generation but could exist as hydrides in the environment. Table compiled from [7-9].

Cd (CH3)2Cd, CH3Cd+ Hg (CH3)2Hg, CH3Hg+, CH3HgH

more efficient.

As (CH3)4As2, (CH3)3As,(CH3)3AsO, (CH3)2AsH,

Ge (CH3)3GeH, (CH3)2GeH2, CH3GeH3, (CH3)2Ge2+,

combined with low molecular weight thiols [13].

CH3Ge3+

Pb (CH3)4Pb, (CH3)3PbH

contrasts will be illustrated.

CH3AsH2, CH3AsO(OH)2, CH3AsO(OH)

**Figure 1.** The primary methylating enzymes for metals and metalloids in organisms: a) SAM, b) tetrahydrofolate and c) methylcobalamin.

For example, methylation of Hg by sulfate-reducing and iron-reducing bacteria [2, 3] in‐ volves methylcobalamin, or related Co-containing enzymes. As ionic Hg (HgII) is the form that is methylated, methylation requires a methyl *carbanion* (CH3 - ), with methylation via a SN2 reaction, and this process is not possible via the other methylation pathways. In con‐ trast, the methyl group given up by S-adenosylmethionine (SAM) and N-methyltetrahydro‐ folate is a *carbocation* (CH3 <sup>+</sup> ). Thus there are fundamental differences in the potential methylating biochemicals and the pathways by which they react with metals and metal‐ loids. For example, the so-called "Challenger" pathway of methylation of As by SAM re‐ quires that initially the AsV is reduced to AsIII and then methylated [4, 5]. The methylated product is oxidized to the AsV state during the methylation step and must be further re‐ duced before addition of more methyl groups. Methylation of Se appears to follow a similar mechanism. In contrast, the methylation of Sn and other cations is thought to involve mainly the cobalamin pathway, in a similar fashion to Hg, with these elements being methylated while in their most oxidized form. There is no concrete evidence for the methylation of re‐ duced Hg (HgI or Hg0 ) in the environment [2]. This is probably a result of the unstable na‐ ture of HgI in the environment and the chemical nature of Hg0 . As a dissolved gas in most environmental aquatic systems it is not accumulated to any significant degree by microor‐ ganisms [6], or other organisms, unless it is oxidized upon uptake. So, therefore while Hg0


could likely be methylated by SAM, methylation of HgII by other pathways appears to be more efficient.

reactions within the aquatic system will also be discussed. Methylation within cells is a fun‐ damental biochemical process and can be carried out by a number of biochemical pathways. It appears, however, that the mechanisms of methylation of metals and metalloids are car‐ ried out by one of three pathways, involving either: S-adenosylmethionine, methylcobala‐

**Figure 1.** The primary methylating enzymes for metals and metalloids in organisms: a) SAM, b) tetrahydrofolate and

For example, methylation of Hg by sulfate-reducing and iron-reducing bacteria [2, 3] in‐ volves methylcobalamin, or related Co-containing enzymes. As ionic Hg (HgII) is the form

SN2 reaction, and this process is not possible via the other methylation pathways. In con‐ trast, the methyl group given up by S-adenosylmethionine (SAM) and N-methyltetrahydro‐

methylating biochemicals and the pathways by which they react with metals and metal‐ loids. For example, the so-called "Challenger" pathway of methylation of As by SAM re‐ quires that initially the AsV is reduced to AsIII and then methylated [4, 5]. The methylated product is oxidized to the AsV state during the methylation step and must be further re‐ duced before addition of more methyl groups. Methylation of Se appears to follow a similar mechanism. In contrast, the methylation of Sn and other cations is thought to involve mainly the cobalamin pathway, in a similar fashion to Hg, with these elements being methylated while in their most oxidized form. There is no concrete evidence for the methylation of re‐

environmental aquatic systems it is not accumulated to any significant degree by microor‐ ganisms [6], or other organisms, unless it is oxidized upon uptake. So, therefore while Hg0

). Thus there are fundamental differences in the potential

) in the environment [2]. This is probably a result of the unstable na‐

), with methylation via a

. As a dissolved gas in most

that is methylated, methylation requires a methyl *carbanion* (CH3 -

in the environment and the chemical nature of Hg0

min or N-methyltetrahydrofolate (Figure 1) [1].

272 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

c) methylcobalamin.

duced Hg (HgI

ture of HgI

folate is a *carbocation* (CH3 <sup>+</sup>

or Hg0

**Table 1.** Known forms of the various methylated compounds in the environment, including those that are formed by hydride generation but could exist as hydrides in the environment. Table compiled from [7-9].

Therefore, the various pathways of methylation are entirely distinct and which pathway dominates is a function of the differences in the speciation and oxidation state of the metal and metalloid in aquatic systems. For example, there is some evidence for the methylation of As and other metalloids in anoxic sediments via the cobalamin pathway [5, 10, 11]. In con‐ trast to metal cations, the metalloids are present in environmental solutions mostly as an oxyanion of a weak acid, and therefore their form depends on pH. It has been proposed that Hg is taken up as a neutral Hg-sulfide complex via passive diffusion prior to methylation by sulfate-reducing bacteria (SRB), which are thought to be the most important methylating or‐ ganisms [12]. However, there is also the potential for active transport of metals, especially through their uptake via channels designed for major ion transport, such as the channels for phosphate assimilation, or for the acquisition of required metals, such as Fe and Zn, or when combined with low molecular weight thiols [13].

Arsenic is a compound that appears to be methylated by organisms to reduce its toxicity [14]. In low phosphate environments, concentrations of arsenate can rival those of phos‐ phate and given their similar chemistry, organisms can take up AsV inadvertently. Methyla‐ tion appears to be a way to detoxify the As and allow for its secretion into the environment. This is more prevalent in marine waters where As concentrations are higher, as discussed further below. In contrast, methylation of Hg and other metals enhances their toxicity [15]. However, most of the methylated Se compounds are derived from the decomposition of larger organic Se biochemicals such as selenoproteins [16]. Throughout the chapter such contrasts will be illustrated.

In summary, this chapter will specifically consider the methylation of elements in Groups 12-16 of the periodic table in aquatic systems, with a focus of those present as minor or trace elements. Most of these elements, especially those lower in the periodic table (i.e. higher atomic mass) form covalent bonds with carbon because of the shielding of the nuclear charge that occurs due to the presence of electrons in filled d and f orbitals. For the metals in Group 12, the tendency to form methylated compounds increases down the group. Howev‐ er, for Groups 15 and 16, the metalloids As and Se are methylated more readily than the ele‐ ments below them in the periodic table, as in these cases the ionic character increases with atomic number. These and other differences within groups result in both similarities and differences in the methylation of the elements and in their stability, fate and transport in the environment, and their ability to biomagnify in aquatic food chains. Before discussing the processes whereby the metal(loids) are is methylated in the environment, it is useful to dis‐ cuss briefly the distribution and fate of these elements in marine and freshwater environ‐ ments.

concentrations at depth are low or below detection, depending on the stability of the methy‐ lated form. In estuarine environments and freshwaters, AsIII and the methylated forms can be a larger fraction of the total dissolved As [20, 26]. The distribution and speciation of Sb is similar to that of As in the ocean water column [14]. The dominant oxidation state is +V, but with the presence of the +III oxidation state in the upper waters and the presence of methy‐ lated species, making up about 10% of the total dissolved Sb. The ancillary parameters shown in Figure 4 for the upper Pacific Ocean [27] give some indication of the physical fac‐ tors influencing the distribution of the metalloids. The density profile shows an area of rapid changing density which coincides with rapid changes in temperature and salinity, which oc‐ curs closer to the surface in the northern latitudes. This stratification separates the more pro‐ ductive surface ocean, where most primary production occurs, from the deeper colder waters where microbial activity is much lower. Therefore, most of the microbial methylation of the metalloids occurs in the upper waters of the ocean, overall within the top 1000 m, and

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**Figure 2.** Vertical profiles of the total metal concentration in open North Pacific Ocean water as compiled by Nozaki 1997. The y-axis is the depth from the surface to 5 km. symbols represent the concentration range: squares for nmol/kg circles for pmol/kg and filled triangle for mBq/kg. Figure reprinted with permission from the American Geo‐

For Se, methylated Se compounds are found in natural waters, as well as both the oxidized and reduced inorganic forms (Figure 3d). The two main inorganic redox states are found but their concentrations are somewhat depleted in the surface waters (Figure 3d), likely due to their uptake and incorporation into biota [14]. Selenium is an essential element although on‐ ly required at low concentrations. The distribution of organic Se (Se-II) suggests its persis‐ tence through the water column, either due to its continual formation and release from microbes and/or from organic matter dissolution, or due to its stability. Volatile methylated

if driven mostly by phytoplankton, within the top 100 m.

physical Union (AGU).

#### **2. The Distribution of Methylated Species in the Ocean and in Freshwaters**

Representative vertical profiles in the North Pacific Ocean for the metal(loid)s discussed in this chapter are shown in Figure 2 [17]. Many of the elements that form methylated species in marine systems (e.g. As, Se, Sb, Ge) exist as oxyanions, and in a number of oxidation states [14, 18-20]. The reduced form of the element is often present because the methylation pathway involves reduction prior to methylation (the so-called Challenger mechanism), as discussed further below, or because of biological or photochemical reduction within organ‐ isms or the water column. In addition to be found as methylated compounds, many metal‐ loids are incorporated into larger metalloid-containing species, such as arsenobetaine and selenoproteins [21, 22]. For example, As is found as AsIII, AsV, mono-, di- and tri-methyl ar‐ senic in marine waters (Figure 3a), and as arsenobetaine, arsenosugars and other As-con‐ taining carbohydrates in organisms. In the open ocean water column, methylation of As is mostly by phytoplankton as this is a detoxification/elimination mechanism. Arsenic (AsV) is taken up inadvertently by microorganisms in low phosphate waters (both exist as polyprotic acids with similar pKa's) and these relatively high concentrations of As can interfere with phosphate biochemistry within the cells.

The methylation of Hg, and other cations (e.g. Sn, Pb), does not involve oxidative methyla‐ tion and therefore the reduced form of these metals, if present, is due to reductive processes that are not directly related to methylation. For Hg, however, the reduced form, elemental Hg (Hg0 ) can be formed through the demethylation of CH3Hg and its subsequent reduction [23, 24]. The presence of the reduced forms of the elements in ocean surface waters, such as Hg0 and AsIII is contrary to what is expected based on thermodynamic equilibrium calcula‐ tions, and this further suggests their formation through microbial processes. However, re‐ duction of Hg can be abiotic as well [25], as its redox couple is such that photochemical reduction is possible in surface waters.

As an example of the distribution of the metalloids in the ocean, profiles of the various spe‐ cies of As and Sb are shown in Figure 3a-c for the North Pacific Ocean. Methylated As and Sb species are present in the surface waters where phytoplankton activity occurs and the concentrations at depth are low or below detection, depending on the stability of the methy‐ lated form. In estuarine environments and freshwaters, AsIII and the methylated forms can be a larger fraction of the total dissolved As [20, 26]. The distribution and speciation of Sb is similar to that of As in the ocean water column [14]. The dominant oxidation state is +V, but with the presence of the +III oxidation state in the upper waters and the presence of methy‐ lated species, making up about 10% of the total dissolved Sb. The ancillary parameters shown in Figure 4 for the upper Pacific Ocean [27] give some indication of the physical fac‐ tors influencing the distribution of the metalloids. The density profile shows an area of rapid changing density which coincides with rapid changes in temperature and salinity, which oc‐ curs closer to the surface in the northern latitudes. This stratification separates the more pro‐ ductive surface ocean, where most primary production occurs, from the deeper colder waters where microbial activity is much lower. Therefore, most of the microbial methylation of the metalloids occurs in the upper waters of the ocean, overall within the top 1000 m, and if driven mostly by phytoplankton, within the top 100 m.

charge that occurs due to the presence of electrons in filled d and f orbitals. For the metals in Group 12, the tendency to form methylated compounds increases down the group. Howev‐ er, for Groups 15 and 16, the metalloids As and Se are methylated more readily than the ele‐ ments below them in the periodic table, as in these cases the ionic character increases with atomic number. These and other differences within groups result in both similarities and differences in the methylation of the elements and in their stability, fate and transport in the environment, and their ability to biomagnify in aquatic food chains. Before discussing the processes whereby the metal(loids) are is methylated in the environment, it is useful to dis‐ cuss briefly the distribution and fate of these elements in marine and freshwater environ‐

274 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**2. The Distribution of Methylated Species in the Ocean and in**

Representative vertical profiles in the North Pacific Ocean for the metal(loid)s discussed in this chapter are shown in Figure 2 [17]. Many of the elements that form methylated species in marine systems (e.g. As, Se, Sb, Ge) exist as oxyanions, and in a number of oxidation states [14, 18-20]. The reduced form of the element is often present because the methylation pathway involves reduction prior to methylation (the so-called Challenger mechanism), as discussed further below, or because of biological or photochemical reduction within organ‐ isms or the water column. In addition to be found as methylated compounds, many metal‐ loids are incorporated into larger metalloid-containing species, such as arsenobetaine and selenoproteins [21, 22]. For example, As is found as AsIII, AsV, mono-, di- and tri-methyl ar‐ senic in marine waters (Figure 3a), and as arsenobetaine, arsenosugars and other As-con‐ taining carbohydrates in organisms. In the open ocean water column, methylation of As is mostly by phytoplankton as this is a detoxification/elimination mechanism. Arsenic (AsV) is taken up inadvertently by microorganisms in low phosphate waters (both exist as polyprotic acids with similar pKa's) and these relatively high concentrations of As can interfere with

The methylation of Hg, and other cations (e.g. Sn, Pb), does not involve oxidative methyla‐ tion and therefore the reduced form of these metals, if present, is due to reductive processes that are not directly related to methylation. For Hg, however, the reduced form, elemental

[23, 24]. The presence of the reduced forms of the elements in ocean surface waters, such as

As an example of the distribution of the metalloids in the ocean, profiles of the various spe‐ cies of As and Sb are shown in Figure 3a-c for the North Pacific Ocean. Methylated As and Sb species are present in the surface waters where phytoplankton activity occurs and the

 and AsIII is contrary to what is expected based on thermodynamic equilibrium calcula‐ tions, and this further suggests their formation through microbial processes. However, re‐ duction of Hg can be abiotic as well [25], as its redox couple is such that photochemical

) can be formed through the demethylation of CH3Hg and its subsequent reduction

ments.

**Freshwaters**

Hg (Hg0

Hg0

phosphate biochemistry within the cells.

reduction is possible in surface waters.

**Figure 2.** Vertical profiles of the total metal concentration in open North Pacific Ocean water as compiled by Nozaki 1997. The y-axis is the depth from the surface to 5 km. symbols represent the concentration range: squares for nmol/kg circles for pmol/kg and filled triangle for mBq/kg. Figure reprinted with permission from the American Geo‐ physical Union (AGU).

For Se, methylated Se compounds are found in natural waters, as well as both the oxidized and reduced inorganic forms (Figure 3d). The two main inorganic redox states are found but their concentrations are somewhat depleted in the surface waters (Figure 3d), likely due to their uptake and incorporation into biota [14]. Selenium is an essential element although on‐ ly required at low concentrations. The distribution of organic Se (Se-II) suggests its persis‐ tence through the water column, either due to its continual formation and release from microbes and/or from organic matter dissolution, or due to its stability. Volatile methylated Se compounds can be formed (analogs to methylated sulfide species, i.e. the suite of com‐ pounds: (CH3)2SySeZ; y=z=0-2; y+z=1 or 2)) in the surface ocean through the decomposition of larger Se-containing biomolecules [28, 29]. The evasion of these compounds could also result in depletion of Se from surface waters [28, 30]. The cycling of Se in the upper ocean is not well understood and needs further study [14].

pounds were in the sub-pM range while concentrations were in the pM range for (CH3)2Se. In each case, there was some evidence of a gradient across the estuary that is likely related to

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Germanium (Ge) is a most unusual element in that its ocean distribution is dominated by its methylated forms (Figure 3e). Germanium is found as the mono- and dimethylated species with the monomethyl species being the dominant form throughout the water column [14]. It has a conservative distribution which indicates its high stability and little is known about its formation mechanisms [32]. There is also little information on the ocean distribution of tin (Sn) (Figure 2) [33]. Much of the focus of study of this element has been due to its use as antifouling agents (e.g. tetrabutyltin) and these compounds will not be discussed in detail. In the surface ocean its concentration is higher than at depth, showing that it is scavenged from the ocean by sinking particles. Bismuth is also higher in surface waters and has a deep

**Figure 4.** Distributions of total mercury, total methylated mercury (sum of mono and dimethylmercury), and their ra‐ tio, temperature, salinity and sigma-theta (a measure of water density) for a transect sampling waters at latitude 152°W in the North Pacific. Taken from Sunderland et al. (2009). Figure reprinted with permssion from AGU.

In the late 1980's, methylated Hg compounds were detected in the water column of the remote ocean [25] and in lakes, and it was shown that the formation of CH3Hg was rela‐ tively ubiquitous in aquatic systems [34]. In ocean waters, and in some freshwater sys‐ tems, dimethylmercury ((CH3)2Hg) has been measured and its presence is mostly found in the water column, although there is some evidence for (CH3)2Hg in sediment porewaters

water mixing and changes in ecosystem productivity.

water scavenged profile (Figure 2) [33].

**Figure 3.** a) Arsenic speciation and distribution in the North Pacific Ocean. Taken from Cutter and Cutter (2006); b) relationship between the relative amount of organic As compounds in seawater and the water temperature. Taken from Santosa et al., 1997; c) Antimony speciation and distribution in the North Pacific Ocean. Taken from Cutter and Cutter (2006); d) Selenium speciation and distribution in the North Pacific Ocean. Taken from Cutter (2010). e) Specia‐ tion of germanium in the North Pacific Ocean. Taken from Cutter (2010). Figures a) and c) reprinted with permission from AGU; b) with permission from Wiley; c) and e) with permission from Elsevier

Tessier et al. [31] developed methods for the simultaneous measurement of a number of vol‐ atile metal(loid) species in various estuaries in France. They were able to detect the methy‐ lated Se species, a variety of Sn compounds, as well as Hg0 . Many of the compounds were found at low concentrations (e.g. (CH3)2Se2, (CH3)2SSe, (C4H9)xSn(CH3)4-x (x=1-3), (CH3)4Sn and (CH3)2Hg) and the dominant species for each element was Hg0 , (CH3)2Se and (C4H9)3SnCH3.The concentration of (CH3)2Se was higher in spring, suggesting production in conjunction with phytoplankton activity. For Sn and Hg, concentrations of volatile com‐ pounds were in the sub-pM range while concentrations were in the pM range for (CH3)2Se. In each case, there was some evidence of a gradient across the estuary that is likely related to water mixing and changes in ecosystem productivity.

Se compounds can be formed (analogs to methylated sulfide species, i.e. the suite of com‐ pounds: (CH3)2SySeZ; y=z=0-2; y+z=1 or 2)) in the surface ocean through the decomposition of larger Se-containing biomolecules [28, 29]. The evasion of these compounds could also result in depletion of Se from surface waters [28, 30]. The cycling of Se in the upper ocean is not

**Figure 3.** a) Arsenic speciation and distribution in the North Pacific Ocean. Taken from Cutter and Cutter (2006); b) relationship between the relative amount of organic As compounds in seawater and the water temperature. Taken from Santosa et al., 1997; c) Antimony speciation and distribution in the North Pacific Ocean. Taken from Cutter and Cutter (2006); d) Selenium speciation and distribution in the North Pacific Ocean. Taken from Cutter (2010). e) Specia‐ tion of germanium in the North Pacific Ocean. Taken from Cutter (2010). Figures a) and c) reprinted with permission

Tessier et al. [31] developed methods for the simultaneous measurement of a number of vol‐ atile metal(loid) species in various estuaries in France. They were able to detect the methy‐

found at low concentrations (e.g. (CH3)2Se2, (CH3)2SSe, (C4H9)xSn(CH3)4-x (x=1-3), (CH3)4Sn

(C4H9)3SnCH3.The concentration of (CH3)2Se was higher in spring, suggesting production in conjunction with phytoplankton activity. For Sn and Hg, concentrations of volatile com‐

and (CH3)2Hg) and the dominant species for each element was Hg0

. Many of the compounds were

, (CH3)2Se and

from AGU; b) with permission from Wiley; c) and e) with permission from Elsevier

lated Se species, a variety of Sn compounds, as well as Hg0

well understood and needs further study [14].

276 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Germanium (Ge) is a most unusual element in that its ocean distribution is dominated by its methylated forms (Figure 3e). Germanium is found as the mono- and dimethylated species with the monomethyl species being the dominant form throughout the water column [14]. It has a conservative distribution which indicates its high stability and little is known about its formation mechanisms [32]. There is also little information on the ocean distribution of tin (Sn) (Figure 2) [33]. Much of the focus of study of this element has been due to its use as antifouling agents (e.g. tetrabutyltin) and these compounds will not be discussed in detail. In the surface ocean its concentration is higher than at depth, showing that it is scavenged from the ocean by sinking particles. Bismuth is also higher in surface waters and has a deep water scavenged profile (Figure 2) [33].

**Figure 4.** Distributions of total mercury, total methylated mercury (sum of mono and dimethylmercury), and their ra‐ tio, temperature, salinity and sigma-theta (a measure of water density) for a transect sampling waters at latitude 152°W in the North Pacific. Taken from Sunderland et al. (2009). Figure reprinted with permssion from AGU.

In the late 1980's, methylated Hg compounds were detected in the water column of the remote ocean [25] and in lakes, and it was shown that the formation of CH3Hg was rela‐ tively ubiquitous in aquatic systems [34]. In ocean waters, and in some freshwater sys‐ tems, dimethylmercury ((CH3)2Hg) has been measured and its presence is mostly found in the water column, although there is some evidence for (CH3)2Hg in sediment porewaters [35]. The relative importance of different bacteria in the production of CH3Hg and (CH3)2Hg in these diverse natural aquatic environments is not clearly understood, although it has been a topic of recent investigation. The distribution of Hg in the North Pacific Ocean is shown in Fig 4 [27]. As can be seen, the distribution of methylated Hg is different from that of As, Sb and the other metalloids as the concentration is low in the surface waters, and maximal in the mid-depth waters. This study and a number of other investigations [25, 27, 36-38] have concluded that the profiles are best explained by formation of methy‐ lated Hg in conjunction with the decomposition of organic matter that sinks from the surface ocean, and that the formation is microbially-mediated. Studies in the Arctic Ocean have shown that in situ methylation occurs in the water column in this region [39], fur‐ ther confirming this pathway. This contrasts lakes where most of the methylation is thought to occur in the sediments [2].

of methylated Hg formation in the ocean and in sediments, although truly anoxic condi‐

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**Figure 5.** a) Changes in the concentration of various parameters including the concentration of bioavailable Hg (HgS0) as well as the methylation rate for sites across the eutrophication gradient in the Florida Everglades. Taken from Be‐ noit et al. (2003); b) The vertical distribution of parameters, including mercury speciation (total (HgT) and methyl-mer‐ cury (MeHg)), and also the rates of microbial processes (mercury methylation (kmeth), sulfate reduction, CO2 and methane production). The sediment reduced sulfide content is indicated by the concentration of labile reduced sul‐ fide (AVS) and pyrite (CRS). The four sample locations include sites in the Chesapeake Bay (A and B) and sites on the shelf (C and D). Taken from Hollweg et al. (2009). Figure a) reprinted with permission of the American Chemical Soci‐

Arsenic speciation and methylation in freshwater ecosystems has also received attention and this has been driven mostly by studies in As contaminated environments. Methylation processes are similar to that found in the ocean with most of the methylated species being confined to the surface waters and levels in deeper waters depending on a variety of condi‐ tions [46-50]. Both seasonally and permanently stratified lakes have been examined and while high levels of the inorganic forms of As are found in anoxic waters, this is not the case for the methylated species. A study in the hydrothermal environment at Yellowstone Na‐ tional Park, USA found that there were volatile As compounds emitted in locations with high aqueous As (up to 50 µM levels) [51]. Concentrations of volatile As in surface waters were from <1 to 2.5 µM. Species identified were (CH3)2AsCl, (CH3)3As, (CH3)2AsCCH3 and (CH3)2AsCl2. There are also some reports of volatile As compounds in some marine environ‐ ments [52]. Studies of the other methylated metal(loid)s in freshwater environments are very limited, with some examination of the formation pathways and speciation of Sb and Se [53-55]. Aspects of the factors influencing formation will be discussed further below. Over‐ all, there is a need for further research on the methylation of metal(loid)s besides As and Hg

tions are not found in the ocean water column.

ety (ACS), b) with permission from Elsevier.

in aquatic systems.

Overall, given the limited data for some elements it is difficult to examine in detail the loca‐ tions and processes of formation of many of the metal(loid)s in marine systems. In summa‐ ry, the metal(loid)s show the following distributions (Figure 2): a) distributions with lower surface ocean concentrations, reflecting uptake in the surface ocean, either into plankton and the food web or by abiotic particles, and release at depth; or b) higher surface water concen‐ trations, reflecting either the dominance of atmospheric inputs (e.g. Pb) or the strong scav‐ enging of the element from deep waters by particles. Overall, however, their distributions are modified to a degree by the formation of methylated or reduced species and the stability and fate of these compounds relative to the more oxidized forms.

Studies of many of the metal(loid)s are also limited for freshwater environments. Mercu‐ ry has been the most studied and the factors controlling Hg methylation in sediments in the Florida Everglades is summarized in Fig 5a [2]. Across the Everglades there is a strong organic matter and sulfate gradient that drives the activity of the SRB's (sulfate reduc‐ tion rate (SRR) in the figure). The product of sulfate reduction, sulfide, complexes Hg and reduces its bioavailability to the methylating organisms, as discussed further below. The bioavailable fraction is represented in Figure 5a as HgS0 , the neutral complex concentra‐ tion. The maximum in methylation rate and the highest fraction as CH3Hg in sediments (%MeHg in the figure) is found at the right combination of bioavailability and bacterial activity, at low sulfide but sufficient sulfate concentrations [2, 40]. Similar results have been found for estuarine and coastal sediments [41-44], as shown in Figure 5b. Higher sulfide levels in coastal sediments result in a strong hindrance of methylation at concentrations above 0.1 mM sulfide in sediment porewaters [2, 42]. This is shown in Figure 5. In the estuarine sediments, lower methylation rates and %CH3Hg are found at depth as sulfide levels increase. The highest fractional conversion of Hg to CH3Hg occurs in low organic matter, shelf sediments where microbial activity is lower but Hg is much more bioavaila‐ ble [41-44]. Studies in lakes show comparable data, and also suggest that there is the po‐ tential for methylation within the water column of seasonally or permanently stratified lakes where oxygen is depleted and sulfide is present [45]. In these environments, the highest methylation is within the redox interface, which is consistent with the discussion of methylated Hg formation in the ocean and in sediments, although truly anoxic condi‐ tions are not found in the ocean water column.

[35]. The relative importance of different bacteria in the production of CH3Hg and (CH3)2Hg in these diverse natural aquatic environments is not clearly understood, although it has been a topic of recent investigation. The distribution of Hg in the North Pacific Ocean is shown in Fig 4 [27]. As can be seen, the distribution of methylated Hg is different from that of As, Sb and the other metalloids as the concentration is low in the surface waters, and maximal in the mid-depth waters. This study and a number of other investigations [25, 27, 36-38] have concluded that the profiles are best explained by formation of methy‐ lated Hg in conjunction with the decomposition of organic matter that sinks from the surface ocean, and that the formation is microbially-mediated. Studies in the Arctic Ocean have shown that in situ methylation occurs in the water column in this region [39], fur‐ ther confirming this pathway. This contrasts lakes where most of the methylation is thought

Overall, given the limited data for some elements it is difficult to examine in detail the loca‐ tions and processes of formation of many of the metal(loid)s in marine systems. In summa‐ ry, the metal(loid)s show the following distributions (Figure 2): a) distributions with lower surface ocean concentrations, reflecting uptake in the surface ocean, either into plankton and the food web or by abiotic particles, and release at depth; or b) higher surface water concen‐ trations, reflecting either the dominance of atmospheric inputs (e.g. Pb) or the strong scav‐ enging of the element from deep waters by particles. Overall, however, their distributions are modified to a degree by the formation of methylated or reduced species and the stability

Studies of many of the metal(loid)s are also limited for freshwater environments. Mercu‐ ry has been the most studied and the factors controlling Hg methylation in sediments in the Florida Everglades is summarized in Fig 5a [2]. Across the Everglades there is a strong organic matter and sulfate gradient that drives the activity of the SRB's (sulfate reduc‐ tion rate (SRR) in the figure). The product of sulfate reduction, sulfide, complexes Hg and reduces its bioavailability to the methylating organisms, as discussed further below. The

tion. The maximum in methylation rate and the highest fraction as CH3Hg in sediments (%MeHg in the figure) is found at the right combination of bioavailability and bacterial activity, at low sulfide but sufficient sulfate concentrations [2, 40]. Similar results have been found for estuarine and coastal sediments [41-44], as shown in Figure 5b. Higher sulfide levels in coastal sediments result in a strong hindrance of methylation at concentrations above 0.1 mM sulfide in sediment porewaters [2, 42]. This is shown in Figure 5. In the estuarine sediments, lower methylation rates and %CH3Hg are found at depth as sulfide levels increase. The highest fractional conversion of Hg to CH3Hg occurs in low organic matter, shelf sediments where microbial activity is lower but Hg is much more bioavaila‐ ble [41-44]. Studies in lakes show comparable data, and also suggest that there is the po‐ tential for methylation within the water column of seasonally or permanently stratified lakes where oxygen is depleted and sulfide is present [45]. In these environments, the highest methylation is within the redox interface, which is consistent with the discussion

, the neutral complex concentra‐

and fate of these compounds relative to the more oxidized forms.

bioavailable fraction is represented in Figure 5a as HgS0

to occur in the sediments [2].

278 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

**Figure 5.** a) Changes in the concentration of various parameters including the concentration of bioavailable Hg (HgS0) as well as the methylation rate for sites across the eutrophication gradient in the Florida Everglades. Taken from Be‐ noit et al. (2003); b) The vertical distribution of parameters, including mercury speciation (total (HgT) and methyl-mer‐ cury (MeHg)), and also the rates of microbial processes (mercury methylation (kmeth), sulfate reduction, CO2 and methane production). The sediment reduced sulfide content is indicated by the concentration of labile reduced sul‐ fide (AVS) and pyrite (CRS). The four sample locations include sites in the Chesapeake Bay (A and B) and sites on the shelf (C and D). Taken from Hollweg et al. (2009). Figure a) reprinted with permission of the American Chemical Soci‐ ety (ACS), b) with permission from Elsevier.

Arsenic speciation and methylation in freshwater ecosystems has also received attention and this has been driven mostly by studies in As contaminated environments. Methylation processes are similar to that found in the ocean with most of the methylated species being confined to the surface waters and levels in deeper waters depending on a variety of condi‐ tions [46-50]. Both seasonally and permanently stratified lakes have been examined and while high levels of the inorganic forms of As are found in anoxic waters, this is not the case for the methylated species. A study in the hydrothermal environment at Yellowstone Na‐ tional Park, USA found that there were volatile As compounds emitted in locations with high aqueous As (up to 50 µM levels) [51]. Concentrations of volatile As in surface waters were from <1 to 2.5 µM. Species identified were (CH3)2AsCl, (CH3)3As, (CH3)2AsCCH3 and (CH3)2AsCl2. There are also some reports of volatile As compounds in some marine environ‐ ments [52]. Studies of the other methylated metal(loid)s in freshwater environments are very limited, with some examination of the formation pathways and speciation of Sb and Se [53-55]. Aspects of the factors influencing formation will be discussed further below. Over‐ all, there is a need for further research on the methylation of metal(loid)s besides As and Hg in aquatic systems.

#### **3. Biotic Transformations**

Of the species that exist in solution as cations, Hg is the element that is methylated to the highest extent, and has been the most studied, primarily as a result of concerns of the hu‐ man and environmental impact of the accumulation of CH3Hg in fish, and their consump‐ tion [56]. The results of these extensive studies can be extrapolated to other cations that are methylated, but more study is also required to investigate their fate and how they are me‐ thylated and bioaccumulated. In terms of the oxyanions, As has received the most attention and again this is driven by concerns over toxicity and the fact that methylation reduces the toxicity of As relative to reduced inorganic As. Methylated compounds of both As and Se can also be formed by the decomposition of larger biochemicals produced in microbes and larger organisms. These pathways will also be discussed in addition to the discussion of the direct methylation pathways.

environmental sulfide concentrations. At higher sulfide, negatively charged Hg-S species and Hg polysulfide species dominate [69]. Therefore, methylation is most prevalent in envi‐ ronments where sulfate reduction is high, but sulfide accumulation is low (Figure 5) [2]. These locations are often the upper layers of sediments, as inputs of fresh organic matter and oxygen diffusion into sediments result in high rates of Fe and sulfate reduction in re‐ gions of relatively low sulfide content. Other complexes of Hg are potentially bioavailable and complexes with small organic ligands can have a relatively high Kow [70]. This is dem‐ onstrated by the fact that a bacterial culture of *Geobacter sulfurreducens* can methylate HgII when present in the medium as a cysteine complex at a much faster rate than as either sul‐ fide or chloride complexes [71, 72]. Most studies therefore highlight the importance of speci‐ ation in determining the rate of methylation in sediments and freshwater systems but the role and activity of the methylating bacteria cannot be ignored. It has been well demonstrat‐ ed that speciation affects the uptake and methylation of inorganic Hg and the same impact is probable on the rate of demethylation of CH3Hg, and on the uptake and methylation of

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The discussion above about methylation in sediments and freshwater systems does not ex‐ plain the sources and sinks for CH3Hg in the open ocean. A recent examination of the sour‐ ces and sinks for CH3Hg and (CH3)2Hg (hereafter represented as ∑CH3Hg) to the ocean [25] suggest that external sources (riverine inputs and coastal sources and atmospheric deposi‐ tion) are insufficient to account for the ∑CH3Hg sinks in the ocean, which include accumula‐ tion into biota and removal by fisheries, photochemical and biological degradation into inorganic Hg, and net removal to the deep ocean and deep sea sediments. This indicates that production within the ocean water column is important. While both CH3Hg and (CH3)2Hg are broadly distributed throughout the ocean water column, they have observed concentra‐ tions are difficult to explain without *in situ* production [25]. Initial studies in the equatorial Pacific Ocean suggested sub-thermocline maxima in both CH3Hg and (CH3)2Hg [38, 73-75] have been confirmed by more recent studies in the North and South Atlantic and Pacific Oceans, the Mediterranean Sea, the Southern Ocean and other locations [27, 36, 37, 76-80]. Figure 4 is representative of a typical profile for the upper ocean. These vertical distributions are most consistent with *in situ* formation of ∑CH3Hg in association with the decomposition of organic matter. The link to organic carbon degradations is demonstrated, for example, by the relationship between the amount of ∑CH3Hg and the extent of organic carbon reminer‐ alization [27], and correlations between ∑CH3Hg and apparent oxygen utilization, another measure of carbon degradation [36-38, 75, 78]. The ∑CH3Hg distribution across studies sug‐ gests that the transition regions (the base of the euphotic zone), and subsurface waters where particulate organic matter is being degraded, are locations of enhanced net methyla‐ tion of Hg. However, there is still little concrete information about the microbes responsible for Hg methylation and for the primary factors driving the relative magnitude of this proc‐

other cations.

ess across ocean basins.

#### **3.1. The biotic methylation of mercury**

Acute exposure incidents in Japan and elsewhere in the 1960's provided the first evidence that inorganic Hg could be converted into the more toxic and bioaccumulative CH3Hg [15, 56]. Methylation of Hg in sediments by the resident microorganisms was subsequently dem‐ onstrated and many of the early studies examining the role of microbes in methylation were in coastal and estuarine sediments [34]. Selective microbial inhibition studies indicated that methylation was primarily mediated by SRB's [57, 58] but recent studies suggest that phylo‐ genetically similar microbes (e.g. Fe-reducers in the genera *Geobacter*) can also methylate Hg [3, 59]. Additionally, there has been recent demonstration of methylation being related to the activity of methanogens in lake periphyton [60] and it is possible that further examination of different environments may lead to further understanding of the Hg methylation process. The site, or more likely sites, of methylation within cells has not been clearly identified and not all SRB's methylate Hg, and even organisms from the same genus has different abilities to methylate Hg [2]. While initial studies suggested that methylation was associated with the Acetyl-CoA pathway and cobalamin in one estuarine organism (*Desulfovibrio desulfuri‐ cans* LS) [61-63], this link does not hold across a variety of organisms, as organisms without this pathway methylate Hg, and *vice versa* [64, 65]. For SRB's, both complete and incomplete organic carbon oxidizers are methylators of Hg. Recent studies have also shown differences in the rate of methylation in biofilms compared to free living SRB in culture [66]. It was shown that the gene expression, and in particular the activity of the Acetyl-CoA pathway, was different for the same organism when growing under these different conditions and that the expression of the pathway coincided with the increased methylation in the biofilms. These results appear to confirm the notion that methylation is accidental and not a detoxify‐ ing mechanism in these organisms.

In a number of ecosystems, CH3Hg production has been shown to be strongly related to Hg interactions with sulfide, e.g. [2, 43, 44, 67]. Neutrally charged dissolved Hg-S species (HOHgSH and Hg(SH)2) are believed to be much more bioavailable to sulfate-reducing bac‐ teria [12, 68] than charged complexes. They have a relatively high Kow and dominate at low environmental sulfide concentrations. At higher sulfide, negatively charged Hg-S species and Hg polysulfide species dominate [69]. Therefore, methylation is most prevalent in envi‐ ronments where sulfate reduction is high, but sulfide accumulation is low (Figure 5) [2]. These locations are often the upper layers of sediments, as inputs of fresh organic matter and oxygen diffusion into sediments result in high rates of Fe and sulfate reduction in re‐ gions of relatively low sulfide content. Other complexes of Hg are potentially bioavailable and complexes with small organic ligands can have a relatively high Kow [70]. This is dem‐ onstrated by the fact that a bacterial culture of *Geobacter sulfurreducens* can methylate HgII when present in the medium as a cysteine complex at a much faster rate than as either sul‐ fide or chloride complexes [71, 72]. Most studies therefore highlight the importance of speci‐ ation in determining the rate of methylation in sediments and freshwater systems but the role and activity of the methylating bacteria cannot be ignored. It has been well demonstrat‐ ed that speciation affects the uptake and methylation of inorganic Hg and the same impact is probable on the rate of demethylation of CH3Hg, and on the uptake and methylation of other cations.

**3. Biotic Transformations**

280 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

direct methylation pathways.

**3.1. The biotic methylation of mercury**

ing mechanism in these organisms.

Of the species that exist in solution as cations, Hg is the element that is methylated to the highest extent, and has been the most studied, primarily as a result of concerns of the hu‐ man and environmental impact of the accumulation of CH3Hg in fish, and their consump‐ tion [56]. The results of these extensive studies can be extrapolated to other cations that are methylated, but more study is also required to investigate their fate and how they are me‐ thylated and bioaccumulated. In terms of the oxyanions, As has received the most attention and again this is driven by concerns over toxicity and the fact that methylation reduces the toxicity of As relative to reduced inorganic As. Methylated compounds of both As and Se can also be formed by the decomposition of larger biochemicals produced in microbes and larger organisms. These pathways will also be discussed in addition to the discussion of the

Acute exposure incidents in Japan and elsewhere in the 1960's provided the first evidence that inorganic Hg could be converted into the more toxic and bioaccumulative CH3Hg [15, 56]. Methylation of Hg in sediments by the resident microorganisms was subsequently dem‐ onstrated and many of the early studies examining the role of microbes in methylation were in coastal and estuarine sediments [34]. Selective microbial inhibition studies indicated that methylation was primarily mediated by SRB's [57, 58] but recent studies suggest that phylo‐ genetically similar microbes (e.g. Fe-reducers in the genera *Geobacter*) can also methylate Hg [3, 59]. Additionally, there has been recent demonstration of methylation being related to the activity of methanogens in lake periphyton [60] and it is possible that further examination of different environments may lead to further understanding of the Hg methylation process. The site, or more likely sites, of methylation within cells has not been clearly identified and not all SRB's methylate Hg, and even organisms from the same genus has different abilities to methylate Hg [2]. While initial studies suggested that methylation was associated with the Acetyl-CoA pathway and cobalamin in one estuarine organism (*Desulfovibrio desulfuri‐ cans* LS) [61-63], this link does not hold across a variety of organisms, as organisms without this pathway methylate Hg, and *vice versa* [64, 65]. For SRB's, both complete and incomplete organic carbon oxidizers are methylators of Hg. Recent studies have also shown differences in the rate of methylation in biofilms compared to free living SRB in culture [66]. It was shown that the gene expression, and in particular the activity of the Acetyl-CoA pathway, was different for the same organism when growing under these different conditions and that the expression of the pathway coincided with the increased methylation in the biofilms. These results appear to confirm the notion that methylation is accidental and not a detoxify‐

In a number of ecosystems, CH3Hg production has been shown to be strongly related to Hg interactions with sulfide, e.g. [2, 43, 44, 67]. Neutrally charged dissolved Hg-S species (HOHgSH and Hg(SH)2) are believed to be much more bioavailable to sulfate-reducing bac‐ teria [12, 68] than charged complexes. They have a relatively high Kow and dominate at low

The discussion above about methylation in sediments and freshwater systems does not ex‐ plain the sources and sinks for CH3Hg in the open ocean. A recent examination of the sour‐ ces and sinks for CH3Hg and (CH3)2Hg (hereafter represented as ∑CH3Hg) to the ocean [25] suggest that external sources (riverine inputs and coastal sources and atmospheric deposi‐ tion) are insufficient to account for the ∑CH3Hg sinks in the ocean, which include accumula‐ tion into biota and removal by fisheries, photochemical and biological degradation into inorganic Hg, and net removal to the deep ocean and deep sea sediments. This indicates that production within the ocean water column is important. While both CH3Hg and (CH3)2Hg are broadly distributed throughout the ocean water column, they have observed concentra‐ tions are difficult to explain without *in situ* production [25]. Initial studies in the equatorial Pacific Ocean suggested sub-thermocline maxima in both CH3Hg and (CH3)2Hg [38, 73-75] have been confirmed by more recent studies in the North and South Atlantic and Pacific Oceans, the Mediterranean Sea, the Southern Ocean and other locations [27, 36, 37, 76-80]. Figure 4 is representative of a typical profile for the upper ocean. These vertical distributions are most consistent with *in situ* formation of ∑CH3Hg in association with the decomposition of organic matter. The link to organic carbon degradations is demonstrated, for example, by the relationship between the amount of ∑CH3Hg and the extent of organic carbon reminer‐ alization [27], and correlations between ∑CH3Hg and apparent oxygen utilization, another measure of carbon degradation [36-38, 75, 78]. The ∑CH3Hg distribution across studies sug‐ gests that the transition regions (the base of the euphotic zone), and subsurface waters where particulate organic matter is being degraded, are locations of enhanced net methyla‐ tion of Hg. However, there is still little concrete information about the microbes responsible for Hg methylation and for the primary factors driving the relative magnitude of this proc‐ ess across ocean basins.

#### **3.2. The biotic methylation of arsenic and antimony**

The methylation of As (Figure 6) is thought to be a detoxifying mechanism for bacteria and phytoplankton [5], especially those in the marine environment where the concentration of As is relatively high compared to the required nutrient phosphate, as both exist in solution as oxyanions. However, the first indication of As methylation was the demonstration of the release of volatile As-containing compounds from wallpaper in the 1800's in Europe, from the use of As-containing pigments in their coloration [4, 5]. It was further realized that mold and damp enhanced their formation and therefore a biological role for the production of these volatile compounds was concluded, mostly based on work done by an Italian physician, Gosio. This early work was followed by Challenger and his colleagues in the early 20th century, who developed the original mechanism for As methylation that still bears his name [4].

with the most common compound being arsenobetaine (trimethylarsinio acetate)

of the methylated derivatives and that they are ultimately generated by processes similar to that invoked in the Challenger mechanism. The methylated species are also found in meas‐ urable concentrations in many environmental waters, both freshwater and marine, as dis‐ cussed above, and their presence has mostly been correlated with phytoplankton activity although their production via bacteria present in conjunction with the algae is also possible.

**Figure 6.** Representation of the steps involved in the methylation of arsenic via the Challenger process. Figure taken

Based on the pKa values, it is evident that the reduced form of As is present as an undissoci‐ ated acid at physiological pH's and in most environmental solutions, and therefore that there is the possibility of its loss via passive diffusion from the cells into the surrounding media. There are a number of instances where measurements in the environment have shown that the concentration of reduced inorganic As is greater than that of AsV, and release of AsIII after microbial reduction is the most likely explanation [16]. There have been a num‐ ber of As reductases indentified in microbes and these processes appear to be separate from the methylation pathways, and therefore involve different reductants, such as thioredoxin and glutaredoxin [4]. Similar to other metal reductases, the operons are attached to plasmids

Antimony (Sb) has a similar chemistry to As and therefore it is expected to behave in a simi‐ lar manner to As in terms of uptake and methylation [7]. Its compounds have been widely

from Feldman (2003) and used with permission from Wiley.

in bacteria.

It is suggested that these compounds are actively exported by the phytoplankton.

) [21]. It is thought that these products are formed from further reactions

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((CH3)3AsCH2COO-

It is now known that methylation of As and other metalloids is carried out by a wide variety of fungi, yeasts and bacteria, and eukaryotes [4, 21, 81]. Methylation of As is the most preva‐ lent of the elements in Group 15 occurring in microbes, algae, plants and animals while methylation of Sb is more restricted to eukaryotic and prokaryotic microorganisms. It is thought that As is mainly taken up by phytoplankton through phosphate uptake channels. It is however also possible that the methylated As species are formed during the degrada‐ tion of As biochemicals during degradation of cellular material [4]. Inorganic As is present in environmental waters as an oxyacid in both oxidation states: arsenic acid (arsenate) AsO(OH)3 or H3AsO4 (pKa1 = 2.2; pKa2 = 7.0; pKa3 = 11.5) and arsenous acid (arsenite) As(OH)3 (pKa1 = 9.3). The pKa's of arsenic acid are very similar to those of phosphate (re‐ spectively, 2.1, 7.2 and 12.4) and therefore the speciation of AsV and phosphate in most envi‐ ronmental waters is analogous. At the typical pH of environmental waters, the major species will be H2AsO4 and HAsO4 2-. It generally appears that phosphate uptake is active and in conjunction with Na+ co-transport and therefore a similar mechanism is likely for As. In open ocean surface waters, phosphate concentrations are often very low (<0.5 µM) and As concentrations are typically around 10-20 nM [14, 20]. Therefore, given this relatively small difference in concentration, inadvertent uptake of As is possible [21]. The methylated forms of As can be released from the cells into the environment or they can be incorporated into larger molecules and their toxicity reduced as a result.

The Challenger mechanism of As, and other metalloid, methylation involves a series of re‐ ductive methylation steps where the addition of the methyl group is via a carbocation addi‐ tion reaction (Figure 6). The details of the mechanism are relatively well known although some aspects of the biochemistry within the cell are not entirely elucidated. It is thought that cellular thiols, such as glutathione, are involved in the reduction steps and that S-adenosyl‐ methionine (SAM) is the main methylating agent. The reduction is linked to glutathione oxi‐ dation and is enzymatically controlled. Arsenite and related methyltransferase enzymes are associated with the methylation step [4, 8]. The products are the simple reduced methylated species ((CH3)xAsO(OH)3-x, x=1-3) as well as oxidized forms, such as trimethylarsine oxide ((CH3)3AsO- ). Volatile forms are also produced by some organisms (Table 1), and these are the methylated As hydrides, such as (CH3)xAsH3-x; x = 1-3) [9]. In marine algae, As is found in a variety of organic compounds, such as arsenosugars and As-containing carbohydrates, with the most common compound being arsenobetaine (trimethylarsinio acetate) ((CH3)3AsCH2COO- ) [21]. It is thought that these products are formed from further reactions of the methylated derivatives and that they are ultimately generated by processes similar to that invoked in the Challenger mechanism. The methylated species are also found in meas‐ urable concentrations in many environmental waters, both freshwater and marine, as dis‐ cussed above, and their presence has mostly been correlated with phytoplankton activity although their production via bacteria present in conjunction with the algae is also possible. It is suggested that these compounds are actively exported by the phytoplankton.

**3.2. The biotic methylation of arsenic and antimony**

282 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

larger molecules and their toxicity reduced as a result.

will be H2AsO4 -

((CH3)3AsO-

conjunction with Na+

The methylation of As (Figure 6) is thought to be a detoxifying mechanism for bacteria and phytoplankton [5], especially those in the marine environment where the concentration of As is relatively high compared to the required nutrient phosphate, as both exist in solution as oxyanions. However, the first indication of As methylation was the demonstration of the release of volatile As-containing compounds from wallpaper in the 1800's in Europe, from the use of As-containing pigments in their coloration [4, 5]. It was further realized that mold and damp enhanced their formation and therefore a biological role for the production of these volatile compounds was concluded, mostly based on work done by an Italian physician, Gosio. This early work was followed by Challenger and his colleagues in the early 20th century, who

developed the original mechanism for As methylation that still bears his name [4].

It is now known that methylation of As and other metalloids is carried out by a wide variety of fungi, yeasts and bacteria, and eukaryotes [4, 21, 81]. Methylation of As is the most preva‐ lent of the elements in Group 15 occurring in microbes, algae, plants and animals while methylation of Sb is more restricted to eukaryotic and prokaryotic microorganisms. It is thought that As is mainly taken up by phytoplankton through phosphate uptake channels. It is however also possible that the methylated As species are formed during the degrada‐ tion of As biochemicals during degradation of cellular material [4]. Inorganic As is present in environmental waters as an oxyacid in both oxidation states: arsenic acid (arsenate) AsO(OH)3 or H3AsO4 (pKa1 = 2.2; pKa2 = 7.0; pKa3 = 11.5) and arsenous acid (arsenite) As(OH)3 (pKa1 = 9.3). The pKa's of arsenic acid are very similar to those of phosphate (re‐ spectively, 2.1, 7.2 and 12.4) and therefore the speciation of AsV and phosphate in most envi‐ ronmental waters is analogous. At the typical pH of environmental waters, the major species

open ocean surface waters, phosphate concentrations are often very low (<0.5 µM) and As concentrations are typically around 10-20 nM [14, 20]. Therefore, given this relatively small difference in concentration, inadvertent uptake of As is possible [21]. The methylated forms of As can be released from the cells into the environment or they can be incorporated into

The Challenger mechanism of As, and other metalloid, methylation involves a series of re‐ ductive methylation steps where the addition of the methyl group is via a carbocation addi‐ tion reaction (Figure 6). The details of the mechanism are relatively well known although some aspects of the biochemistry within the cell are not entirely elucidated. It is thought that cellular thiols, such as glutathione, are involved in the reduction steps and that S-adenosyl‐ methionine (SAM) is the main methylating agent. The reduction is linked to glutathione oxi‐ dation and is enzymatically controlled. Arsenite and related methyltransferase enzymes are associated with the methylation step [4, 8]. The products are the simple reduced methylated species ((CH3)xAsO(OH)3-x, x=1-3) as well as oxidized forms, such as trimethylarsine oxide

the methylated As hydrides, such as (CH3)xAsH3-x; x = 1-3) [9]. In marine algae, As is found in a variety of organic compounds, such as arsenosugars and As-containing carbohydrates,

). Volatile forms are also produced by some organisms (Table 1), and these are

and HAsO4 2-. It generally appears that phosphate uptake is active and in

co-transport and therefore a similar mechanism is likely for As. In

**Figure 6.** Representation of the steps involved in the methylation of arsenic via the Challenger process. Figure taken from Feldman (2003) and used with permission from Wiley.

Based on the pKa values, it is evident that the reduced form of As is present as an undissoci‐ ated acid at physiological pH's and in most environmental solutions, and therefore that there is the possibility of its loss via passive diffusion from the cells into the surrounding media. There are a number of instances where measurements in the environment have shown that the concentration of reduced inorganic As is greater than that of AsV, and release of AsIII after microbial reduction is the most likely explanation [16]. There have been a num‐ ber of As reductases indentified in microbes and these processes appear to be separate from the methylation pathways, and therefore involve different reductants, such as thioredoxin and glutaredoxin [4]. Similar to other metal reductases, the operons are attached to plasmids in bacteria.

Antimony (Sb) has a similar chemistry to As and therefore it is expected to behave in a simi‐ lar manner to As in terms of uptake and methylation [7]. Its compounds have been widely used in industry, medicine, as a poison, and as cosmetics. The methylated forms, (CH3)xSb3-x (x = 1-3), are well-known (Table 1), as are many other organo-Sb compounds, and many have been synthesized for industrial purposes. Volatile Sb compounds were suggested as a possible cause for Sudden Infant Death Syndrome, as Sb compounds were used as flame re‐ tardants in mattresses, and this lead to the examination of the mechanisms of their forma‐ tion [4, 7]. A number of fungi and bacteria have been identified that can methylate Sb, and methylation is much higher in the presence of SbIII and it is apparent that many organisms are not able to reduce and methylate SbV [54]. Therefore, the methylation of Sb may be less ubiquitous than that of As, and there appears to be complex interactions if both As and Sb are present in the same culture in terms of relative methylation [4, 7]. Overall, little is known of the speciation and form of Sb in natural waters, and the formula is given either as Sb(OH)5 or HSb(OH)6, which dissociates in water to form the anion, Sb(OH)6 - (pKa1 =2.2). There is evidence from measurements in seawater and in the presence of marine algae that reduction to SbIII occurs and that the monomethylated form exists in environmental waters [54]. It is also present in freshwaters, and evidence for its formation in environments such as landfills [56]. Overall, however, it appears that SbV is the major form in environmental solu‐ tions. The presence of volatile Sb compounds in landfill gases and other methanogenic envi‐ ronments confirms that methylation of Sb is microbially mediated [4, 82].

Also, in most cases the porewater concentrations were similar or lower than those of the

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The relative concentration of the various compounds changed over the course of the incuba‐ tions in both these experiments [83, 84] and these changes are likely related to the changing microbial community with time and the sequential nature of the methylation processes, as well as demethylation. Field sampling of sediment porewater confirmed the presence of these compounds in the environment, but also showed that these compounds were a small fraction (<1%) of the total As. Another study of estuarine porewaters also found the pres‐ ence of methylated As, but again these were a relative small fraction of the total (<4%) [85]. This contrasts the oxic water column where higher relative amounts occur (Figure 3). Re‐ sults with lake sediments [86] were similar as these studies also suggested that anaerobic bacteria (Fe, Mn or sulfate-reducing bacteria) were mostly responsible for the transforma‐ tions found. Overall, these studies suggest that methylation of As and other metalloids can occur in sediments and that anaerobic bacteria are responsible for the methylation, contrast‐ ing the formation mechanisms in the oxic environment. Overall, the links and interactions between the various pathways are complex, and it is difficult to distinguish the relative im‐ portance of the various processes in As methylation in sediments and other environments.

Selenium, an element in Group 16, has important and complex organometallic chemistry, and is readily methylated. In organisms, Se has a large number of biochemical roles as it is incorporated into a number of enzymes [52]. The methylation of Se was first investigated by Challenger and since these early studies it has been shown that Se has an extensive bio‐ chemistry and that compounds, such as selenoproteins are important constituents of organ‐ isms as they act as anti-oxidants and have other roles in the cellular machinery [16, 81, 87]. Compounds such as selenomethionine and Se-adenosylselenomethionine (SeSAM), are found in cells, where Se has replaced S, as could be expected as these elements are from the same group of the periodic table. The methylating ability of SeSAM has been shown to be

sponsible for the toxicity of Se. Additionally, analogs to the methylated S-containing com‐ pounds (CH3)2S (DMS) and (CH3)2S2 have been identified ((CH3)2Se, (CH3)2SSe and (CH3)2Se2) [7]. It appears likely that these compounds are the degradation products pro‐ duced by certain microorganisms that exist in ocean waters, being formed from the decom‐

processes likely account for their formation in terrestrial waters and other environments. The importance of this pathway was confirmed by growing a freshwater green algae on selenate in the absence of sulfate and showing the formation of the volatile methylated spe‐ cies [53]. This production was reduced when sulfate was added to the medium. Another study also demonstrated the release of volatile methylated Se compounds from green algae. This study also showed the potential for As to inhibit the formation of the Se compounds

CH2CH2COO-

are some of the compounds re‐

) (Figure 7) [7]. Similar

**3.3. The biotic formation of methylated selenium compounds**

It has been suggested that CH3SeH and the cation CH3Se+

position of 3-dimethyl-selenopropionate ((CH3)2Se+

overlying water.

greater than that of SAM [7].

Most studies have invoked the Challenger mechanism to explain the methylation of As and other metalloids by SAM in the environment, especially in oxic waters but there is some speculation that the pathway of methylation may be different for anaerobic organisms. In this case, it has been suggested that methylation may involve cobalamin and therefore involve a different mechanism whereby a carbanion or a radical from methylcobalamin is added to AsIII in the presence of mediating enzymes [4, 7]. Methanogenic Achaea, for example, were shown to methylate a variety of metalloids of Groups 15 and 16 (As, Se, Sb, Te, and Bi) [10]. The mechanism was attributed to side reactions with methylcobalamin. Additionally, it was dem‐ onstrated that methylation of As(V) did not occur and that methylation of As(III) did not involve oxidation, and was therefore similar in process to the methylation of Hg whereby the methyl group is added as a carbanion rather than a carbocation. While Weufel et al. [10] were able to demonstrate the formation of higher methylated compounds, another study [11] suggested that the reaction pathway produces only monomethylated forms, which contrasts the environment where higher methylated forms are often more abundant. In addition to methylation of As, cobalamin has also been shown to methylate Sb [11].

Studies have also focused on the methylation of As and other metalloids in sediments [83]. Laboratory incubation of marine sediments have produced volatile arsines ((CH3)xAsH3-x; x=0-3) and other methylated compounds ((CH3)xAsO(OH)3-x; x=1-3). In other experiments, incubation of sediments showed the production of both methylated As and Sb species [84]. It appeared that the dimethylated species dominated for both metalloids. Also, the initial rate of formation of the methylated species was faster for As than for Sb. After the comple‐ tion of the experiment (76 days) all methylated forms of As and Sb were found distributed through the sediment column (0-12 cm) and in the overlying water. While the experiment lasted 76 days, in many instances the peak in concentration occurred relatively early in the experiment suggesting demethylation was occurring in the latter parts of the experiment. Also, in most cases the porewater concentrations were similar or lower than those of the overlying water.

The relative concentration of the various compounds changed over the course of the incuba‐ tions in both these experiments [83, 84] and these changes are likely related to the changing microbial community with time and the sequential nature of the methylation processes, as well as demethylation. Field sampling of sediment porewater confirmed the presence of these compounds in the environment, but also showed that these compounds were a small fraction (<1%) of the total As. Another study of estuarine porewaters also found the pres‐ ence of methylated As, but again these were a relative small fraction of the total (<4%) [85]. This contrasts the oxic water column where higher relative amounts occur (Figure 3). Re‐ sults with lake sediments [86] were similar as these studies also suggested that anaerobic bacteria (Fe, Mn or sulfate-reducing bacteria) were mostly responsible for the transforma‐ tions found. Overall, these studies suggest that methylation of As and other metalloids can occur in sediments and that anaerobic bacteria are responsible for the methylation, contrast‐ ing the formation mechanisms in the oxic environment. Overall, the links and interactions between the various pathways are complex, and it is difficult to distinguish the relative im‐ portance of the various processes in As methylation in sediments and other environments.

#### **3.3. The biotic formation of methylated selenium compounds**

used in industry, medicine, as a poison, and as cosmetics. The methylated forms, (CH3)xSb3-x (x = 1-3), are well-known (Table 1), as are many other organo-Sb compounds, and many have been synthesized for industrial purposes. Volatile Sb compounds were suggested as a possible cause for Sudden Infant Death Syndrome, as Sb compounds were used as flame re‐ tardants in mattresses, and this lead to the examination of the mechanisms of their forma‐ tion [4, 7]. A number of fungi and bacteria have been identified that can methylate Sb, and methylation is much higher in the presence of SbIII and it is apparent that many organisms are not able to reduce and methylate SbV [54]. Therefore, the methylation of Sb may be less ubiquitous than that of As, and there appears to be complex interactions if both As and Sb are present in the same culture in terms of relative methylation [4, 7]. Overall, little is known of the speciation and form of Sb in natural waters, and the formula is given either as Sb(OH)5 or HSb(OH)6, which dissociates in water to form the anion, Sb(OH)6 - (pKa1 =2.2). There is evidence from measurements in seawater and in the presence of marine algae that reduction to SbIII occurs and that the monomethylated form exists in environmental waters [54]. It is also present in freshwaters, and evidence for its formation in environments such as landfills [56]. Overall, however, it appears that SbV is the major form in environmental solu‐ tions. The presence of volatile Sb compounds in landfill gases and other methanogenic envi‐

284 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

ronments confirms that methylation of Sb is microbially mediated [4, 82].

methylation of As, cobalamin has also been shown to methylate Sb [11].

Most studies have invoked the Challenger mechanism to explain the methylation of As and other metalloids by SAM in the environment, especially in oxic waters but there is some speculation that the pathway of methylation may be different for anaerobic organisms. In this case, it has been suggested that methylation may involve cobalamin and therefore involve a different mechanism whereby a carbanion or a radical from methylcobalamin is added to AsIII in the presence of mediating enzymes [4, 7]. Methanogenic Achaea, for example, were shown to methylate a variety of metalloids of Groups 15 and 16 (As, Se, Sb, Te, and Bi) [10]. The mechanism was attributed to side reactions with methylcobalamin. Additionally, it was dem‐ onstrated that methylation of As(V) did not occur and that methylation of As(III) did not involve oxidation, and was therefore similar in process to the methylation of Hg whereby the methyl group is added as a carbanion rather than a carbocation. While Weufel et al. [10] were able to demonstrate the formation of higher methylated compounds, another study [11] suggested that the reaction pathway produces only monomethylated forms, which contrasts the environment where higher methylated forms are often more abundant. In addition to

Studies have also focused on the methylation of As and other metalloids in sediments [83]. Laboratory incubation of marine sediments have produced volatile arsines ((CH3)xAsH3-x; x=0-3) and other methylated compounds ((CH3)xAsO(OH)3-x; x=1-3). In other experiments, incubation of sediments showed the production of both methylated As and Sb species [84]. It appeared that the dimethylated species dominated for both metalloids. Also, the initial rate of formation of the methylated species was faster for As than for Sb. After the comple‐ tion of the experiment (76 days) all methylated forms of As and Sb were found distributed through the sediment column (0-12 cm) and in the overlying water. While the experiment lasted 76 days, in many instances the peak in concentration occurred relatively early in the experiment suggesting demethylation was occurring in the latter parts of the experiment.

Selenium, an element in Group 16, has important and complex organometallic chemistry, and is readily methylated. In organisms, Se has a large number of biochemical roles as it is incorporated into a number of enzymes [52]. The methylation of Se was first investigated by Challenger and since these early studies it has been shown that Se has an extensive bio‐ chemistry and that compounds, such as selenoproteins are important constituents of organ‐ isms as they act as anti-oxidants and have other roles in the cellular machinery [16, 81, 87]. Compounds such as selenomethionine and Se-adenosylselenomethionine (SeSAM), are found in cells, where Se has replaced S, as could be expected as these elements are from the same group of the periodic table. The methylating ability of SeSAM has been shown to be greater than that of SAM [7].

It has been suggested that CH3SeH and the cation CH3Se+ are some of the compounds re‐ sponsible for the toxicity of Se. Additionally, analogs to the methylated S-containing com‐ pounds (CH3)2S (DMS) and (CH3)2S2 have been identified ((CH3)2Se, (CH3)2SSe and (CH3)2Se2) [7]. It appears likely that these compounds are the degradation products pro‐ duced by certain microorganisms that exist in ocean waters, being formed from the decom‐ position of 3-dimethyl-selenopropionate ((CH3)2Se+ CH2CH2COO- ) (Figure 7) [7]. Similar processes likely account for their formation in terrestrial waters and other environments. The importance of this pathway was confirmed by growing a freshwater green algae on selenate in the absence of sulfate and showing the formation of the volatile methylated spe‐ cies [53]. This production was reduced when sulfate was added to the medium. Another study also demonstrated the release of volatile methylated Se compounds from green algae. This study also showed the potential for As to inhibit the formation of the Se compounds [55]. Overall, the formation of methylated Se compounds is relatively ubiquitous as they are apparently produced by both bacteria and algae.

(CH3)2Cd is relatively unstable in water. There have been few studies of organic Cd com‐ pounds in the environment in contrast to the inorganic chemistry of Cd that has been wellstudied. Initial evidence for the formation of CH3Cd+ was by bacterial cultures isolated from polar waters [91], and these same cultures also produced methylated Pb and Hg com‐ pounds. In the environment, evidence suggests that the peak concentration of these methy‐ lated species coincided with that of chlorophyll a, suggesting a microbial role in the formation of these compounds in polar waters. These studies of the formation of methylated Cd compounds in ocean waters require more confirmation. It is known that Cd can be abiot‐ ically methylated by methylcobalamin so this represents the potential biotic methylation

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For the other Group 13 elements, there is little evidence for the formation of methylated compounds except for Tl. This is a potentially toxic and bioaccumulative metal and the

methylated at the same time (i.e. likely via the Challenger mechanism) producing a mono-, di- or trimethylated product of, respectively, +2, +1 and 0 charge. This process occurs under anaerobic conditions with little evidence of aerobic methylation [82]. None of these com‐ pounds are highly stable but there is some initial evidence for the presence of these com‐ pounds at low pM concentrations in the Atlantic Ocean, mostly as (CH3)2Tl+ [92], and ranging up to nearly 50% of the total Tl, and present throughout the water column. The pro‐ file of the methylated species correlated with chlorophyll a in the upper waters, suggesting its microbial production. These authors also found methylation to occur in an anaerobic lake sediment [92]. There is no evidence for methylated In and Ga complexes in the environment.

Alkylation of Group 14 elements occurs for Sn and Pb and Ge. Alkyl Pb and Sn compounds have been widely produced for use in industrial and other applications, but these will not be discussed here. Studies have shown that a variety of microbes can methylate Sn [8]. For Ge, the mono-, di-, tri- and tetramethylated compounds have been found in the environment. In the ocean, GeIV has its highest concentrations in deep ocean waters with depleted concentra‐ tions in surface waters (Figure 3e) ([82] and references therein). For much of the ocean, the dominant form is monomethylated (CH3Ge3+), and its distribution suggests it is relatively

somewhat lower concentrations (Figure 3e). It is apparent that methylated Ge compounds can be produced under anaerobic conditions and that these species are not produced in oxic waters in the presence of algae. These compounds can also be made in the laboratory through reactions with CH3I and methylcobalamin, suggesting that this is the pathway for methylation in the environment, although it has also been suggested that these compounds

Besides As and Sb, bismuth (Bi) is found under some conditions as methylated compounds, although the methylation is restricted to prokaryotes [5, 82]. As the compounds of Bi are used in industrial and pharmaceutical applications, such as Pepto-Bismol, its presence in sewage treatment plants, and in municipal waste deposits, and the loss of volatile forms of Bi from these environments is not surprising [9]. Bismuth, in contrast to As and Sb, exists in environmental media as a +3 ion rather than as an oxyanion. It also can be found in the

is oxidized and

) occur in seawater at

chemistry of inorganic Tl has received some attention [7]. It appears that TlI

stable. The other methylated Ge species ((CH3)2Ge2+ and (CH3)3Ge+

are formed via the Challenger mechanism [7].

pathway in aquatic systems [7].

**Figure 7.** Pathways for the formation and decomposition of methylated selenium compounds in the environment

However, direct methylation is also possible. In freshwater environments, γ-Proteobacteria, such as *Pseudomonas* spp. have been identified as Se methylators. Additionally, a gene en‐ coding for the bacterial thiopurine methyltransferase has been shown to methylate selenite and (methyl)selenocysteine into dimethylselenide and dimethyldiselenide [88]. One poten‐ tial pathway of formation that interlinks the Se and Hg cycles relates to the potential for CH3Hg, present in the cell as a thiol complex (represented here as CH3HgR), to bind to Secontaining amino acids or thiols (represented here as SeR). It is proposed that two CH3HgSeR react to form (CH3Hg)2Se and R2Se, and then (CH3Hg)2Se decomposes to (CH3)2Hg and HgSe [89]. A similar pathway was proposed many years ago involving (CH3Hg)2S for the formation of (CH3)2Hg, which was purportedly formed in sediments [90]. Demethylation of (CH3)2Se is found in anoxic sediments and it is speculated that methano‐ gens are using this compound for growth in an analogous fashion to their utilization of DMS. In summary, it is likely that in sediments and other low oxygen/anoxic environments, direct methylation is occurring while in the water column the production of the methylated Se compounds results from decomposition of Se-containing biomolecules.

#### **3.4. The biotic formation of other methylated compounds**

A number of metal(loid)s not yet discussed are known to form organometallic compounds although there is little information on the formation, stability and toxicity of many of them, or on how they are formed in aquatic systems. In Group 12, Zn does not form any stable small methylated compounds in aquatic systems. In contrast, Cd, above Hg in the periodic table, has been isolated from the environment as methylated compounds [9], although (CH3)2Cd is relatively unstable in water. There have been few studies of organic Cd com‐ pounds in the environment in contrast to the inorganic chemistry of Cd that has been wellstudied. Initial evidence for the formation of CH3Cd+ was by bacterial cultures isolated from polar waters [91], and these same cultures also produced methylated Pb and Hg com‐ pounds. In the environment, evidence suggests that the peak concentration of these methy‐ lated species coincided with that of chlorophyll a, suggesting a microbial role in the formation of these compounds in polar waters. These studies of the formation of methylated Cd compounds in ocean waters require more confirmation. It is known that Cd can be abiot‐ ically methylated by methylcobalamin so this represents the potential biotic methylation pathway in aquatic systems [7].

[55]. Overall, the formation of methylated Se compounds is relatively ubiquitous as they are

**Figure 7.** Pathways for the formation and decomposition of methylated selenium compounds in the environment

Se compounds results from decomposition of Se-containing biomolecules.

**3.4. The biotic formation of other methylated compounds**

However, direct methylation is also possible. In freshwater environments, γ-Proteobacteria, such as *Pseudomonas* spp. have been identified as Se methylators. Additionally, a gene en‐ coding for the bacterial thiopurine methyltransferase has been shown to methylate selenite and (methyl)selenocysteine into dimethylselenide and dimethyldiselenide [88]. One poten‐ tial pathway of formation that interlinks the Se and Hg cycles relates to the potential for CH3Hg, present in the cell as a thiol complex (represented here as CH3HgR), to bind to Secontaining amino acids or thiols (represented here as SeR). It is proposed that two CH3HgSeR react to form (CH3Hg)2Se and R2Se, and then (CH3Hg)2Se decomposes to (CH3)2Hg and HgSe [89]. A similar pathway was proposed many years ago involving (CH3Hg)2S for the formation of (CH3)2Hg, which was purportedly formed in sediments [90]. Demethylation of (CH3)2Se is found in anoxic sediments and it is speculated that methano‐ gens are using this compound for growth in an analogous fashion to their utilization of DMS. In summary, it is likely that in sediments and other low oxygen/anoxic environments, direct methylation is occurring while in the water column the production of the methylated

A number of metal(loid)s not yet discussed are known to form organometallic compounds although there is little information on the formation, stability and toxicity of many of them, or on how they are formed in aquatic systems. In Group 12, Zn does not form any stable small methylated compounds in aquatic systems. In contrast, Cd, above Hg in the periodic table, has been isolated from the environment as methylated compounds [9], although

apparently produced by both bacteria and algae.

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For the other Group 13 elements, there is little evidence for the formation of methylated compounds except for Tl. This is a potentially toxic and bioaccumulative metal and the chemistry of inorganic Tl has received some attention [7]. It appears that TlI is oxidized and methylated at the same time (i.e. likely via the Challenger mechanism) producing a mono-, di- or trimethylated product of, respectively, +2, +1 and 0 charge. This process occurs under anaerobic conditions with little evidence of aerobic methylation [82]. None of these com‐ pounds are highly stable but there is some initial evidence for the presence of these com‐ pounds at low pM concentrations in the Atlantic Ocean, mostly as (CH3)2Tl+ [92], and ranging up to nearly 50% of the total Tl, and present throughout the water column. The pro‐ file of the methylated species correlated with chlorophyll a in the upper waters, suggesting its microbial production. These authors also found methylation to occur in an anaerobic lake sediment [92]. There is no evidence for methylated In and Ga complexes in the environment.

Alkylation of Group 14 elements occurs for Sn and Pb and Ge. Alkyl Pb and Sn compounds have been widely produced for use in industrial and other applications, but these will not be discussed here. Studies have shown that a variety of microbes can methylate Sn [8]. For Ge, the mono-, di-, tri- and tetramethylated compounds have been found in the environment. In the ocean, GeIV has its highest concentrations in deep ocean waters with depleted concentra‐ tions in surface waters (Figure 3e) ([82] and references therein). For much of the ocean, the dominant form is monomethylated (CH3Ge3+), and its distribution suggests it is relatively stable. The other methylated Ge species ((CH3)2Ge2+ and (CH3)3Ge+ ) occur in seawater at somewhat lower concentrations (Figure 3e). It is apparent that methylated Ge compounds can be produced under anaerobic conditions and that these species are not produced in oxic waters in the presence of algae. These compounds can also be made in the laboratory through reactions with CH3I and methylcobalamin, suggesting that this is the pathway for methylation in the environment, although it has also been suggested that these compounds are formed via the Challenger mechanism [7].

Besides As and Sb, bismuth (Bi) is found under some conditions as methylated compounds, although the methylation is restricted to prokaryotes [5, 82]. As the compounds of Bi are used in industrial and pharmaceutical applications, such as Pepto-Bismol, its presence in sewage treatment plants, and in municipal waste deposits, and the loss of volatile forms of Bi from these environments is not surprising [9]. Bismuth, in contrast to As and Sb, exists in environmental media as a +3 ion rather than as an oxyanion. It also can be found in the mono-, di- and tri- methylated form, and (CH3)3Bi is non-polar while the other forms are ions, which exist as complexes in solution (Table 1). The trimethylated form is less stable than its As and Sb analogs. For example, methylated Bi compounds are produced by metha‐ nogens in culture [7]. The hydride (BiH3) has also been isolated from bacterial cultures (Ta‐ ble 1). In the biotic formation of methylated Bi compounds, it is possible that the methyl group is donated by methylcobalamin, which is consistent with its form in solution as ionic complexes [5, 82]. This contrasts the methylation of As and Sb. However, there is little de‐ tailed information available about the exact nature of the methylation process for Bi.

at high CH3Hg and Hg concentrations it has been shown that a series of inducible genes can be activated (the *mer* operon) that can aid in detoxification of CH3Hg via demethyla‐ tion (the *mer B* gene which encodes for organomercury lyase), and reduction of HgII to Hg0 (the *mer A* gene which produces mercury reductase) (Figure 8). The *mer B* gene can decom‐ pose a variety of organomercury compounds. There is also a regulatory gene (*mer R*), as well as transport genes and their transport proteins in the cell membrane [23]. The overall operon is contained on a plasmid and is readily transferred between bacteria in the environ‐ ment. However, while membrane HgII transport proteins are present in bacteria with the *mer* operon, they are absent in SRB and therefore are not involved in the transport of Hg

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**Figure 8.** Representation of the *mer* operon and the processes whereby mercury species are transformed. Taken from

However, while methane and Hg0 are the primary products of *mer*-mediated Hg demethyla‐ tion, CO2 has also been observed as a major demethylation product in many studies [24, 94, 95] and this oxidative demethylation is not considered an active detoxification pathway. A variety of aerobes and anaerobes have been implicated in carrying out oxidative demethyla‐ tion which has been observed in freshwater, estuarine and alkaline-hypersaline sediments

Barkay et al. (2003) and used with permission from Elsevier.

associated with methylation.

There is evidence for the formation of organo-Te compounds and the mechanisms for their formation appear to be similar to the mechanisms for the formation of organic Se-containing compounds [82], which is also in Group 16 of the periodic table. Certain bacteria have been shown to methylate Te and form dimethylated compounds, and there is a complex interac‐ tion between the ability of these organisms to methylate and the Se concentration [8]. This suggests that Se and Te, which are electronically similar, behave similarly in this regard. Both exist in solution as oxyacids although the pKa's of the selenic and selenous acid are much lower than the corresponding values for telluric and tellurous acid [93]. There has been some suggestion that Po can be methylated but the conditions under which this occur‐ red suggests that these compounds are unlikely in natural environments [7]. The mechanism of methylation is not known and as Po exists in various oxidation states, there are a number of potential methylation pathways. More research is needed to examine in more detail the methylation and cycling of Po and the other less-studied heavy metals and metalloids.

#### **4. Microbially-Mediated Decomposition of Methylated Compounds**

The pathways for the decomposition of organometal(loid)s often occur in a stepwise fashion with the removal of successive methyl (or other alkyl) groups from the central metal(loid) atom. Examples include the decomposition of (CH3)2Hg to CH3Hg+ to HgII/Hg0 and (CH3)4Sn or (CH3)4Pb to the tri, di and monomethyl forms. Microbial demethylation is likely to be a detoxifying mechanism in many cases, but there is also evidence that some microbes can use the low molecular weight methyl compounds as a carbon source. Both of these pathways will be highlighted with specific examples. For most of the metal(loid)s, the degradation pathways are less studied than the methylation reactions. The one exception is the demethy‐ lation of CH3Hg using the *mer* operon [23] (Figure 8). This pathway can decompose other alkyl as well as phenyl Hg compounds.

A number of microbes appear to be important in the demethylation of CH3Hg although the mechanisms are not as well understood in many environments [23, 24]. In uncontaminat‐ ed environments, the major products appear to be Hg and CO2 and therefore this path‐ way has been termed *oxidative demethylation*, in contrast to *reductive demethylation*, where CH4 is the major carbon product [24]. The mechanism of oxidative demethylation may be anal‐ ogous to monomethylamine degradation by methanogens or acetate oxidation by SRB's. Reductive demethylation appears to be prevalent in more contaminated environments and at high CH3Hg and Hg concentrations it has been shown that a series of inducible genes can be activated (the *mer* operon) that can aid in detoxification of CH3Hg via demethyla‐ tion (the *mer B* gene which encodes for organomercury lyase), and reduction of HgII to Hg0 (the *mer A* gene which produces mercury reductase) (Figure 8). The *mer B* gene can decom‐ pose a variety of organomercury compounds. There is also a regulatory gene (*mer R*), as well as transport genes and their transport proteins in the cell membrane [23]. The overall operon is contained on a plasmid and is readily transferred between bacteria in the environ‐ ment. However, while membrane HgII transport proteins are present in bacteria with the *mer* operon, they are absent in SRB and therefore are not involved in the transport of Hg associated with methylation.

mono-, di- and tri- methylated form, and (CH3)3Bi is non-polar while the other forms are ions, which exist as complexes in solution (Table 1). The trimethylated form is less stable than its As and Sb analogs. For example, methylated Bi compounds are produced by metha‐ nogens in culture [7]. The hydride (BiH3) has also been isolated from bacterial cultures (Ta‐ ble 1). In the biotic formation of methylated Bi compounds, it is possible that the methyl group is donated by methylcobalamin, which is consistent with its form in solution as ionic complexes [5, 82]. This contrasts the methylation of As and Sb. However, there is little de‐

There is evidence for the formation of organo-Te compounds and the mechanisms for their formation appear to be similar to the mechanisms for the formation of organic Se-containing compounds [82], which is also in Group 16 of the periodic table. Certain bacteria have been shown to methylate Te and form dimethylated compounds, and there is a complex interac‐ tion between the ability of these organisms to methylate and the Se concentration [8]. This suggests that Se and Te, which are electronically similar, behave similarly in this regard. Both exist in solution as oxyacids although the pKa's of the selenic and selenous acid are much lower than the corresponding values for telluric and tellurous acid [93]. There has been some suggestion that Po can be methylated but the conditions under which this occur‐ red suggests that these compounds are unlikely in natural environments [7]. The mechanism of methylation is not known and as Po exists in various oxidation states, there are a number of potential methylation pathways. More research is needed to examine in more detail the methylation and cycling of Po and the other less-studied heavy metals and metalloids.

tailed information available about the exact nature of the methylation process for Bi.

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**4. Microbially-Mediated Decomposition of Methylated Compounds**

atom. Examples include the decomposition of (CH3)2Hg to CH3Hg+

alkyl as well as phenyl Hg compounds.

The pathways for the decomposition of organometal(loid)s often occur in a stepwise fashion with the removal of successive methyl (or other alkyl) groups from the central metal(loid)

or (CH3)4Pb to the tri, di and monomethyl forms. Microbial demethylation is likely to be a detoxifying mechanism in many cases, but there is also evidence that some microbes can use the low molecular weight methyl compounds as a carbon source. Both of these pathways will be highlighted with specific examples. For most of the metal(loid)s, the degradation pathways are less studied than the methylation reactions. The one exception is the demethy‐ lation of CH3Hg using the *mer* operon [23] (Figure 8). This pathway can decompose other

A number of microbes appear to be important in the demethylation of CH3Hg although the mechanisms are not as well understood in many environments [23, 24]. In uncontaminat‐ ed environments, the major products appear to be Hg and CO2 and therefore this path‐ way has been termed *oxidative demethylation*, in contrast to *reductive demethylation*, where CH4 is the major carbon product [24]. The mechanism of oxidative demethylation may be anal‐ ogous to monomethylamine degradation by methanogens or acetate oxidation by SRB's. Reductive demethylation appears to be prevalent in more contaminated environments and

to HgII/Hg0

and (CH3)4Sn

**Figure 8.** Representation of the *mer* operon and the processes whereby mercury species are transformed. Taken from Barkay et al. (2003) and used with permission from Elsevier.

However, while methane and Hg0 are the primary products of *mer*-mediated Hg demethyla‐ tion, CO2 has also been observed as a major demethylation product in many studies [24, 94, 95] and this oxidative demethylation is not considered an active detoxification pathway. A variety of aerobes and anaerobes have been implicated in carrying out oxidative demethyla‐ tion which has been observed in freshwater, estuarine and alkaline-hypersaline sediments [24, 94, 96]. However, the identity of the organisms responsible for oxidative demethylation in the environment remains poorly understood and no specific organism has been isolated. One study confirmed the ability of two sulfate reducing bacterial strains and one methano‐ gen strain to demethylate mercury in pure culture [97]. The authors however argued that the CO2 seen in these studies resulted from oxidation of methane released from CH3Hg after cleavage via organomercurial lyase and was actually a secondary product and not the pri‐ mary product of demethylation. However, this view is not universally accepted based on other their studies under both aerobic and anaerobic conditions [24]. Overall, the relative importance of *mer*-mediated versus oxidative demethylation is poorly understood. In sys‐ tems that are not highly contaminated, oxidative demethylation appears to dominate, under both aerobic and anaerobic conditions. The Hg concentrations that would cause a switch from one pathway to the other are only loosely defined. The end-product of oxidative deme‐ thylation has been presumed to be Hg(II), but that has not been confirmed in most studies.

containing compounds such as methyliodide (CH3I). For example, experiments examining the potential for the abiotic methylation of AsIII by CH3I found that the reaction proceeded, but only at very high pH's (>10), above those typically found in the environment [100]. Mon‐ omethylarsenic was the only product formed. This study therefore suggests that abiotic for‐ mation of methylated As is not likely in the environment. Other studies have shown that CH3I can methylate Pb, Sn and Ge [8], and there are also studies showing the methylation of

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 by CH3I, but not with HgII [101]. Again, these studies do not suggest that these reactions are important in the environment. For example, the rate of formation of CH3Hg from Hg0

the presence of CH3I, given typical environmental concentrations of these species, is insuffi‐ cient to account for any substantial portion of the CH3Hg found in natural waters. For Hg, the same conclusion is reached in terms of methyl transfer reactions between methylated tin compounds and HgII, even though it appeared that the presence of Cl and high pH, which would be found in seawater, enhanced the reaction rate [101]. Overall, the results of a num‐ ber of studies over time [52, 101-104] lead to the conclusion that abiotic formation of CH3Hg

There have been a number of studies that have shown the potential for the transfer of a methyl group from an organic compound to HgII, and the formation of CH3Hg in environ‐ mental waters [102, 104, 105], including precipitation [106]. However, most of these experi‐ ments have been conducted at unrealistic concentrations of both Hg and the organic compound, as well as having a ratio of Hg/organic matter much greater than found in the environment. Additionally, the reactions are often done at low pH or high temperature. Clearly, these experiments show that CH3Hg can be manufactured abiotically, which is no surprise, but the results of these studies have little environmental relevance. For example, in the laboratory, CH3Hg is routinely manufactured through the reaction with cobalamin. This

In terms of abiotic decomposition, the stability of organometal(loid)s in water is related to the polarity of the metal carbon bond with more polarity enhancing hydrolysis and decom‐ position [107]. However, while many organometal(loid)s may be thermodynamically unsta‐ ble, they are often kinetically stable and are not degraded abiotically as readily as may be expected. For example, CH3Hg which is less stable in water at low pH but the reaction is kinetically hindered [108]. Photochemical processes however can enhance the degrada‐ tion rate. This is an important loss process for methylated Hg. Dimethylmercury is much less stable to photochemical degradation than CH3Hg [109]. In the ocean, the rate of CH3Hg photodecomposition varies, with some studies showing relative rapid rates of degrada‐ tion, while others have shown little degradation [110-112]. It is likely that complexation to

or NOM in seawater impacts this rate, but these effects have been little studied com‐ pared to in freshwater systems [111, 113, 114]. The degradation of CH3Hg in freshwater has received more study and photochemical oxidants and UV radiation are important in driving the degradation [114-117] with various reactive oxygen species implicated in the reactions [113, 118]. For the methylated Se compounds, it is likely that oxidation occurs in the presence of light, especially UV radiation, as has been found for methylated S com‐

does not however suggest that this is occurring abiotically in the environment.

in the environment by these pathways is not important.

in

291

Hg0

Cl-

pounds (e.g. (CH3)2S) [29, 119].

Studies in freshwater and marine sediments [24, 42, 67, 94, 98] however confirm that the rate of demethylation is rapid, and that the rate constant for this process is higher than that of methylation, and that demethylation occurs across the redox gradient. Many of these studies have used stable isotope approaches, where isotopically-labelled inorganic Hg, and CH3Hg made using a different isotope of Hg, are spiked into sediments or water and the transfor‐ mations of each followed under the same conditions. Another approach is to use radiolabel‐ led Hg and 14C-labeled CH3Hg but these approaches cannot be done at the trace levels of the stable isotope method. There is an obvious advantage of simultaneously examining both of the reactions in the same experiment. These studies [42, 67, 98] have shown overall that the fraction of Hg as CH3Hg in sediments is often closely related to the ratio of the rate con‐ stants, which suggests that both reactions are pseudo first order and that the system reaches steady state relatively rapidly. Given that demethylation rate constants on the order of 10 d-1 have been measured in some sediment, the time to steady state is a few days.

Biodegradation studies of other methylated compounds are limited but these are likely to occur in sediments and reduced environments [82, 99]. One study examining the degrada‐ tion of methylated As compounds in a lake showed that the decomposition occurred in the presence of suspended particulate but not in filtered water, implicating microbial processes in the decomposition. Degradation occurred in the dark under anaerobic conditions. Studies of alkylated Sn compounds have similarly shown their degradation in the sediment and it is likely that anaerobic microbes can demethylate most of the commonly found methylated metal(loid)s in the environment.

#### **5. Abiotic Formation and Degradation Pathways in Aquatic Systems**

A variety of methyl donors exist in environmental solutions and these have the potential to methylate the metal(loid)s discussed here. Pathways include the following primary mecha‐ nisms: 1) cross-methylation i.e. transfer of a methyl group from one metal(loid) compound to another e.g. (CH3)4Pb + HgII -> (CH3)3Pb+ + CH3Hg+ ; and 2) methylation by other methylcontaining compounds such as methyliodide (CH3I). For example, experiments examining the potential for the abiotic methylation of AsIII by CH3I found that the reaction proceeded, but only at very high pH's (>10), above those typically found in the environment [100]. Mon‐ omethylarsenic was the only product formed. This study therefore suggests that abiotic for‐ mation of methylated As is not likely in the environment. Other studies have shown that CH3I can methylate Pb, Sn and Ge [8], and there are also studies showing the methylation of Hg0 by CH3I, but not with HgII [101]. Again, these studies do not suggest that these reactions are important in the environment. For example, the rate of formation of CH3Hg from Hg0 in the presence of CH3I, given typical environmental concentrations of these species, is insuffi‐ cient to account for any substantial portion of the CH3Hg found in natural waters. For Hg, the same conclusion is reached in terms of methyl transfer reactions between methylated tin compounds and HgII, even though it appeared that the presence of Cl and high pH, which would be found in seawater, enhanced the reaction rate [101]. Overall, the results of a num‐ ber of studies over time [52, 101-104] lead to the conclusion that abiotic formation of CH3Hg in the environment by these pathways is not important.

[24, 94, 96]. However, the identity of the organisms responsible for oxidative demethylation in the environment remains poorly understood and no specific organism has been isolated. One study confirmed the ability of two sulfate reducing bacterial strains and one methano‐ gen strain to demethylate mercury in pure culture [97]. The authors however argued that the CO2 seen in these studies resulted from oxidation of methane released from CH3Hg after cleavage via organomercurial lyase and was actually a secondary product and not the pri‐ mary product of demethylation. However, this view is not universally accepted based on other their studies under both aerobic and anaerobic conditions [24]. Overall, the relative importance of *mer*-mediated versus oxidative demethylation is poorly understood. In sys‐ tems that are not highly contaminated, oxidative demethylation appears to dominate, under both aerobic and anaerobic conditions. The Hg concentrations that would cause a switch from one pathway to the other are only loosely defined. The end-product of oxidative deme‐ thylation has been presumed to be Hg(II), but that has not been confirmed in most studies.

290 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Studies in freshwater and marine sediments [24, 42, 67, 94, 98] however confirm that the rate of demethylation is rapid, and that the rate constant for this process is higher than that of methylation, and that demethylation occurs across the redox gradient. Many of these studies have used stable isotope approaches, where isotopically-labelled inorganic Hg, and CH3Hg made using a different isotope of Hg, are spiked into sediments or water and the transfor‐ mations of each followed under the same conditions. Another approach is to use radiolabel‐ led Hg and 14C-labeled CH3Hg but these approaches cannot be done at the trace levels of the stable isotope method. There is an obvious advantage of simultaneously examining both of the reactions in the same experiment. These studies [42, 67, 98] have shown overall that the fraction of Hg as CH3Hg in sediments is often closely related to the ratio of the rate con‐ stants, which suggests that both reactions are pseudo first order and that the system reaches steady state relatively rapidly. Given that demethylation rate constants on the order of 10 d-1

Biodegradation studies of other methylated compounds are limited but these are likely to occur in sediments and reduced environments [82, 99]. One study examining the degrada‐ tion of methylated As compounds in a lake showed that the decomposition occurred in the presence of suspended particulate but not in filtered water, implicating microbial processes in the decomposition. Degradation occurred in the dark under anaerobic conditions. Studies of alkylated Sn compounds have similarly shown their degradation in the sediment and it is likely that anaerobic microbes can demethylate most of the commonly found methylated

**5. Abiotic Formation and Degradation Pathways in Aquatic Systems**

A variety of methyl donors exist in environmental solutions and these have the potential to methylate the metal(loid)s discussed here. Pathways include the following primary mecha‐ nisms: 1) cross-methylation i.e. transfer of a methyl group from one metal(loid) compound

+ CH3Hg+

; and 2) methylation by other methyl-

have been measured in some sediment, the time to steady state is a few days.

metal(loid)s in the environment.

to another e.g. (CH3)4Pb + HgII -> (CH3)3Pb+

There have been a number of studies that have shown the potential for the transfer of a methyl group from an organic compound to HgII, and the formation of CH3Hg in environ‐ mental waters [102, 104, 105], including precipitation [106]. However, most of these experi‐ ments have been conducted at unrealistic concentrations of both Hg and the organic compound, as well as having a ratio of Hg/organic matter much greater than found in the environment. Additionally, the reactions are often done at low pH or high temperature. Clearly, these experiments show that CH3Hg can be manufactured abiotically, which is no surprise, but the results of these studies have little environmental relevance. For example, in the laboratory, CH3Hg is routinely manufactured through the reaction with cobalamin. This does not however suggest that this is occurring abiotically in the environment.

In terms of abiotic decomposition, the stability of organometal(loid)s in water is related to the polarity of the metal carbon bond with more polarity enhancing hydrolysis and decom‐ position [107]. However, while many organometal(loid)s may be thermodynamically unsta‐ ble, they are often kinetically stable and are not degraded abiotically as readily as may be expected. For example, CH3Hg which is less stable in water at low pH but the reaction is kinetically hindered [108]. Photochemical processes however can enhance the degrada‐ tion rate. This is an important loss process for methylated Hg. Dimethylmercury is much less stable to photochemical degradation than CH3Hg [109]. In the ocean, the rate of CH3Hg photodecomposition varies, with some studies showing relative rapid rates of degrada‐ tion, while others have shown little degradation [110-112]. It is likely that complexation to Clor NOM in seawater impacts this rate, but these effects have been little studied com‐ pared to in freshwater systems [111, 113, 114]. The degradation of CH3Hg in freshwater has received more study and photochemical oxidants and UV radiation are important in driving the degradation [114-117] with various reactive oxygen species implicated in the reactions [113, 118]. For the methylated Se compounds, it is likely that oxidation occurs in the presence of light, especially UV radiation, as has been found for methylated S com‐ pounds (e.g. (CH3)2S) [29, 119].

A photochemical degradation study will tetraethyllead showed that first order decrease of the reactant with the subsequent buildup and decay of the intermediates [120]. The final product was PbII. It is likely that tetramethyllead would be degraded in a similar fashion. For Se, photodecomposition of selenoamino acids can produce significant amounts of vola‐ tile selenium species in both light and dark conditions in the laboratory, with (CH3)2SSe and (CH3)2Se2 being the major products, with small amounts of (CH3)2Se being formed [29]. Inor‐ ganic selenium oxyanions did not produce any volatile products. It was hypothesized that formation of H2O2 under the experimental conditions initiated the decomposition reactions. Overall, it is likely that multi-methylated species can be photochemically decomposed by the stepwise removal of methyl groups. This is discussed above and is true for the decompo‐ sition of (CH3)2Hg.

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#### **6. Conclusion**

The methylation of metal(loid)s in the environment is important to the fate and transport of many of the elements in Groups 12-16 of the periodic table. This is true for elements besides those that have received the most attention (Hg, As and Se). Much of the research done on these three elements can be extrapolated to the other elements based on knowledge about the chemistry of the elements in environmental waters and the uptake and fate of the ele‐ ments within cells. While there is still more research needed to fully understand the methyl‐ ation, demethylation and fate and transport of Hg, As and Se, this is even more necessary for the other metal(loid)s discussed in this chapter.

#### **Acknowledgements**

Acknowledgements The support of NSF and NIH, especially grant P42 ES007373 from NIEHS, is acknowledged for support of studies and work cited and in supporting the chap‐ ter prepartion. The contents of this chapter are solely the responsibility of the author and do not represent the official views of NIH.

#### **Author details**

Robert P. Mason\*

Address all correspondence to: robert.mason@uconn.edu

Departments of Marine Sciences and Chemistry, University of Connecticut, USA

#### **References**

A photochemical degradation study will tetraethyllead showed that first order decrease of the reactant with the subsequent buildup and decay of the intermediates [120]. The final product was PbII. It is likely that tetramethyllead would be degraded in a similar fashion. For Se, photodecomposition of selenoamino acids can produce significant amounts of vola‐ tile selenium species in both light and dark conditions in the laboratory, with (CH3)2SSe and (CH3)2Se2 being the major products, with small amounts of (CH3)2Se being formed [29]. Inor‐ ganic selenium oxyanions did not produce any volatile products. It was hypothesized that formation of H2O2 under the experimental conditions initiated the decomposition reactions. Overall, it is likely that multi-methylated species can be photochemically decomposed by the stepwise removal of methyl groups. This is discussed above and is true for the decompo‐

The methylation of metal(loid)s in the environment is important to the fate and transport of many of the elements in Groups 12-16 of the periodic table. This is true for elements besides those that have received the most attention (Hg, As and Se). Much of the research done on these three elements can be extrapolated to the other elements based on knowledge about the chemistry of the elements in environmental waters and the uptake and fate of the ele‐ ments within cells. While there is still more research needed to fully understand the methyl‐ ation, demethylation and fate and transport of Hg, As and Se, this is even more necessary

Acknowledgements The support of NSF and NIH, especially grant P42 ES007373 from NIEHS, is acknowledged for support of studies and work cited and in supporting the chap‐ ter prepartion. The contents of this chapter are solely the responsibility of the author and do

sition of (CH3)2Hg.

**6. Conclusion**

**Acknowledgements**

**Author details**

Robert P. Mason\*

not represent the official views of NIH.

for the other metal(loid)s discussed in this chapter.

292 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment

Address all correspondence to: robert.mason@uconn.edu

Departments of Marine Sciences and Chemistry, University of Connecticut, USA


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300 Methylation - From DNA, RNA and Histones to Diseases and Treatment Methylation - From DNA, RNA and Histones to Diseases and Treatment


### *Edited by Anica Dricu*

Alterations in the normal DNA methylation processes are known to have major consequences for embryonic development and are associated with congenital defects, autoimmunity, aging and malignant transformation. The main purpose of this book is to provide information about the importance of methylation mechanisms in human health and disease. The book, covers the basic mechanism of DNA and protein methylation, aiming at the advanced undergraduate and graduate biomedical students and researchers working in the epigenetic area. The textbook chapters provide background as well as advanced information in the methylation area. On the other hand, it provide readers with both classical and relevant recent discoveries that have been made in the field, pointing out pathways where questions remain.

Methylation - From DNA, RNA and Histones to Diseases and Treatment

Methylation

From DNA, RNA and Histones

to Diseases and Treatment

*Edited by Anica Dricu*

Photo by farakos / iStock