**Chromatin Damage Patterns Shift According to Eu/ Heterochromatin Replication**

María Vittoria Di Tomaso, Pablo Liddle, Laura Lafon-Hughes, Ana Laura Reyes-Ábalos and Gustavo Folle

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51847

### **1. Introduction**

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350 The Mechanisms of DNA Replication

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In order to maintain genetic stability, strictly controlled mechanisms are essential to assure the accuracy of genetic functions. Precise genome replication and correct control of gene ex‐ pression mostly *via* epigenetic mechanisms are critical in maintaining the stability of ge‐ nomes. Moreover, the characteristic chromatin compartmentalization of mammalian genomes contributes to regulate the housekeeping or tissue-specific genetic activities [1, 2].

Table 1 summarizes the distinct chromatin compartments and their foremost properties. Eu‐ chromatin (*eu*: true) and heterochromatin (*hetero*: different) are two major compartments or chromatin states of the DNA originally distinguished by their isopycnotic or heteropycnotic interphase staining properties, respectively [3]. The heterochromatin compartment differen‐ tiates in both constitutive (permanent) and facultative (developmentally reorganized) states [4]. Facultative heterochromatin represents chromatin regions being facultatively inactivated (heterochromatinized) because of gene dosage compensation (i.e.: mammalian female inac‐ tive X chromosome) randomly silenced at an early stage of embryogenesis or tissue-specific gene expression. Constitutive heterochromatin consists in regions of α- and β-heterochro‐ matin [5, 6].

Distinct features characterize the different chromatin states (Table 1). Interphase open chro‐ matin conformation and transcriptional activity in all cell types distinguish euchromatin. Higher order chromatin compaction characterizes constitutive α- and β-heterochromatic re‐ gions while gene silencing differentiates constitutive α-heterochromatin. Tissue-specific transcriptional activity and low or high chromatin condensation, depending on gene expres‐

© 2013 Di Tomaso 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 Di Tomaso et al.; licensee InTech. This is a paper 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.

sion, correspond to features of facultative heterochromatin [7, 6]. The mammalian genome compartmentalization can be visualized in both banded metaphase chromosomes and stained interphase nuclei.



The C-banding procedure [8] produces a selective staining of specific chromosome regions, mapping at or adjacent to centromeres, telomeres or interstitial arm sites, depending on the species. Occasionally, a chromosome arm is entirely heterochromatic, such as the long arm of the Chinese hamster X chromosome (Figure 1, left). In humans, C-bands are located at centromeres and pericentric regions of all chromosomes, being conspicuous at the pericen‐ tric regions of chromosomes 1, 9 and 16 and the distal long arm of the Y chromosome (Yq) (Figure 1, right).

sion, correspond to features of facultative heterochromatin [7, 6]. The mammalian genome compartmentalization can be visualized in both banded metaphase chromosomes and

**Compartments Euchromatin Facultative heterochromatin Constitutive heterochromatin**

Peripheral compartment and chromocenters

> High order compaction

Dosage inactivated

Gene activity in euchromatic state until silencing

GC-rich AT-rich GC- and AT-rich AT-rich AT-rich

genes No genes

methylated Methylated Methylated Methylated

hypoacetylated Hypoacetylated Hypoacetylated Hypoacetylated

compensation α- heterochromatin β-heterochromatin

chromosome (Xi) C-bands C-bands

Peripheral compartment and chromocenters

> High order compaction

No gene activity

Tandem highly repeated DNA sequences

Peripheral compartment and chromocenters

> High order compaction

Transposable elements and heterochromatic genes

Low gene activity in heterochromatic state

> Tandem highly repeated DNA sequences

**Chromatin types** Euchromatin Tissue-specific Dosage

Open conformation Low or high order

**genes** Housekeeping genes Inactivated tissue-

Gene activity in euchromatic state in all cells

**methylation** Unmethylated Unmethylated or

**acetylation** Hyperacetylated Hyperacetilated or

**Table 1.** Distinguishing properties of chromatin compartments.

Light G-bands Dark G-bands Inactive X

Peripheral compartment and chromocenters

compaction

specific genes

Tissue-specific gene activity in euchromatic state

SINEs LINEs SINEs and LINEs

**timing** Early Early or late Late Late Late or early

The C-banding procedure [8] produces a selective staining of specific chromosome regions, mapping at or adjacent to centromeres, telomeres or interstitial arm sites, depending on the species. Occasionally, a chromosome arm is entirely heterochromatic, such as the long arm of the Chinese hamster X chromosome (Figure 1, left). In humans, C-bands are located at centromeres and pericentric regions of all chromosomes, being conspicuous at the pericen‐ tric regions of chromosomes 1, 9 and 16 and the distal long arm of the Y chromosome (Yq)

stained interphase nuclei.

352 The Mechanisms of DNA Replication

**Location in metaphase chromosomes**

**Location in**

**Interphase chromatin compaction**

**Presence of**

**Gene expression and relation to chromatin state**

**GC or AT DNA sequences richness**

**Repeated DNA sequences**

**CpG island**

**Core histone tail**

**Replication**

(Figure 1, right).

**interphase nuclei** Inner compartment

**Figure 1.** C-banding in CHO and human chromosomes. Left: C-banded metaphase of CHO9 cell line. The CHO cell line was established from a Chinese hamster ovary fibroblast culture [9] and presents a modal number of 21 chromo‐ somes. This cell line contains eight normal and twelve rearranged autosomes with only one X chromosome. Giemsastained C-band regions are visualized in yellow (reflected light microscopy). The CHO X chromosome (X) shows an almost entirely heterochromatic long arm. Right: C-banded caryotype of a human peripheral lymphocyte metaphase showing centromeric, pericentric (chromosomes 1, 9 and 16) and distal Yq heterochromatic blocks.

By digestion with the proteolytic enzyme trypsin followed by Giemsa staining (G-banding procedure) [10], a pattern of alternate light and dark regions along the length of all chromo‐ somes is obtained (light G-bands and dark G-bands, respectively). The G-band pattern is characteristic for each chromosome pair allowing their precise identification and caryotyp‐ ing. Figure 2 shows the CHO9 and human G-band chromosome patterns.

**Figure 2.** G-banded CHO9 metaphase (left) and a male human peripheral lymphocyte caryotype exhibiting G-bands (right).

C- and G-band patterns reveal the heterogeneous organization of chromatin along con‐ densed chromosomes. C-bands enclose constitutive α- and β-heterochromatin. Regions with ubiquitously expressed housekeeping genes (euchromatin) reside in light G-bands, while tissue-specific genes (facultative heterochromatin) dwell in dark G-bands [5, 6, 11 ].

Light and dark G-bands may reflect a differential array of SAR (**S**caffold-**A**ssociated **R**egions), composed by highly AT-rich DNA stretches binding to the chromosome scaffold. Regions of dark G-bands exhibit a tighter chromatin fiber coiling than light G-bands do‐ mains [12]. Constitutive heterochromatin has an even more dense conformation.

Moreover, euchromatic light G-bands are GC-rich and gene-dense regions, containing un‐ methylated CpG islands and moderately repeated **S**hort **In**terspersed **E**lements (SINE), mainly represented by Alu family sequences. Conversely, facultative heterochromatic dark G-bands are AT-rich, gene-poor and harbor hypermethylated CpG and moderately repeated family of Long Interspersed Elements (LINE) sequences. Constitutive α-heterochromatic Cbands are the major locations of tandem non-coding highly repeated satellite DNA sequen‐ ces, devoid of genes [11, 13]. However, constitutive β-heterochromatin presents inserted middle-repetitive transposable elements between the tandem repeats, some of them tran‐ scriptionally active [6]. Moreover, genes residing within regions of pericentric constitutive β-heterochromatin termed "heterochromatic genes" have been reported in *Drosophila,* mam‐ mals and plants [14, 15].

In spite of variations according to cell type or function of mammalian interphase nuclei, the corresponding chromatin of light and dark G-bands as well as C-bands is non-randomly dis‐ tributed in different nuclear compartments, displaying specific chromatin conformation, molecular composition and gene expression patterns.

In most interphase cells, euchromatin (light G-bands) dwells in the inner compartment of nu‐ clei, whereas heterochromatin (dark G-bands and C-bands) resides in the peripheral compart‐ ment, chromocenters and around nucleoli [6, 16]. Figure 3 illustrates a HeLa nucleus where the different interphase chromatin compartments can be recognized.

**Figure 3.** Distinct eu/heterochromatin compartments in DAPI-stained HeLa interphase nucleus. DAPI-bright regions correspond to heterochromatin and dim areas to euchromatin. N: nucleolus.

Constitutive and tissue-specific genes are only expressed in the euchromatic state. There‐ fore, facultative heterochromatin behaves as euchromatin in cells where its tissue-specific genes are transcribed, but holds a packed (heterochromatic) conformation when genes re‐ main silent.

ubiquitously expressed housekeeping genes (euchromatin) reside in light G-bands, while

Light and dark G-bands may reflect a differential array of SAR (**S**caffold-**A**ssociated **R**egions), composed by highly AT-rich DNA stretches binding to the chromosome scaffold. Regions of dark G-bands exhibit a tighter chromatin fiber coiling than light G-bands do‐

Moreover, euchromatic light G-bands are GC-rich and gene-dense regions, containing un‐ methylated CpG islands and moderately repeated **S**hort **In**terspersed **E**lements (SINE), mainly represented by Alu family sequences. Conversely, facultative heterochromatic dark G-bands are AT-rich, gene-poor and harbor hypermethylated CpG and moderately repeated family of Long Interspersed Elements (LINE) sequences. Constitutive α-heterochromatic Cbands are the major locations of tandem non-coding highly repeated satellite DNA sequen‐ ces, devoid of genes [11, 13]. However, constitutive β-heterochromatin presents inserted middle-repetitive transposable elements between the tandem repeats, some of them tran‐ scriptionally active [6]. Moreover, genes residing within regions of pericentric constitutive β-heterochromatin termed "heterochromatic genes" have been reported in *Drosophila,* mam‐

In spite of variations according to cell type or function of mammalian interphase nuclei, the corresponding chromatin of light and dark G-bands as well as C-bands is non-randomly dis‐ tributed in different nuclear compartments, displaying specific chromatin conformation,

In most interphase cells, euchromatin (light G-bands) dwells in the inner compartment of nu‐ clei, whereas heterochromatin (dark G-bands and C-bands) resides in the peripheral compart‐ ment, chromocenters and around nucleoli [6, 16]. Figure 3 illustrates a HeLa nucleus where the

**Figure 3.** Distinct eu/heterochromatin compartments in DAPI-stained HeLa interphase nucleus. DAPI-bright regions

tissue-specific genes (facultative heterochromatin) dwell in dark G-bands [5, 6, 11 ].

mains [12]. Constitutive heterochromatin has an even more dense conformation.

mals and plants [14, 15].

354 The Mechanisms of DNA Replication

molecular composition and gene expression patterns.

different interphase chromatin compartments can be recognized.

correspond to heterochromatin and dim areas to euchromatin. N: nucleolus.

However, some transposons and heterochromatic genes of β-heterochromatin are transcrip‐ tionally active in heterochromatic state suggesting that distinct epigenetic mechanisms of gene regulation and preservation of eu/heterochromatic states may exist in these regions [6, 14, 15].

Once acquired, the chromatin states are somatically maintained as stable heritable epigenet‐ ic states. Euchromatin remodels during mitosis and restores the original organization in ear‐ ly G1 phase of each cell cycle. In addition, during DNA synthesis (S-phase) both euchromatin and heterocromatin transiently lose their typical condensation status recover‐ ing the previous folding level after replication. Establishment and maintenance of chromatin states involve post-translational modification enzymes that act coordinately to methylate CpG islands and to either acetylate, methylate, phosphorylate, ubiquitinate, poly-ADP ribo‐ sylate or SUMOylate the core histone tails of nucleosomes. These epigenetic changes, togeth‐ er with the recruitment of methyl-CpG binding proteins, ATP-dependent chromatin remodeling complexes and the association of specific non-histone proteins, such as HP1 (**H**eterochromatin **P**rotein 1) or RNAi (non-coding interference RNA), also mediate the regu‐ lation of DNA replication, transcription and repair [17, 18].

The N- and C-termini of H3 and H4 core histones are particularly involved in epigenetic regulation. Acetyl groups covalently added to lysines, serines or arginines of the N-terminal histone tails reduce the affinity to DNA, promoting the accessibility of chromatin remodel‐ ing and activating transcription factors. Therefore, histone hyperacetylation usually charac‐ terizes active chromatin regions. Conversely, transcriptionally silenced chromatin regions generally contain hypoacetylated histones (Table 1). For instance, H3 acetylated (ac) in ly‐ sine 9 (H3K9ac) is enriched at the promoter region of active genes although, it was reported that the histone H3 acetylated at lysine 4 (H3K4ac) resides in pericentric heterochromatin of *Schizosaccharomyces pombe,* playing a role in the assembly of repressive heterochromatin [19]. On the other hand, histone methylation (me) can be associated with transcriptional activa‐ tion or repression. For example, methylation of H3 on lysines 4, 36 or 79 (H3K4me, H3K36me, H3K79me) is associated with transcriptional activation whereas methylation of H3 on lysines 9 or 27 (H3K9me, H3K27me) and of H4 on lysine 20 (H4K20me) is involved in transcriptional repression [18]. The concerted action of acetylated and methylated histone core residues is central in creating a "histone code" which delineates distinct genomic loci that recruit factors needed for DNA remodeling, transcription, replication and repair [5, 17].

In general, methylation of CpG islands within 5'regions of genes is associated with hypoace‐ thylated histones, characterizing the heterochromatic state (Table 1). However, DNA meth‐ ylation is not exclusively related to gene silencing. It was reported that methylation of some imprinting centers can displace trans-acting repressor factors, allowing the expression of the linked imprinted genes [20].

The epigenetic mechanisms involved in the maintenance of eu/heterochromatic compart‐ ments and gene expression are connected to DNA replication. There are specific interactions between components of the replication machinery and chromatin related factors, timing the eu- or heterochromatin replication.

### **2. Replication of eu/heterochromatin compartments**

Compartmentalization of vertebrate genomes cooperates in achieving the high fidelity DNA replication necessary for the accurate preservation of the genetic information throughout cell generations. DNA replication is a temporarily and spatially highly ordered and strictly regulated process, occurring during S-phase of the cell cycle, with distinct genome compart‐ ments replicating at different times. The replication timing of the genome compartments are highly conserved within consecutive cell cycles and regulated by specific epigenetic chroma‐ tin conformation domains, DNA features and transcriptional activity [21, 22, 23].

Mammalian chromosome duplication involves clusters or domains of neighboring replicons named **R**eplication **T**iming **D**omains (RTD) which synchronously start and end replication, according to a deterministic replication timing program [21, 22, 24]. When one domain com‐ pletes replication, an adjacent domain successively initiates DNA synthesis [25]. Remarka‐ bly, mouse and human asynchronous replication timing may function randomly between individual replicons within a RTD and non-randomly between RTD [25]. The random firing of replication origins within a RTD generates a different replication pattern during each Sphase, but it has been reported that some origins fire preferentially and more frequently than others [26]. The RTD are stable structures of mammalian interphase nuclei, replicating and transcribing in temporal and spatial coordination [26].

Pulse labeled interphase nuclei of human, mouse and hamster cells with the base analogues 5-bromo-2'-deoxyuridine (BrdU) or 5-ethynyl-2′-deoxiuridine (EdU) demonstrated the asyn‐ chrony and specific spatial distribution of DNA replication. The early replication pattern of S-phase (ES-phase) is characterized by replication foci dispersed throughout the inner envi‐ ronment of the nuclei with scarce or absence of foci at the periphery or adjacent to the nucle‐ oli. The replication pattern changes throughout the progression of S-phase. In mid S-phase (MS-phase) most foci map adjacent to the internal nuclear membrane and around nucleoli, with few foci centrally located. Lastly, late S-phase replication maps next to the nuclear en‐ velope as well as in chromocenters and around nucleoli [16, 27]. Early S-phase and late Sphase replication patterns of CHO9 cells are illustrated in Figure 4.

In general, chromatin with transcriptional activity (euchromatin) replicates early in S-phase whereas constitutive α-heterochromatin duplicates late. Besides, facultative heterochroma‐ tin replicates earlier if its tissue-specific genes are being expressed and later if not [6, 28] (Table 1). It has been reported that genes of mouse embryonic stem cells residing within GCrich and LINE-poor DNA (euchromatin) do not modify their replication timing after differ‐ entiation to neural precursors, whereas genes residing in AT-rich and LINE-rich DNA revealed changes in replication timing accompanied by changes in gene expression and chromatin folding [29]. A change of replication timing from early S to late S-phase is particu‐ larly evident in the female mammalian Xi [30]

The epigenetic mechanisms involved in the maintenance of eu/heterochromatic compart‐ ments and gene expression are connected to DNA replication. There are specific interactions between components of the replication machinery and chromatin related factors, timing the

Compartmentalization of vertebrate genomes cooperates in achieving the high fidelity DNA replication necessary for the accurate preservation of the genetic information throughout cell generations. DNA replication is a temporarily and spatially highly ordered and strictly regulated process, occurring during S-phase of the cell cycle, with distinct genome compart‐ ments replicating at different times. The replication timing of the genome compartments are highly conserved within consecutive cell cycles and regulated by specific epigenetic chroma‐

Mammalian chromosome duplication involves clusters or domains of neighboring replicons named **R**eplication **T**iming **D**omains (RTD) which synchronously start and end replication, according to a deterministic replication timing program [21, 22, 24]. When one domain com‐ pletes replication, an adjacent domain successively initiates DNA synthesis [25]. Remarka‐ bly, mouse and human asynchronous replication timing may function randomly between individual replicons within a RTD and non-randomly between RTD [25]. The random firing of replication origins within a RTD generates a different replication pattern during each Sphase, but it has been reported that some origins fire preferentially and more frequently than others [26]. The RTD are stable structures of mammalian interphase nuclei, replicating

Pulse labeled interphase nuclei of human, mouse and hamster cells with the base analogues 5-bromo-2'-deoxyuridine (BrdU) or 5-ethynyl-2′-deoxiuridine (EdU) demonstrated the asyn‐ chrony and specific spatial distribution of DNA replication. The early replication pattern of S-phase (ES-phase) is characterized by replication foci dispersed throughout the inner envi‐ ronment of the nuclei with scarce or absence of foci at the periphery or adjacent to the nucle‐ oli. The replication pattern changes throughout the progression of S-phase. In mid S-phase (MS-phase) most foci map adjacent to the internal nuclear membrane and around nucleoli, with few foci centrally located. Lastly, late S-phase replication maps next to the nuclear en‐ velope as well as in chromocenters and around nucleoli [16, 27]. Early S-phase and late S-

In general, chromatin with transcriptional activity (euchromatin) replicates early in S-phase whereas constitutive α-heterochromatin duplicates late. Besides, facultative heterochroma‐ tin replicates earlier if its tissue-specific genes are being expressed and later if not [6, 28] (Table 1). It has been reported that genes of mouse embryonic stem cells residing within GCrich and LINE-poor DNA (euchromatin) do not modify their replication timing after differ‐ entiation to neural precursors, whereas genes residing in AT-rich and LINE-rich DNA revealed changes in replication timing accompanied by changes in gene expression and

tin conformation domains, DNA features and transcriptional activity [21, 22, 23].

eu- or heterochromatin replication.

356 The Mechanisms of DNA Replication

**2. Replication of eu/heterochromatin compartments**

and transcribing in temporal and spatial coordination [26].

phase replication patterns of CHO9 cells are illustrated in Figure 4.

**Figure 4.** Early (ES-phase) or late (LS-phase) replication patterns of CHO9 nuclei revealed by incorporation of EdU and subsequent detection with an Alexa Fluor 488 (green) conjugated azide (Click-iT EdU imaging kit, Invitrogen). (a) ESphase nucleus with inner compartment replication. (b) LS-phase nucleus showing replication in the peripheral com‐ partment, chromocenters and around nucleoli.

Early replication seems to be important but not essential for gene transcription. Moreover, late replication is not an obligatory feature of heterochromatin. For example, transcriptional‐ ly active transposons of β-heterochromatin replicate late while the heterochromatic centro‐ meres and the silent mating-type cassettes of *Schizosaccharomyces pombe* replicate in early S phase [14, 15, 31]. There are additional cases reported of early heterochromatin replication such as human telomeres [32]. and mouse pericentric heterochromatin and centromeres [33].

The early replicon clusters of higher eukaryotes alternate their replication and transcription activity. However, correlation between replication and transcription does not exist in S*ac‐ charomyces cerevisiae* [34]. Employing distinct colored fluorescent labels to recognize early replication foci and transcription foci (factories), it was shown that both labels do not coloc‐ alize. In ES-phase, actively replicating foci are transcriptionally inactive and only restart transcription after finishing replication. The replication timing is indirectly related to tran‐ scription through the assembly of a higher-order chromatin state [2]. For example, silencing of the mammalian Xi is initially reversible and only stabilizes when an identifiable higherorder chromatin configuration (Barr body) appears and replication is delayed [35].

The chromatin replication timing is reestablished early in G1 phase of each cell cycle, coinci‐ dent with the anchorage and positioning of chromosomal segments at specific locations within the nucleus named TDP (**T**iming **D**ecision **P**oint) [36]. Both anchorage and position‐ ing of chromosomes are central in the organization of nuclear eu/heterochromatic compart‐ ments and the establishment of replication timing and transcriptional activity [23, 36]. Modifications in subnuclear chromatin organization are associated with changes in replica‐ tion timing during development [37]. For example, the position of the immunoglobulin heavy chain locus (IgH) in B cells shows that its localization in the interphase nuclei de‐ pends on replication timing and gene activity. During early stages of B cell differentiation, both transcribed alleles of the IgH locus are centrally located in the nucleus and replicate early. Conversely, in advanced differentiation stages the IgH locus is repositioned to the nu‐ clear periphery, repressed and late-replicated [38].

Nonetheless, chromatin positioning at the nuclear periphery is indicative but not mandatory for gene silencing and late replication. In fact, the nuclear periphery is heterogeneous with respect to transcription. For instance, in budding yeast, nuclear pores, which mediate the transport between the nucleus and cytoplasm, enhance the transcriptional activity of genes positioned in their proximity [39]. The dosage compensation complex of the hyperacetylated *Drosophila* male X chromosome interacts with nuclear pore proteins determining its tran‐ scription up-regulation and early DNA duplication [40].

Replication clusters correspond to bands of metaphase chromosomes. Tightly coiled C-band (constitutive heterochromatin) replicates in late S-phase. Facultative heterochromatin of the dark G-bands duplicates either early or late depending on its tissue-specific expression. Ear‐ ly replication pattern characterizes the loosely coiled euchromatin of light G-bands. Ubiqui‐ tously expressed housekeeping genes (light G-bands) are therefore early replicating [41, 42]. Duplication timing analysis by quantitative PCR of the boundary region between G-light 13q14.3 and G-dark 13q21.1 bands showed that the G-light side of the frontier replicates ear‐ ly whilst the G-dark interface replicates late. However, analysis using PCR primers spaced at approximately 150 Kb intervals showed that the switch in G-light/G-dark band replication timing takes place gradually from early-mid to late S-phase over a 1-2 Mb region [43]. The DNA segments corresponding to large regions between early and late-S phase replication timing domains are termed TTR (Timing Transition Regions) [44].

A correlation between replication timing and epigenetic modification of chromatin has also been shown. Early replication domains are related to specific combination of changes in his‐ tone lysine residues (H3K9Ac, H3K27Ac, H3K4me, H3K36me and H3K79me) associated with transcriptional activity. On the other hand, the repressive epigenetic modifications (H3K9me, H3K27me and H4K20me) are linked to late replication [18].

Chromatin epigenetic changes occurring throughout DNA replication may provide a repli‐ cation timing mechanism (firing early or late replication origins) in the direction of main‐ taining specific chromatin expression patterns [45]. It was reported that histone hypoacetylation is needed to preserve normal heterochromatin replication dynamics [46] and that histone hyperacetylation may increase the efficiency of replication origins, advanc‐ ing the replication timing of distinct genomic regions [47]. For instance, removal of acetyl groups by HDAC (**H**istone **D**e**AC**etylase) contributes to mantain late replication at imprint‐ ed loci [48] and the generation of neocentromeres [49].

Several proteins, including CpG island-methylating DNMT (**DN**A **M**ethyl **T**ransferase), core histone tail-methylating HMT (**H**istone **M**ethyl **T**ransferase) and HP1 (**H**eterochromatin-as‐ sociated **P**rotein), colocalize with late replicating DNA regions [45]. HP1 binds to hetero‐ chromatin, facilitating the extension of the repressive H3K9me modification [50] and hence delaying replication timing by supporting heterochromatin conformation. HP1 could facili‐ tate the late firing of replication origins within heterochromatin [51]. Furthermore, muta‐ tions of DNMT result in earlier replication of normally late replicating DNA. For instance, patients with mutations in the Dnmt3b gene (coding protein DNMT3b) have hypomethylat‐ ed CpG islands in the Xi chromosome, which replicates at an earlier S-phase stage despite the presence of XIST (**X**-**I**nactive **S**pecific **T**ranscript) RNA [52]. Accordingly, changes in ei‐ ther DNA or histone methylation status in concert with histone acetylation patterns may promote open or tight chromatin conformations and thus modifications in the firing of repli‐ cation origins and/or replication rates [47].

both transcribed alleles of the IgH locus are centrally located in the nucleus and replicate early. Conversely, in advanced differentiation stages the IgH locus is repositioned to the nu‐

Nonetheless, chromatin positioning at the nuclear periphery is indicative but not mandatory for gene silencing and late replication. In fact, the nuclear periphery is heterogeneous with respect to transcription. For instance, in budding yeast, nuclear pores, which mediate the transport between the nucleus and cytoplasm, enhance the transcriptional activity of genes positioned in their proximity [39]. The dosage compensation complex of the hyperacetylated *Drosophila* male X chromosome interacts with nuclear pore proteins determining its tran‐

Replication clusters correspond to bands of metaphase chromosomes. Tightly coiled C-band (constitutive heterochromatin) replicates in late S-phase. Facultative heterochromatin of the dark G-bands duplicates either early or late depending on its tissue-specific expression. Ear‐ ly replication pattern characterizes the loosely coiled euchromatin of light G-bands. Ubiqui‐ tously expressed housekeeping genes (light G-bands) are therefore early replicating [41, 42]. Duplication timing analysis by quantitative PCR of the boundary region between G-light 13q14.3 and G-dark 13q21.1 bands showed that the G-light side of the frontier replicates ear‐ ly whilst the G-dark interface replicates late. However, analysis using PCR primers spaced at approximately 150 Kb intervals showed that the switch in G-light/G-dark band replication timing takes place gradually from early-mid to late S-phase over a 1-2 Mb region [43]. The DNA segments corresponding to large regions between early and late-S phase replication

A correlation between replication timing and epigenetic modification of chromatin has also been shown. Early replication domains are related to specific combination of changes in his‐ tone lysine residues (H3K9Ac, H3K27Ac, H3K4me, H3K36me and H3K79me) associated with transcriptional activity. On the other hand, the repressive epigenetic modifications

Chromatin epigenetic changes occurring throughout DNA replication may provide a repli‐ cation timing mechanism (firing early or late replication origins) in the direction of main‐ taining specific chromatin expression patterns [45]. It was reported that histone hypoacetylation is needed to preserve normal heterochromatin replication dynamics [46] and that histone hyperacetylation may increase the efficiency of replication origins, advanc‐ ing the replication timing of distinct genomic regions [47]. For instance, removal of acetyl groups by HDAC (**H**istone **D**e**AC**etylase) contributes to mantain late replication at imprint‐

Several proteins, including CpG island-methylating DNMT (**DN**A **M**ethyl **T**ransferase), core histone tail-methylating HMT (**H**istone **M**ethyl **T**ransferase) and HP1 (**H**eterochromatin-as‐ sociated **P**rotein), colocalize with late replicating DNA regions [45]. HP1 binds to hetero‐ chromatin, facilitating the extension of the repressive H3K9me modification [50] and hence delaying replication timing by supporting heterochromatin conformation. HP1 could facili‐ tate the late firing of replication origins within heterochromatin [51]. Furthermore, muta‐

clear periphery, repressed and late-replicated [38].

358 The Mechanisms of DNA Replication

scription up-regulation and early DNA duplication [40].

timing domains are termed TTR (Timing Transition Regions) [44].

(H3K9me, H3K27me and H4K20me) are linked to late replication [18].

ed loci [48] and the generation of neocentromeres [49].

In mammals, several distinct discrete or diffuse genomic sequence motifs can potentially act as **O**rigin **R**eplication **I**dentification (ORI), where a large number of proteins bind to load replication complexes. A protein complex, named the pre-**R**eplication **C**omplex (pre-RC) associates with potential replication origins in G1 phase. This complex includes the **O**rigin **R**ecognition **C**omplex (ORC), which recognizes the replication origins, the heli‐ case MCM2-7 (**M**ini **C**hromosome **M**aintenance 2-7), and other essential factors. Early fir‐ ing ORI demonstrated to be rich in MCM proteins. Besides, MCM could be more efficient in early firing than in late firing ORI suggesting that heterochromatin could re‐ press MCM activities [53, 54].

Accessibility of replication initiation factors to redundant or discrete replication origins may be regulated by its nuclear localization in relation to chromatin states. For example, the early replicating α-globin locus is located within a light G-band. Deletions that juxtapose the αglobin locus next to a region of late replicating telomeric condensed heterochromatin (repo‐ sitioning this locus to the nuclear periphery), delay the initiation of α-globin replication by restricting the access of initiation factors to the ORI [55].

There is a complex cell cycle intra-S checkpoint involving the ATR/CHK1-related network in metazoas and ATR/Rad53 in *Saccharomyces cerevisiae* that controls replication asynchrony. The transition from early to late S-phase replication (mid-S replication pause) is coupled with the activation of the intra S-phase checkpoint at mid S-phase which inhibits the initia‐ tion of late replicons. It has been reported that inhibition of CHK1 generates earlier firing of a late-firing subset of ORI [56, 57]. Accordingly, the checkpoint function may play a role in regulating replication asynchrony and S-phase progression [25, 58].

Both DNA and histone methylation can affect replication timing *via* the ATR/CHK1 con‐ trol pathway. There is a complex and so far not completely understood relationship be‐ tween checkpoint function and epigenetic modifications (DNA methylation, histone methylation and histone acetylation) in the regulation of replication origins firing during S-phase [47, 59].

Following pre-RC loading to ORI, a protein pre-**I**nitiation **C**omplex (pre-IC) assembles upon MCM proteins together with factors required for loading replicative polymerase. The chro‐ matin association of pre-RC and pre-IC is asynchronous, allowing pre-RC inhibition and pre-IC activation (from S-phase initiation toward the end of mitosis) by the cell cycle CDK proteins (**C**yclin-**D**ependent **K**inases). This regulation licenses replication to occur at a spe‐ cific time, only once per cell cycle, and ensures that cell cycle cannot progress until check‐ points are satisfied. In *Xenopus laevis* and mammalian cells there is an additional system to control licensing by means of the geminin protein, which also inhibits pre-RC. Degradation of geminin at the end of mitosis is essential for a new license of replication [56, 60].

Completion of replication is necessary for entire chromosome condensation. *Drosophila* ORC mutants unable to complete S-phase have defects not only in DNA replication (with some euchromatic regions replicating even later than heterochromatin) but also in cell cycle pro‐ gression and chromatin condensation [61]. Although some levels of chromosome condensa‐ tion occur in the absence of a complete replication cycle, mitotic chromosomes are shorter and thicker than in wild type *Drosophila*. Even though ORC is principally involved in the ini‐ tiation of DNA replication, additional roles in mitotic chromosome condensation, centro‐ mere function as well as the establishment and maintenance of gene silencing and heterochromatin have been suggested [61, 62, 63].

### **3. Eu/heterochromatin replication and distribution of genetic damage**

The S-phase of the cell cycle has proved to be very sensitive to genetic damage. S-phase has been considered as one of the sources of genomic instability. There are several lines of evi‐ dence that correlate genomic instability with chromosomal aberrations (CA), birth defects and infertility [64]. Besides, oncogene activation or tumor suppressor gene repression can arise as a consequence of primary DNA damage or CA [65]. Several authors have reported the colocalization of induced CA breakpoints (BP) (sites of chromosomal breaks in a CA) with regions harboring fragile sites, oncogenes or cancer-associated CA [66-72].

The human genome holds long stretches of AT-rich sequences as well as inverted, mirror or direct tandem repeats, prone to be arranged in unusual DNA secondary structures that may inhibit replication. The presence of secondary structures, unstable single-stranded or non-re‐ plicated regions could lead to chromosome fragility expressed as gaps or breaks in meta‐ phase chromosomes [73, 74].

DNA replication in mammals slows down significantly when the 1-2 Mb regions of TTR are replicated [57]. It was reported that after replication of euchromatic light G-bands, the repli‐ cation fork stalls at TTR of the interband regions, restarting DNA synthesis at the adjacent dark G-band after a mid S-phase pause [6]. This interband region devoid of replication ori‐ gins is often replicated by means of a single replication fork [75]. Such genomic segments could generate damage-prone regions that frequently overlap with DNA fragile sites [43, 76]. For example, the common fragile site FRA3B is devoid of replication origins and thus completes replication very late in S-phase [77]. In addition, it was observed that mutation rates increase with the distance from replication origins [78, 79].

Furthermore, it was reported an increase in mutation rate as S-phase advances. Early replicating housekeeping genes are more conserved than later replicating tissue-specific genes [57, 80]. Genes corresponding to mutational hot spots involved in speciation and adaptive radiation response are late replicating [57]. CpG methylation status of late repli‐ cating regions may contribute to the rise in mutation rate mostly due to 5meCpG substitu‐ tions [81, 82].

#### **3.1. Eu/heterochromatin replication and induced-damage distribution in a mitotic chromosome model**

control licensing by means of the geminin protein, which also inhibits pre-RC. Degradation

Completion of replication is necessary for entire chromosome condensation. *Drosophila* ORC mutants unable to complete S-phase have defects not only in DNA replication (with some euchromatic regions replicating even later than heterochromatin) but also in cell cycle pro‐ gression and chromatin condensation [61]. Although some levels of chromosome condensa‐ tion occur in the absence of a complete replication cycle, mitotic chromosomes are shorter and thicker than in wild type *Drosophila*. Even though ORC is principally involved in the ini‐ tiation of DNA replication, additional roles in mitotic chromosome condensation, centro‐ mere function as well as the establishment and maintenance of gene silencing and

**3. Eu/heterochromatin replication and distribution of genetic damage**

with regions harboring fragile sites, oncogenes or cancer-associated CA [66-72].

rates increase with the distance from replication origins [78, 79].

The S-phase of the cell cycle has proved to be very sensitive to genetic damage. S-phase has been considered as one of the sources of genomic instability. There are several lines of evi‐ dence that correlate genomic instability with chromosomal aberrations (CA), birth defects and infertility [64]. Besides, oncogene activation or tumor suppressor gene repression can arise as a consequence of primary DNA damage or CA [65]. Several authors have reported the colocalization of induced CA breakpoints (BP) (sites of chromosomal breaks in a CA)

The human genome holds long stretches of AT-rich sequences as well as inverted, mirror or direct tandem repeats, prone to be arranged in unusual DNA secondary structures that may inhibit replication. The presence of secondary structures, unstable single-stranded or non-re‐ plicated regions could lead to chromosome fragility expressed as gaps or breaks in meta‐

DNA replication in mammals slows down significantly when the 1-2 Mb regions of TTR are replicated [57]. It was reported that after replication of euchromatic light G-bands, the repli‐ cation fork stalls at TTR of the interband regions, restarting DNA synthesis at the adjacent dark G-band after a mid S-phase pause [6]. This interband region devoid of replication ori‐ gins is often replicated by means of a single replication fork [75]. Such genomic segments could generate damage-prone regions that frequently overlap with DNA fragile sites [43, 76]. For example, the common fragile site FRA3B is devoid of replication origins and thus completes replication very late in S-phase [77]. In addition, it was observed that mutation

Furthermore, it was reported an increase in mutation rate as S-phase advances. Early replicating housekeeping genes are more conserved than later replicating tissue-specific genes [57, 80]. Genes corresponding to mutational hot spots involved in speciation and adaptive radiation response are late replicating [57]. CpG methylation status of late repli‐ cating regions may contribute to the rise in mutation rate mostly due to 5meCpG substitu‐

of geminin at the end of mitosis is essential for a new license of replication [56, 60].

heterochromatin have been suggested [61, 62, 63].

phase chromosomes [73, 74].

360 The Mechanisms of DNA Replication

tions [81, 82].

DNA lesions trigger a **D**NA **D**amage **R**esponse (DDR) characterized by activation of cell cy‐ cle checkpoints, damage sensor proteins, DNA repair mechanisms and apoptotic pathways [83, 84]. The DNA **D**ouble-**S**trand **B**reak (DSB) is the critical DNA lesion involved in CA production [85]. DSB can be generated by DNA-damaging agents or spontaneously through the endogenous production of reactive oxygen species (ROS) or cellular processes such as DNA replication, repair, transposition or mitotic recombination. Agents inducing DSB and CA are named clastogens. The S-phase independent clastogens, like ionizing radiation and the radiomimetic agent bleomycin, directly induce DSB. Conversely, S-phase dependent clastogens such as UV-C and alkylating compounds need the intervention of DNA repair and replication in order to generate DSB, which could ultimately lead to CA. Hence, DNA replication constitutes a relevant step in the transformation of DNA lesions into CA. Be‐ sides, some clastogenic agents such as the anti-topoisomerase II cleavable complex trappers behave as S-phase independent clastogens. Eukaryotic topoisomerases II alleviate tensional DNA stress by the generation of a DNA topoisomerase II complex (cleavable complex) with‐ in which the topoisomerase II component introduces transient breaks in both DNA strands (DSB) allowing the DNA to pass through the breaks [86]. Drugs that act by trapping cleava‐ ble complexes hamper the resealing of DSB produced by topoisomerase II and, as a conse‐ quence, DNA DSB persist [87, 88].

As shown in Figure 1, the CHO9 X-chromosome exhibits an almost entire constitutive heter‐ ochromatic long arm (Xq) with the exception of a medial secondary constriction. Besides, Xq replicates in late S-phase whereas the euchromatin of the short arm (Xp) and the Xq secon‐ dary constriction duplicates during early S-phase (Figure 5) [89, 90]. Differential replication timing of Xp and Xq of CHO cells provided a valuable experimental model to analyze the relationship between eu/heterochromatin DNA replication and CA induced by different types of clastogens: UV-C light, the methylating agent methylmethane sulphonate (MMS) and the anti-topoisomerase II inhibitor etoposide (a cleavable complex trapper) in BrdU pulse-labeled CHO9 chromosomes [91, 92].

CHO9 cells were treated with MMS (20 mM) or etoposide (20 μM) and simultaneously ex‐ posed to 30 mM BrdU (40 min) or otherwise exposed to UV-C (30 J/m2; 0.1 J/m2/s) and im‐ mediately labeled with BrdU (40 min). Incorporation of BrdU in Xp or Xq was disclosed by immunolabeling either treated or control CHO9 metaphases with anti-BrdU antibodies cou‐ pled to FITC. The relationship between replication timing, chromatin conformation and ge‐ netic damage was investigated by mapping induced BP in Xp and Xq in cells treated both in early and late S-phase [91, 92].

Examples of CA induced by MMS, etoposide and UV-C in replicating CHO9 Xp or Xq are shown in Figure 5. Figure 6 illustrates Xp/Xq distribution of etoposide, UV-C and MMS-in‐ duced BP in relation to replication.

The application of χ<sup>2</sup> test to analyze the association between Xp/Xq replication pattern and Xp/Xq BP localization showed that when Xp replicates, BP produced by either MMS, UV-C or etoposide clustered in Xp. On the other hand, during Xq replication, BP induced by the clastogens concentrated in Xq [91, 92] (Figure 6).

**Figure 5.** Illustrations of CA involving CHO9 Xp or Xq induced by (a) MMS, (b) etoposide, or (c) UV-C in (a) early (Xp replication) or (b and c) late (Xq replication) S-phase. Different types of CA are shown: (a) symmetric quadrirradial af‐ fecting Xp; (b) asymmetric quadrirradial with acentric fragment involving Xq; (c) duplication-deletion in Xq (arrow). Chromosomes exhibit BrdU immunolabeling (yellow) and either PI (red) or DAPI (blue) counterstaining.

**Figure 6.** Bar diagram illustrating CHO9 X chromosome BP distribution induced by etoposide (ETO, 20 μM), methyl‐ methane sulphonate (MMS, 20 mM) and UV-C (30 J/m2; 0.1 J/m2/s) in Xp (grey) and Xq (blue) during early (ES-phase) and late (LS-phase) cell cycle phases. The bar on the left side (E) indicates the expected frequencies of induced BP ac‐ cording to Xp and Xq relative length.

Since UV-C and MMS are S-phase dependent clastogens, the observed predominance of BP produced in Xp or Xq according to replication timing could be explained based on their re‐ quirement of DNA synthesis to produce CA. DNA base damage induced by MMS as well as cyclobutane pyrimidine dimers (CPD) and 6-4 photoproducts (6-4 PP) produced by UV-C are preferentially repaired through **B**ase **E**xcision **R**epair (BER) and **N**ucleotide **E**xcision **R**epair (NER) mechanisms, respectively. Both repair systems create an excision repair **S**in‐ gle-**S**trand **B**reak (SSB) intermediate at the site of DNA lesion which is then filled by DNA repair synthesis [93]. If DNA replication initiates with an excision repair SSB intermediate, another SSB can be generated in the complementary DNA strand, thus forming a DSB [94, 95]. Additionally, CPD, 6-4 PP or base damage in a single strand (unrepaired before DNA replication) may stall the replication fork and as a result, may produce a SSB in the opposite DNA strand [96, 97]. Furthermore, two nearby SSB in each DNA strand may behave as a DSB [98]. The DSB generated could be ultimately processed and transformed in CA [91, 92].

or etoposide clustered in Xp. On the other hand, during Xq replication, BP induced by the

**Figure 5.** Illustrations of CA involving CHO9 Xp or Xq induced by (a) MMS, (b) etoposide, or (c) UV-C in (a) early (Xp replication) or (b and c) late (Xq replication) S-phase. Different types of CA are shown: (a) symmetric quadrirradial af‐ fecting Xp; (b) asymmetric quadrirradial with acentric fragment involving Xq; (c) duplication-deletion in Xq (arrow).

**Figure 6.** Bar diagram illustrating CHO9 X chromosome BP distribution induced by etoposide (ETO, 20 μM), methyl‐ methane sulphonate (MMS, 20 mM) and UV-C (30 J/m2; 0.1 J/m2/s) in Xp (grey) and Xq (blue) during early (ES-phase) and late (LS-phase) cell cycle phases. The bar on the left side (E) indicates the expected frequencies of induced BP ac‐

Since UV-C and MMS are S-phase dependent clastogens, the observed predominance of BP produced in Xp or Xq according to replication timing could be explained based on their re‐ quirement of DNA synthesis to produce CA. DNA base damage induced by MMS as well as cyclobutane pyrimidine dimers (CPD) and 6-4 photoproducts (6-4 PP) produced by UV-C are preferentially repaired through **B**ase **E**xcision **R**epair (BER) and **N**ucleotide **E**xcision **R**epair (NER) mechanisms, respectively. Both repair systems create an excision repair **S**in‐ gle-**S**trand **B**reak (SSB) intermediate at the site of DNA lesion which is then filled by DNA

Chromosomes exhibit BrdU immunolabeling (yellow) and either PI (red) or DAPI (blue) counterstaining.

clastogens concentrated in Xq [91, 92] (Figure 6).

362 The Mechanisms of DNA Replication

cording to Xp and Xq relative length.

Nonetheless, the preferential location of CA in replicating Xp or Xq during etoposide treat‐ ment (independently of its eu/heterochromatic states) may occur due to the inhibition of topoisomerase II activity during DNA synthesis [87, 88]. The local unraveling and subse‐ quent rewinding of eu or heterochromatin regions undergoing replication require topoiso‐ merase II activities to alleviate DNA torsional stress [86]. Etoposide stabilizes DNAtopoisomerase II cleavable complex and hinders the resealing of DSB introduced by the enzyme generating the accumulation of DSB unable to reach resolution. In addition, chro‐ matin unwinding during replication may turn DNA more accessible to S-independent and S-dependent chemical agents including etoposide and MMS, respectively [91, 92].

#### **3.2. Eu/heterochromatin replication and primary induced-damage distribution in interphase nuclei**

Few minutes after exposure of mammalian cells to DSB-inducing agents, the nucleosomal histone variant H2AX is phosphorylated at serine 139 (humans) or 129 (mouse) of C-termi‐ nal tails reaching a peak of phosphorylation 30 min later. H2AX phosphorylation (named γH2AX) initiates around the induced DSB and spreads through a large chromatin region (~2000 H2AX molecules) flanking the lesion, which can be visualized as discrete γH2AX foci in interphase nuclei and mitotic chromosomes by means of specific fluorochrome-conjugat‐ ed antibodies [99].

γH2AX is involved in the DDR by coordination with other damage response proteins to re‐ cruit signaling, remodeling, checkpoint and repair proteins. At sites of DSB, the DNA-PK (**DN**A **D**ependent **P**rotein **K**inase) binds to activate the **N**on **H**omologous **E**nd **J**oining (NHEJ) DSB repair pathway. If DSB are produced after replication, RAD51 and BRCA2 are recruited to DSB sites initiating the **H**omologous **R**ecombination repair pathway (HR). Si‐ multaneously, the sensing complex MRN (MRE11, RAD50, NBS1) associates to DSB, facili‐ tating the recruitment and activation (auto-phosphorylation) of ATM (**A**taxia **T**elangiectasia **M**utated), MDC1, BRCA1 and 53BP1 [100].

ATM, ATR (**AT**M- and **R**ad3-related) and DNA-PK are members of the phosphatidylinositol 3-kinase-like family of serine/threonine protein kinases that phosphorylate H2AX. Unlike ATM, which appears to be mainly activated by DSB, ATR seems to be activated by induced SSB and the excision repair SSB intermediates generated during DNA repair. Since ATR ac‐ tivation was observed in replicating cells, it was suggested that the blockage of replication forks by SSB is required to initiate ATR-mediated phosphorylation of H2AX. Besides, it was reported that stalled replication forks may also trigger H2AX phosphorylation when bulky lesions (i.e.: CPD and 6-4 PP) collide with replication forks [101, 102].

NBS1, MDC1, 53BP1, and BRCA1 may all function as mediators and amplifiers of the DDR, recruiting diverse repair and checkpoint proteins (including ATM and ATR) and generating an amplification loop that also extends H2AX phosphorylation [99]. 53BP1 can bind directly to H3K79me and H4K20me accumulated at sites of DSB collaborating with a global chroma‐ tin unwinding following the formation of DSB in concert with other proteins like TIP60 (member of an histone acetyltransferases family) and KAP1 [103, 104, 105].

Several immunofluorescence studies have demonstrated that induced-γH2AX foci are locat‐ ed preferentially within euchromatic regions of the genome, suggesting that heterochroma‐ tin could be refractory to γH2AX foci formation. Employing immuno-FISH to analyze radiation induced-DSB (γH2AX foci) in chromatin regions with known chromatin compac‐ tion (human chromosome 18 versus chromosome 19; RIDGE versus anti-RIDGE region of human chromosome 11), it has been observed that condensed regions of gene-poor chroma‐ tin are less susceptible to DSB induction compared with decondensed, gene-rich chromatin [106-109].

Different hypothesis have been raised to explain the non-homogeneous distribution of γH2AX foci in nuclei. The highly condensed state or abundance of binding proteins may re‐ duce the accesibility of chemical DNA damaging agents to heterochromatin. Besides, since condensed chromatin is less hydrated than euchromatin, a lower amount of free radicals could be induced by radiation [110]. Furthermore, compact heterochromatin could contain a lower proportion of H2AX isoform or be less accessible to kinases due to compaction or pro‐ tein coating [106]. Additionally, a wave of chromatin unwinding starting at DSB sites and spreading throughout the entire chromatin was described (as a result of KAP1 phosphoryla‐ tion by ATM kinase) implying that the preferential location of γH2AX foci in decondensed chromatin perhaps reflects chromatin reorganization [105, 111-113].

Finally, a short-range migration of DSB from packed chromatin toward specific decon‐ densed DSB repair domains could also take place [106, 110]. Using carbon ion microirradia‐ tion to induce DSB combined to a modified TUNEL assay to directly visualize these lesions and γH2AX immunodetection, a bending of the linear ion-induced γH2AX track around heterochromatic regions was observed [114]. The γH2AX foci migration from the interior to the periphery of heterochromatin appears to initiate within 20 min post-irradiation and be almost complete 1 h after damage induction. The decondensation of heterochromatin at sites of ion hits possibly promotes the movement of DSB to peripheral regions of lower chroma‐ tin density where repair may potentially proceed [114].

To assess the influence of replication in the distribution of chromatin damage, we analyzed the localization of bleomycin-induced γH2AX foci in relation to replication of eu- or hetero‐ chromatin interphase compartments in 5-ethynyl-2'deoxiuridine (EdU) pulsed-labeled CHO9 nuclei. Bleomycin (BLM) is a radiomimetic S-independent clastogen that induces oxi‐ dative damage, SSB and mainly DSB as well as a rapid phosphorylation of H2AX [115].

Asynchronously growing CHO9 cultures were pulse-exposed (30 min) to EdU (controls) or simultaneously (30 min) treated with BLM (40 μg/ml). Early and late replication regions and γH2AX foci were detected with an azide conjugated to Alexa Fluor 488 (Click-iT EdU, Invi‐ trogen) and mouse anti-γH2AX (Abcam) followed by Cy3-conjugated antimouse antibodies, respectively. Single-cell z-stacks from control (n=25) and treated (n=63) nuclei were obtained by confocal microscopy and processed with Image J software. Using binary masks for each channel, the relation (ratio) between the percentage of damaged (γH2AX) area in replicating chromatin (EdU) area and the percentage of damaged area in the whole nuclear area (DAPI) was calculated for each nucleus. Finally, the arithmetic mean of the ratios corresponding to early S (n=30) and late S (n=33) nuclei was calculated.

NBS1, MDC1, 53BP1, and BRCA1 may all function as mediators and amplifiers of the DDR, recruiting diverse repair and checkpoint proteins (including ATM and ATR) and generating an amplification loop that also extends H2AX phosphorylation [99]. 53BP1 can bind directly to H3K79me and H4K20me accumulated at sites of DSB collaborating with a global chroma‐ tin unwinding following the formation of DSB in concert with other proteins like TIP60

Several immunofluorescence studies have demonstrated that induced-γH2AX foci are locat‐ ed preferentially within euchromatic regions of the genome, suggesting that heterochroma‐ tin could be refractory to γH2AX foci formation. Employing immuno-FISH to analyze radiation induced-DSB (γH2AX foci) in chromatin regions with known chromatin compac‐ tion (human chromosome 18 versus chromosome 19; RIDGE versus anti-RIDGE region of human chromosome 11), it has been observed that condensed regions of gene-poor chroma‐ tin are less susceptible to DSB induction compared with decondensed, gene-rich chromatin

Different hypothesis have been raised to explain the non-homogeneous distribution of γH2AX foci in nuclei. The highly condensed state or abundance of binding proteins may re‐ duce the accesibility of chemical DNA damaging agents to heterochromatin. Besides, since condensed chromatin is less hydrated than euchromatin, a lower amount of free radicals could be induced by radiation [110]. Furthermore, compact heterochromatin could contain a lower proportion of H2AX isoform or be less accessible to kinases due to compaction or pro‐ tein coating [106]. Additionally, a wave of chromatin unwinding starting at DSB sites and spreading throughout the entire chromatin was described (as a result of KAP1 phosphoryla‐ tion by ATM kinase) implying that the preferential location of γH2AX foci in decondensed

Finally, a short-range migration of DSB from packed chromatin toward specific decon‐ densed DSB repair domains could also take place [106, 110]. Using carbon ion microirradia‐ tion to induce DSB combined to a modified TUNEL assay to directly visualize these lesions and γH2AX immunodetection, a bending of the linear ion-induced γH2AX track around heterochromatic regions was observed [114]. The γH2AX foci migration from the interior to the periphery of heterochromatin appears to initiate within 20 min post-irradiation and be almost complete 1 h after damage induction. The decondensation of heterochromatin at sites of ion hits possibly promotes the movement of DSB to peripheral regions of lower chroma‐

To assess the influence of replication in the distribution of chromatin damage, we analyzed the localization of bleomycin-induced γH2AX foci in relation to replication of eu- or hetero‐ chromatin interphase compartments in 5-ethynyl-2'deoxiuridine (EdU) pulsed-labeled CHO9 nuclei. Bleomycin (BLM) is a radiomimetic S-independent clastogen that induces oxi‐ dative damage, SSB and mainly DSB as well as a rapid phosphorylation of H2AX [115].

Asynchronously growing CHO9 cultures were pulse-exposed (30 min) to EdU (controls) or simultaneously (30 min) treated with BLM (40 μg/ml). Early and late replication regions and γH2AX foci were detected with an azide conjugated to Alexa Fluor 488 (Click-iT EdU, Invi‐

(member of an histone acetyltransferases family) and KAP1 [103, 104, 105].

chromatin perhaps reflects chromatin reorganization [105, 111-113].

tin density where repair may potentially proceed [114].

[106-109].

364 The Mechanisms of DNA Replication

**Figure 7.** Distribution of BLM induced-γH2AX foci revealed by immunolabelling (Cy3; red) in early (top) or late (bot‐ tom) S-phase CHO9 nuclei. Replicating patterns were obtained by EdU incorporation and chemical detection (azide-Alexa Fluor 488; green). Nuclei were counterstained with DAPI (blue). Early S (a-c) and late S replicating nuclei are shown. Panels (a, d) and (d, e) contain DAPI/γH2AX/EdU and γH2AX/EdU merged images, respectively. Panels (c) and (f) illustrate binary masks of red (γH2AX) and green (EdU) channels overlaying the respective DAPI images.

Preliminary results (arithmetic mean of the ratios: 1.57 in early S- and 1.45 in late S-nuclei) suggest a bias in damage distribution towards replicating areas (~50 % higher than expect‐ ed) probably due to local unwinding of chromatin down to naked DNA in both eu- and het‐ erochromatin during DNA synthesis. Chromatin decondensation may increase the susceptibility to DNA damage as well as the accessibility of kinases that phosphorylate H2AX. Noteworthy, detailed visual analysis of fluorescent images or the corresponding bi‐ nary masks in both early and late S-phase revealed that these results were not due to a large amount of γH2AX foci dwelling within replicating area and few of them outside. Instead, γH2AX foci recurrently mapped to the interfaces between replicating and non-replicating regions (Figure 7; Liddle P, unpublished observations).

The fact that in late-replicating cells γH2AX foci tend to map to the boundaries of rep‐ licating compartments (Figure 7, panels d-f) may be due to repositioning of damaged sites to less condensed peripheral heterochromatin regions, as it has been suggested in other models [112, 113]. However, this peculiar distribution of γH2AX foci in replicating/ non-replicating interfaces was also observed in early S-phase when the less compact eu‐ chromatin replicates (Figure 7, panels a-c). In this respect, BLM-induced DNA lesions could preferentially map at the damage-prone TTR located at the boundaries of early and late replicating compartments.

#### **4. Conclusions**

We assayed the influence of eu/heterochromatin replication timing in the distribution of chromatin induced damage using two different approaches: (1) the analysis of UV-C, MMS and etoposide-induced BP in Xp or Xq replicating CHO9 X mitotic chromosome and; (2) the analysis of primary BLM-induced damage (γH2AX foci) in CHO9 early and late replicating interphase nuclei. Our findings support the assumption that induced damage patterns shift according to eu- or heterochromatin replication. The asynchronic replication of eu- or heter‐ ochromatin compartments could influence the distribution of primary DNA lesions and CA, prevailing in replicating chromatin regions, irrespective of its eu- or heterochromatic state. Thus, eu/heterochromatin replication timing seems to play an overriding role in the produc‐ tion and localization of chromosome damage in S-phase cells.

#### **Acknowledgments**

We are indebted to the PEDECIBA Postgraduate Program, the National Agency of Investi‐ gation and Innovation (ANII) and the Alexander von Humboldt Foundation (AvH). Liddle P. is a former Fellow of the AvH Förderung Program at the LMU Biozentrum (Munich).

#### **Author details**

María Vittoria Di Tomaso, Pablo Liddle, Laura Lafon-Hughes, Ana Laura Reyes-Ábalos and Gustavo Folle

\*Address all correspondence to: marvi@iibce.edu.uy

Department of Genetics, Instituto de Investigaciones Biológicas Clemente Estable, Montevi‐ deo, Uruguay

### **References**

The fact that in late-replicating cells γH2AX foci tend to map to the boundaries of rep‐ licating compartments (Figure 7, panels d-f) may be due to repositioning of damaged sites to less condensed peripheral heterochromatin regions, as it has been suggested in other models [112, 113]. However, this peculiar distribution of γH2AX foci in replicating/ non-replicating interfaces was also observed in early S-phase when the less compact eu‐ chromatin replicates (Figure 7, panels a-c). In this respect, BLM-induced DNA lesions could preferentially map at the damage-prone TTR located at the boundaries of early

We assayed the influence of eu/heterochromatin replication timing in the distribution of chromatin induced damage using two different approaches: (1) the analysis of UV-C, MMS and etoposide-induced BP in Xp or Xq replicating CHO9 X mitotic chromosome and; (2) the analysis of primary BLM-induced damage (γH2AX foci) in CHO9 early and late replicating interphase nuclei. Our findings support the assumption that induced damage patterns shift according to eu- or heterochromatin replication. The asynchronic replication of eu- or heter‐ ochromatin compartments could influence the distribution of primary DNA lesions and CA, prevailing in replicating chromatin regions, irrespective of its eu- or heterochromatic state. Thus, eu/heterochromatin replication timing seems to play an overriding role in the produc‐

We are indebted to the PEDECIBA Postgraduate Program, the National Agency of Investi‐ gation and Innovation (ANII) and the Alexander von Humboldt Foundation (AvH). Liddle P. is a former Fellow of the AvH Förderung Program at the LMU Biozentrum (Munich).

María Vittoria Di Tomaso, Pablo Liddle, Laura Lafon-Hughes, Ana Laura Reyes-Ábalos and

Department of Genetics, Instituto de Investigaciones Biológicas Clemente Estable, Montevi‐

tion and localization of chromosome damage in S-phase cells.

\*Address all correspondence to: marvi@iibce.edu.uy

and late replicating compartments.

**4. Conclusions**

366 The Mechanisms of DNA Replication

**Acknowledgments**

**Author details**

Gustavo Folle

deo, Uruguay


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### **Chapter 15**

## **A Histone Cycle**

Douglas Maya, Macarena Morillo-Huesca, Lidia Delgado Ramos, Sebastián Chávez and Mari-Cruz Muñoz-Centeno

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53993

#### **1. Introduction**

Each time a cell divides it must duplicate its DNA content and segregate it equally between two daughter cells. Once a cell has decided to replicate its DNA, hundreds of different pro‐ teins must carefully interact with each other in a very orchestrated process. Defects in any of these steps can lead to cell death and genetic instability and have been shown to be present in many human diseases including cancer [1]. Each step of DNA replication must take place in a correct spatial and temporal window, so cells have evolved complex regulatory net‐ works allowing an efficient regulation of this process. One important feature of eukaryotic cells is that DNA is strongly associated with histones, basic proteins that wrap DNA around octameric structures called nucleosomes. The association of DNA and nucleosomes is com‐ monly known as chromatin.

Nucleosomes are, among others, one of the principal differences between eukaryotic and prokaryotic DNA. Unlike bacteria, eukaryotic cells are not able to live without DNA packed into chromatin [2]. Replication involves dramatic changes in the whole chromatin land‐ scape, since nucleosomes must be removed transiently from the front of the replication ma‐ chinery and repositioned after it. Nucleosome disassembly and assembly involves the action of chromatin remodeling factors, proteins able to destabilize interactions between histones and DNA allowing the interaction of other complexes with DNA. Restoration of chromatin after the replication bubble is a very important step because nucleosomes are not repetitive units of information and contain a specific epigenetic signature or code [3]. In order to en‐ sure enough histones for the nascent DNA, cells must increase the pool of free histones. In human cells, each passage through S-phase requires the synthesis and assembly of almost 30 million nucleosomes that are synthesized mainly during S-phase and are rapidly packaged

© 2013 Maya 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 Maya et al.; licensee InTech. This is a paper 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.

to DNA. Histone production is very tightly coupled to DNA synthesis and is rapidly shut off when replication finishes or is halted by treatment with mutagenic agents. Histone regu‐ lation is very important and accumulation of free histones in the cell has been shown to be highly deleterious for the cell and lead to chromosome loss [2].

Chromatin is a structural barrier for replication but can also play an important role in the regulation of some of the steps within. In this chapter, we will focus on how the chromatin landscape influences DNA replication and show that histones and DNA must adapt to each other in order to ensure a correct genomic duplication. We will describe the influence that chromatin plays at the different stages of DNA replication and then jump to the accurate control that cells exert on histone levels during the cell cycle. We will finally show different situations that uncouple DNA replication from histone deposition and synthesis, and dis‐ cuss if chromatin state can influence the decision of cells to replicate their DNA or not.

### **2. Replication from a chromatin point of view**

#### **2.1. Chromatin influences early steps of replication**

The initiation of DNA replication in any organism requires a series of proteins able to recruit and ultimately load two hexameric DNA helicases. These proteins are able to unwind DNA, a process required to start replication. In eukaryotic cells the pre-initiation complex is formed by two MCM2-7 rings that are loaded in an inactive form next to the Origin Recogni‐ tion Complex (ORC). The MCM helicase must be activated by the sequential action of Dbf4 Dependent Kinase (DDK) and Cyclin Dependent Kinase (CDK), and the addition of other accessory proteins. In mammalian cells, 30,000 to 50,000 origins are sequentially activated each time a cell divides [4].

The nature of the sequence and the structure of replication origins is still a matter of debate, and most higher eukaryotes lack a specific consensus sequence for ORC binding. Origins of DNA Replication Initiation (ORIs) are normally regions of DNA sequence rich in AT that contain a nucleosome-free region (NFR) [5] [6] and it has been suggested that the chromatin environment is important for the establishment of the ORC complex [7] [8]. In *Drosophila* follicle cells histones that localize around ORIs are hyperacetylated and changes in the ace‐ tylation levels of these histones affects ORC binding [9].

As ilustrated in figure 1, methylation of histone H4 has also been shown to be important for ORC recruitment and artificial tethering of the methyltransferase SET8 to a random locus promotes ORC1 binding [10]. Once ORC is bound to DNA, proteins CDC6 and CDT1 help to load the two MCM2-7 helicases to DNA [11]. Loading could also be influenced by the ace‐ tylation of histone H4, since CDT1 is able to recruit the histone acetyltransferase HBO1 to the ORC and enhances the recruitment of MCM2-7 to the origin [12, 13] [14]. Nevertheless, some ORC and MCM subunits are acetylated by Hbo1 in yeast and could therefore be the real targets of this enzyme [14]. Although all origins that are selected are able to load the pre-initiation complex, only one out of every ten will fire. Regulation of firing depends on the activation of the MCM helicases by sequential phosphorylation of some of its subunits by DDK and CDK kinases, that allow the recruitment of CDC45 and the GINS complex [15]. Once these proteins are loaded, the rest of the replication machinery is recruited and replica‐ tion starts.

to DNA. Histone production is very tightly coupled to DNA synthesis and is rapidly shut off when replication finishes or is halted by treatment with mutagenic agents. Histone regu‐ lation is very important and accumulation of free histones in the cell has been shown to be

Chromatin is a structural barrier for replication but can also play an important role in the regulation of some of the steps within. In this chapter, we will focus on how the chromatin landscape influences DNA replication and show that histones and DNA must adapt to each other in order to ensure a correct genomic duplication. We will describe the influence that chromatin plays at the different stages of DNA replication and then jump to the accurate control that cells exert on histone levels during the cell cycle. We will finally show different situations that uncouple DNA replication from histone deposition and synthesis, and dis‐ cuss if chromatin state can influence the decision of cells to replicate their DNA or not.

The initiation of DNA replication in any organism requires a series of proteins able to recruit and ultimately load two hexameric DNA helicases. These proteins are able to unwind DNA, a process required to start replication. In eukaryotic cells the pre-initiation complex is formed by two MCM2-7 rings that are loaded in an inactive form next to the Origin Recogni‐ tion Complex (ORC). The MCM helicase must be activated by the sequential action of Dbf4 Dependent Kinase (DDK) and Cyclin Dependent Kinase (CDK), and the addition of other accessory proteins. In mammalian cells, 30,000 to 50,000 origins are sequentially activated

The nature of the sequence and the structure of replication origins is still a matter of debate, and most higher eukaryotes lack a specific consensus sequence for ORC binding. Origins of DNA Replication Initiation (ORIs) are normally regions of DNA sequence rich in AT that contain a nucleosome-free region (NFR) [5] [6] and it has been suggested that the chromatin environment is important for the establishment of the ORC complex [7] [8]. In *Drosophila* follicle cells histones that localize around ORIs are hyperacetylated and changes in the ace‐

As ilustrated in figure 1, methylation of histone H4 has also been shown to be important for ORC recruitment and artificial tethering of the methyltransferase SET8 to a random locus promotes ORC1 binding [10]. Once ORC is bound to DNA, proteins CDC6 and CDT1 help to load the two MCM2-7 helicases to DNA [11]. Loading could also be influenced by the ace‐ tylation of histone H4, since CDT1 is able to recruit the histone acetyltransferase HBO1 to the ORC and enhances the recruitment of MCM2-7 to the origin [12, 13] [14]. Nevertheless, some ORC and MCM subunits are acetylated by Hbo1 in yeast and could therefore be the real targets of this enzyme [14]. Although all origins that are selected are able to load the pre-initiation complex, only one out of every ten will fire. Regulation of firing depends on

highly deleterious for the cell and lead to chromosome loss [2].

**2. Replication from a chromatin point of view**

**2.1. Chromatin influences early steps of replication**

tylation levels of these histones affects ORC binding [9].

each time a cell divides [4].

378 The Mechanisms of DNA Replication

**Figure 1. Chromatin influences DNA replication fork establishment.** Schematic representation of the different steps during the assembly and activation of the replication fork machinery that are influenced by histone modifica‐ tions. 1.0RC recruitment is influenced by methylation and acetylation levels of histone I-I4. 2.Acetylation of histone H4 by the Hbo1 might influence loading of the two Mcm2-7 helicases. 3. H4K 16 aoetytation is related to the timing of origin firing distribution.

Two interesting observations have lead to the hypothesis that chromosome architecture can also be important for origin usage. One of them is that origins seem to be organized into clusters of 5 to 10 origins that fire simultaneously [18]. Cohesins are enriched next to origins and depletion of the RAD21 cohesin subunit greatly reduces the number of active origins. This ring-like complex is able to wrap two chromatin fibers and creates a chromatin loop. It has been suggested that this spatial organization of chromatin could define replication do‐ mains that are activated synchronously [16, 17]. In agreement with this hypothesis, analysis of the oriGNA13 replication origin of hamster cells shows that active origins localize close to the base of chromatin loops [18]. The second feature that directly relates chromatin structure and replication is that genome replication does not take place in a single and continuous way. Domains containing several megabases of contiguous DNA are replicated earlier than others [19] and this replication timing is somehow correlated with acetylation levels of his‐ tone H4 at the K16 residue [20].

Chromatin influences the recruitment and the activity of different elements of the replica‐ tion machinery. Once this machinery has been set up and is fully active, fork progression must now cope with the fact that approximately every 192 bp there is a nucleosome that must be displaced from DNA in order to continue with replication.

#### **2.2. Nucleosome reorganization around the replication fork**

Replication fork progression involves many proteins that interact closely with DNA. Elec‐ tron micrography of replicating SV40 mini-chromosomes has shown that 300bp ahead of the replication fork, DNA remains *naked*, or at least contains nucleosomes that are unstable when compared to a canonical nucleosome [21] [22]. MCM progression in mammalian cells suggests that chromatin is decondensed in front of the replication fork [23] and artificial tethering of Cdc45 to DNA is able to promote a partial decondensation of chromatin without DNA synthesis [24]. This initial decondensation could be related to an increase in the mobi‐ lity of histone H1 due to its phosphorylation by the cyclin A-CDK2 complex. It is still un‐ clear if nucleosome disruption in front of the replication fork is due to specific chromatin remodeling in front of the fork or to the passage of the replication machinery itself [3].

Nucleosome disassembly and reassembly are processes quite well described for transcrip‐ tion. The efficiency of these processes is largely dependent on chromatin-remodeling com‐ plexes, proteins able to interact with and change the stability of chromatin, allowing the transcription machinery to interact with DNA. There are many different chromatin-remod‐ eling complexes and all of them are possible candidates for nucleosome eviction during rep‐ lication. The fact that chromatin disassembly and assembly occur in such a small spatial window makes it very difficult to distinguish between the complexes required for one or the other process. There are two major complexes that could be involved in H2A/H2B displace‐ ment during replication: FACT and NAP1, and another two for H3/H4: Asf1 and CAF1.

The FACT complex is composed of two main subunits, SPT16 and SSRP1 (Pob3 in *S.cere‐ visiae*), and plays a key role in nucleosome reorganization during transcription elonga‐ tion. FACT function has been mostly related to the displacement of an H2A/H2B dimer during transcription, but it has also been proposed that displacement could be an indi‐ rect effect of nucleosome reorganization by this complex [25, 26]. There are many differ‐ ent items of evidence suggesting that FACT plays a role as a histone chaperone during DNA replication. FACT is required for DNA replication in *Xenopus* extracts, is present at human replication origins [27] and has been co-purified as part of the replication fork progression complex in yeast [28]. The other H2A/H2B histone chaperone candidate is Nap1, which has been shown to disassemble nucleosomes in vitro when combined with the RSC complex [29, 30]. Once H2A/H2B dimers are displaced, the H3/H4 tetramer is now accessible for an H3/H4 chaperone.

To date, it is not known if H2A/H2B dimers removed during replication are recycled. On the contrary, it is well established that the original H3/H4 tetramer present in front of the repli‐ cation fork machinery is restored after the replication fork in a random semi-conservative process [31]. The fact that the H3/H4 tetramer is recycled suggests that a histone chaperone must disassemble this tetramer in front of the replication machinery and reassemble it after. One good candidate for this process is Asf1. This protein along with Chromatin Assembly Factor 1 (CAF1) plays a key role in deposition of new H3/H4 following passage of the repli‐ cation fork. Asf1 binds PCNA and replication factor C [32], and can also bind the MCM heli‐ case complex through histones H3 and H4. Upon a replication fork progression block, Asf1 can be found associated with post-translationally modified H3/H4 histones, which most likely belong to the parental chromatin [33].

and replication is that genome replication does not take place in a single and continuous way. Domains containing several megabases of contiguous DNA are replicated earlier than others [19] and this replication timing is somehow correlated with acetylation levels of his‐

Chromatin influences the recruitment and the activity of different elements of the replica‐ tion machinery. Once this machinery has been set up and is fully active, fork progression must now cope with the fact that approximately every 192 bp there is a nucleosome that

Replication fork progression involves many proteins that interact closely with DNA. Elec‐ tron micrography of replicating SV40 mini-chromosomes has shown that 300bp ahead of the replication fork, DNA remains *naked*, or at least contains nucleosomes that are unstable when compared to a canonical nucleosome [21] [22]. MCM progression in mammalian cells suggests that chromatin is decondensed in front of the replication fork [23] and artificial tethering of Cdc45 to DNA is able to promote a partial decondensation of chromatin without DNA synthesis [24]. This initial decondensation could be related to an increase in the mobi‐ lity of histone H1 due to its phosphorylation by the cyclin A-CDK2 complex. It is still un‐ clear if nucleosome disruption in front of the replication fork is due to specific chromatin remodeling in front of the fork or to the passage of the replication machinery itself [3].

Nucleosome disassembly and reassembly are processes quite well described for transcrip‐ tion. The efficiency of these processes is largely dependent on chromatin-remodeling com‐ plexes, proteins able to interact with and change the stability of chromatin, allowing the transcription machinery to interact with DNA. There are many different chromatin-remod‐ eling complexes and all of them are possible candidates for nucleosome eviction during rep‐ lication. The fact that chromatin disassembly and assembly occur in such a small spatial window makes it very difficult to distinguish between the complexes required for one or the other process. There are two major complexes that could be involved in H2A/H2B displace‐ ment during replication: FACT and NAP1, and another two for H3/H4: Asf1 and CAF1.

The FACT complex is composed of two main subunits, SPT16 and SSRP1 (Pob3 in *S.cere‐ visiae*), and plays a key role in nucleosome reorganization during transcription elonga‐ tion. FACT function has been mostly related to the displacement of an H2A/H2B dimer during transcription, but it has also been proposed that displacement could be an indi‐ rect effect of nucleosome reorganization by this complex [25, 26]. There are many differ‐ ent items of evidence suggesting that FACT plays a role as a histone chaperone during DNA replication. FACT is required for DNA replication in *Xenopus* extracts, is present at human replication origins [27] and has been co-purified as part of the replication fork progression complex in yeast [28]. The other H2A/H2B histone chaperone candidate is Nap1, which has been shown to disassemble nucleosomes in vitro when combined with the RSC complex [29, 30]. Once H2A/H2B dimers are displaced, the H3/H4 tetramer is

must be displaced from DNA in order to continue with replication.

**2.2. Nucleosome reorganization around the replication fork**

tone H4 at the K16 residue [20].

380 The Mechanisms of DNA Replication

now accessible for an H3/H4 chaperone.

**Figure 2. Nucleosome reorganization around the replication fork**. Representation of the different nucleosome re‐ organization events that take place during replication fork progression. In order to simplify the figure, the DNA repli‐ cation machinery and other accessory proteins that are important during fork progression are not shown. Interrogation marks are used when the protein/s involved in such process remain unknown or when the pathway has not been directly demonstrated.

Deposition of histone octamers occurs as soon as DNA is long enough to wrap around a nu‐ cleosome [21]. Since one H3/H4 tetramer is recycled after DNA replication, one new H3/H4 tetramer must be deposited on the other strand. The mechanism of reposition of the original H3/H4 tetramer remains unclear but probably involves Asf1 (see previous paragraph). In‐ corporation of the new H3/H4 tetramer on the other hand is more defined and involves the action of CAF1 and Asf1. CAF1 is recruited to both leading and lagging strands by the pro‐ liferating cell nuclear antigen PCNA. Depletion of CAF1 produces a clear decrease in the as‐ sembly of new chromatin during replication [34], activates the DNA Damage Checkpoint (DDR), and stalls replication forks [35], suggesting that efficient chromatin repositioning af‐ ter replication is important for replication fork progression. Asf1 plays a role in this process as a histone pool protein that delivers H3/H4 dimers to CAF1. After the H3/H4 tetramer is assembled, H2A/H2B becomes incorporated into chromatin in a process that probably in‐ volves FACT or NAP1. Finally, H1 protein is incorporated to allow further compacting of chromatin. Incorporation of H1 is probably mediated by the NASP protein and is required for efficient S-phase progression [36].

One interesting feature recently described is that the chromatin landscape influences the length of the Okazaki fragments synthesized at the lagging strand during DNA replication [37]. Due to the 5´to 3´ polarity of DNA polymerase, synthesis of DNA in the lagging strand is discontinuous and generates short fragments of DNA named Okazaki fragments. These fragments must then join to form a unique DNA strand in a process known as maturation. Okazaki fragment maturation requires the sequential action of the flap endonuclease 1 (FEN1) and DNA ligase I. This group has demonstrated that the ligation junctions of Okaza‐ ki fragments are preferentially located in the nucleosome midpoint. The length of Okazaki fragments depends on the chromatin behind the replication fork and mutations that impair chromatin repositioning increase the average size of Okazaki fragments. According to their model, Pol runs into the nucleosome assembled into the previous Okazaki fragment and this triggers termination, flap processing and ligation.

#### **2.3. Chromatin maturation and centromere formation**

After nucleosome incorporation to DNA, two major processes must take place: chromatin maturation and centromere formation. Histones start to acquire certain modifications in their tails as soon as they are repositioned to DNA. Nascent chromatin is highly acetylated and must be deacetylated and methylated to reach a more compact state. Deacetylation nor‐ mally takes place by the histone deacetylases HDAC1-3 and DNA methylation by the DNA methyltransferase 1 (DNMT1). In addition to chromatin compacting, there are also some specific post-transcriptional modifications that must be acquired to establish a specific epi‐ genetic code that is transmitted to daughter cells. Maintaining this "epigenetic memory" of daughter cells is important and has implications during cell differentiation (23). Restoration of all these marks does not take place exclusively in replication and can also take place dur‐ ing mitosis or even in daughter cells [38, 39]. Replication of the chromatin near the centro‐ mere is also vital to ensure an efficient segregation during mitosis. This heterochromatin presents a specific variant of histone H3 named CENP-A, which is essential for the efficient binding of the kinetochore during mitosis. The kinetochore is a huge structure that attaches to centromeric DNA and mediates the interaction of chromosomes with the mitotic spindle and their movement to the spindle poles during mitosis [40]. Accurate segregation of chro‐ mosomes relies on the correct formation of the spindle apparatus.

H3/H4 tetramer remains unclear but probably involves Asf1 (see previous paragraph). In‐ corporation of the new H3/H4 tetramer on the other hand is more defined and involves the action of CAF1 and Asf1. CAF1 is recruited to both leading and lagging strands by the pro‐ liferating cell nuclear antigen PCNA. Depletion of CAF1 produces a clear decrease in the as‐ sembly of new chromatin during replication [34], activates the DNA Damage Checkpoint (DDR), and stalls replication forks [35], suggesting that efficient chromatin repositioning af‐ ter replication is important for replication fork progression. Asf1 plays a role in this process as a histone pool protein that delivers H3/H4 dimers to CAF1. After the H3/H4 tetramer is assembled, H2A/H2B becomes incorporated into chromatin in a process that probably in‐ volves FACT or NAP1. Finally, H1 protein is incorporated to allow further compacting of chromatin. Incorporation of H1 is probably mediated by the NASP protein and is required

One interesting feature recently described is that the chromatin landscape influences the length of the Okazaki fragments synthesized at the lagging strand during DNA replication [37]. Due to the 5´to 3´ polarity of DNA polymerase, synthesis of DNA in the lagging strand is discontinuous and generates short fragments of DNA named Okazaki fragments. These fragments must then join to form a unique DNA strand in a process known as maturation. Okazaki fragment maturation requires the sequential action of the flap endonuclease 1 (FEN1) and DNA ligase I. This group has demonstrated that the ligation junctions of Okaza‐ ki fragments are preferentially located in the nucleosome midpoint. The length of Okazaki fragments depends on the chromatin behind the replication fork and mutations that impair chromatin repositioning increase the average size of Okazaki fragments. According to their model, Pol runs into the nucleosome assembled into the previous Okazaki fragment and

After nucleosome incorporation to DNA, two major processes must take place: chromatin maturation and centromere formation. Histones start to acquire certain modifications in their tails as soon as they are repositioned to DNA. Nascent chromatin is highly acetylated and must be deacetylated and methylated to reach a more compact state. Deacetylation nor‐ mally takes place by the histone deacetylases HDAC1-3 and DNA methylation by the DNA methyltransferase 1 (DNMT1). In addition to chromatin compacting, there are also some specific post-transcriptional modifications that must be acquired to establish a specific epi‐ genetic code that is transmitted to daughter cells. Maintaining this "epigenetic memory" of daughter cells is important and has implications during cell differentiation (23). Restoration of all these marks does not take place exclusively in replication and can also take place dur‐ ing mitosis or even in daughter cells [38, 39]. Replication of the chromatin near the centro‐ mere is also vital to ensure an efficient segregation during mitosis. This heterochromatin presents a specific variant of histone H3 named CENP-A, which is essential for the efficient binding of the kinetochore during mitosis. The kinetochore is a huge structure that attaches to centromeric DNA and mediates the interaction of chromosomes with the mitotic spindle

for efficient S-phase progression [36].

382 The Mechanisms of DNA Replication

this triggers termination, flap processing and ligation.

**2.3. Chromatin maturation and centromere formation**

CENP-A (also known as CENH3) is an essential protein that replaces histone H3 at centro‐ meric DNA. This protein is highly divergent among different species but is functionally well conserved since the homologue protein of *S. cerevisiae,* Cse4 is able to complement human cells lacking CENP-A or *vice versa* [41]. In human cells, CENP-A is not assembled on to DNA just after DNA replication and CENP-A-containing nucleosomes are interspersed with can‐ onical nucleosomes during replication of centromeres [42, 43]. This organization promotes the folding of centromeric chromatin into a unique structure during metaphase, in which all the nucleosomes containing CENP-A are facing the external side of chromosomes. This structure allows kinetochore assembly and microtubule attachment and favours sister chro‐ matid cohesion [44]. Once chromosomes are separated, CENH3 is fully positioned on cen‐ tromeric chromatin during the period between telophase and G1 in a process that is dependent on the transient incorporation of Mis18 and KNL2 in anaphase [45]. The incorpo‐ ration of CENP-A to centromeric chromatin is mediated at least in part by the HJURP pro‐ tein (Scm3 in *S. cerevisiae*) and is related to low levels of acetylation of the K16 residue of histone H4. Defects in the proper incorporation of this histone variant can lead to cell death, genetic instability and chromosome loss [46-48]. There is also a subset of specific proteins important to prevent the deposition of CENH3 containing nucleosomes out of the centro‐ meric DNA. In *S. cerevisiae*, the ubiquitin E3 ligase Psh1 prevents the spread of Cse4 contain‐ ing nucleosomes out of the centromere [49] [50]. The absence of both CAF1 and HIRA also leads to the presence of this type of nucleosomes in euchromatic regions in both *S. cerevisiae* and *S. pombe* and has been shown to cause genetic instability [51]. Finally, several papers point out that a proper homeostasis between H3 and CENH3 histones is important for the distribution of this centromeric variant and for efficient chromosome segregation [52, 53].

### **3. From gene to protein, histones are highly regulated**

It is clear that there is a strong interdependency between DNA replication and chromatin reorganization. Nucleosomes are more than structural bricks for DNA, and require the mod‐ ification of specific residues or the substitution of certain histone variants for others to main‐ tain a correct epigenetic state. Once cells have decided to replicate their DNA, an increase in the abundance of histone proteins is required to pack the new genome that is about to be generated. Histone genes are among the most highly cell-cycle-regulated genes [54] because cells need to ensure a high demand of histones during replication, but must make sure that these levels are quickly down-regulated when replication slows down or is blocked, to avoid the deleterious effects of free histones on cell survival.

Canonical histone proteins can be regulated at transcriptional, post-transcriptional, transla‐ tional and post-translational levels. The importance of each pathway on histone metabolism largely depends on the organism. In *S. cerevisiae* for example, the transcriptional regulation has been traditionally shown to play a major role in histone regulation, while in mammalian cells, post-transcriptional and translational mechanisms seem to be more important. Never‐ theless, it is clear that all organisms try to produce histones exclusively during the replica‐ tive S-phase and more specifically only when replication is actively taking place.

#### **3.1. Histones are regulated from the beginning: transcriptional regulation**

Histone transcription is tightly regulated during the cell cycle. In some organisms like *S. cer‐ evisiae*, transcription of histones can only be detected in late G1 and during DNA replication [55]. In higher eukaryotes, however, histones mRNAs can be found at all stages but increase as cells enter S-phase [56]. Expression of all canonical histones must be stoichiometric and several studies show that an imbalance between the different histone subtypes can be highly deleterious for the cell [52, 53, 57].

In metazoans, entry into S-phase increases the expression of replication-dependent histone genes three to five-fold [58]. Histone genes are clustered and each cluster normally contains at least one copy of the five canonical histones. Although transcription of all histones is care‐ fully coordinated, no obvious common sequence element has been found at their promoters. Nevertheless, common elements can be found for some particular histone variants, like the Octamer-binding Transcription Factor (OTF1) for H2B promoters [59] or the Coding Region Activating Sequences (CRAS) in H2A, H3 and H4 genes [60]. Activation of histone gene transcription requires the Nuclear Protein Ataxia-Telangiectasia (NPAT), which is essential for S-phase progression [61]. This protein normally locates next to the *Histone locus bodies* and is phosphorylated by cyclin E-CDK2 at the beginning of S-Phase. Phosphorylation per‐ sists through S-phase and increases histone gene transcription [62, 63].

Transcriptional regulation of histones in *S. cerevisiae* is largely dependent on the integri‐ ty of the HIR complex. This complex is conserved from yeast to humans and has been shown to play a role in both of them in replication-independent chromatin assembly. In yeast, this complex is composed of the three subunits Hir1-3 and Hpc2. Deletion of any of the subunits leads to a de-repression of histones outside of S-phase [64]. Histone genes are grouped in 4 clusters, and each of them express simultaneously H2A/H2B or H3/H4 from a bidirectional promoter. These promoters contain upstream activating se‐ quences (UAS) required for the recruitment of activators such as Spt10 and SBF [65]. Three of these four clusters also contain a negative regulatory site (NEG or CCR) able to maintain these genes in a repressed state in cell cycle phases outside of late G1 and Sphase and under replication stress conditions [64, 66, 67]. Deletion of the negative regu‐ latory site is able to de-repress the *HTA1-HTB1* histone locus and allow expression outside of S-phase. The mechanism of repression is not completely understood but in‐ volves changes at the chromatin structure creating a repressive chromatin, which de‐ pends on the HIR complex, proteins Rtt106, Yta7 and Asf1, and the chromatin remodelling complex RSC. Two recent reports coming from the same group have shed some light on how repressive chromatin switches to active chromatin (in terms of tran‐ scription). The first one involves the degradation of Yta7 mediated by a phosphorylation event that involves Casein Kinase II (CKAII) and the cyclin-dependent kinase Cdc28. Degradation of Yta7 allows the efficient expression of histone mRNAs during S-phase through a mechanism that could involve transcription elongation efficiency [68]. The sec‐ ond report is related to the cell cycle regulation of Spt21, an activator of histone gene expression [69]. Spt21 protein levels outside S-phase are regulated by proteolysis, in a mechanism that depends on a complex formed by the Anaphase Promoting Complex (APC) with Cdh1 during G1, and on APC-Cdc20 during G2 and M (Brenda Andrews, EMBO transcription meeting 2012).

cells, post-transcriptional and translational mechanisms seem to be more important. Never‐ theless, it is clear that all organisms try to produce histones exclusively during the replica‐

Histone transcription is tightly regulated during the cell cycle. In some organisms like *S. cer‐ evisiae*, transcription of histones can only be detected in late G1 and during DNA replication [55]. In higher eukaryotes, however, histones mRNAs can be found at all stages but increase as cells enter S-phase [56]. Expression of all canonical histones must be stoichiometric and several studies show that an imbalance between the different histone subtypes can be highly

In metazoans, entry into S-phase increases the expression of replication-dependent histone genes three to five-fold [58]. Histone genes are clustered and each cluster normally contains at least one copy of the five canonical histones. Although transcription of all histones is care‐ fully coordinated, no obvious common sequence element has been found at their promoters. Nevertheless, common elements can be found for some particular histone variants, like the Octamer-binding Transcription Factor (OTF1) for H2B promoters [59] or the Coding Region Activating Sequences (CRAS) in H2A, H3 and H4 genes [60]. Activation of histone gene transcription requires the Nuclear Protein Ataxia-Telangiectasia (NPAT), which is essential for S-phase progression [61]. This protein normally locates next to the *Histone locus bodies* and is phosphorylated by cyclin E-CDK2 at the beginning of S-Phase. Phosphorylation per‐

Transcriptional regulation of histones in *S. cerevisiae* is largely dependent on the integri‐ ty of the HIR complex. This complex is conserved from yeast to humans and has been shown to play a role in both of them in replication-independent chromatin assembly. In yeast, this complex is composed of the three subunits Hir1-3 and Hpc2. Deletion of any of the subunits leads to a de-repression of histones outside of S-phase [64]. Histone genes are grouped in 4 clusters, and each of them express simultaneously H2A/H2B or H3/H4 from a bidirectional promoter. These promoters contain upstream activating se‐ quences (UAS) required for the recruitment of activators such as Spt10 and SBF [65]. Three of these four clusters also contain a negative regulatory site (NEG or CCR) able to maintain these genes in a repressed state in cell cycle phases outside of late G1 and Sphase and under replication stress conditions [64, 66, 67]. Deletion of the negative regu‐ latory site is able to de-repress the *HTA1-HTB1* histone locus and allow expression outside of S-phase. The mechanism of repression is not completely understood but in‐ volves changes at the chromatin structure creating a repressive chromatin, which de‐ pends on the HIR complex, proteins Rtt106, Yta7 and Asf1, and the chromatin remodelling complex RSC. Two recent reports coming from the same group have shed some light on how repressive chromatin switches to active chromatin (in terms of tran‐ scription). The first one involves the degradation of Yta7 mediated by a phosphorylation event that involves Casein Kinase II (CKAII) and the cyclin-dependent kinase Cdc28. Degradation of Yta7 allows the efficient expression of histone mRNAs during S-phase

tive S-phase and more specifically only when replication is actively taking place.

**3.1. Histones are regulated from the beginning: transcriptional regulation**

sists through S-phase and increases histone gene transcription [62, 63].

deleterious for the cell [52, 53, 57].

384 The Mechanisms of DNA Replication

It has been recently shown that the HIR complex is conserved through evolution [70, 71] [55]. In humans, this complex is formed by three proteins: HIRA, Ubinuclein1 and Cabin1. The role described for HIRA in humans has been mostly associated with chromatin assem‐ bly of the transcriptional histone variant H3.3 in cooperation with ASF1 [72]. Nevertheless, several studies suggest that this complex could also play an important role in metazoan his‐ tone regulation. Ectopic over-expression of HIRA is able to repress histone gene transcrip‐ tion and block S-phase progression in human cells. This protein localizes to histone gene clusters in an immunofluorescence essay [73]. Cyclin E-CDK2 and cyclin A-CDK2 can phos‐ phorylate HIRA, and this phosphorylation is inhibited by cyclin inhibitor p21, which has been shown to be important for repression of histone synthesis upon replication stress [74] [69]. HIRA could therefore be acting as a repressor of histone gene expression outside of Sphase regulated through phosphorylation by the cyclin E-CDK2. In this model, phosphory‐ lation by the cyclin E-CDK2 could switch histone expression by activating NPAT and inactivating HIRA.

#### **3.2. Once they are transcribed: post-transcriptional and translational regulation**

Mammalian histone mRNAs lack introns and do not have a poly(A) tail as do most mRNAs. Instead, they contain a special 3´UTR sequence that forms a stem-loop structure [54]. His‐ tone clusters localize to specific Cajal Bodies that are enriched in factors required for expres‐ sion (NPAT) and maturation (U7 snRNA) of histone mRNA named *histone locus bodies* [75]. Maturation requires the formation of the 3´end through an endonucleolytic cleavage that has been shown to be important for transcription termination [76, 77]. Cleavage takes place between the stem-loop and the histone downstream element (HDE). The machinery in‐ volved in this process uses some common elements from the processing machinery of polya‐ denylated mRNAs but has also some specific components like SLBP, the Sm-like proteins (LSM1-11), the U7 snRNA and ZFP100. Additional information on maturation of histone mRNAs can be found in a nice review published some years ago [54]. The Stem Loop Bind‐ ing Protein, SLBP, is one of the most important proteins for post-transcriptional and transla‐ tional regulation of histone mRNAs and accompanies histone mRNA throughout its life.

SLBP is the only known cell cycle regulated protein of all the histone processing machinery. This protein starts accumulating during late G1 and is degraded at the end of S-phase by the phosphorylation of two threonine residues that target it for degradation [78]. There are three major roles for SLBP on histone regulation: 1. Allow an efficient cleavage during mRNA ma‐ turation 2. Facilitate circularization of histone mRNAs, required for their efficient translation by polyribosomes [79, 80] and 3. Increase histone mRNA stability preventing degradation of histone mRNAs by the 3´hExo [81]. Nevertheless, histone mRNA are still degraded when SLBP is artificially present at constitutive levels at the end of S-phase [78] or when DNA rep‐ lication is inhibited [82], indicating that although this protein has a major contribution to histone mRNA stability, it is not able to prevent degradation itself.

Canonical histone mRNAs in lower eukaryotes and plants are polyadenylated. The fact that these transcripts lack any known specific structure at their 3´end and have a short half-life has lead to the conclusion that regulation in these organisms mostly takes place at a transcriptional level. Nevertheless, there is quite a lot of recent evidence that strong‐ ly suggests the importance of the post-transcriptional regulation of histone mRNAs in *S. cerevisiae*. Several reports implicate some of the components of the exosome in the specif‐ ic degradation of the H2B transcripts [83]. One year ago, Herrero and Moreno revealed the importance of the SM-like protein Lsm1 in histone mRNA degradation. Mutants lack‐ ing Lsm1 are sensitive to DNA damaging drugs and histone over-expression, and show a defect in histone mRNA degradation under replication stress conditions [84]. This pro‐ tein is part of the Lsm1-7-Pat1 mRNA degradation complex, which has an important role in histone mRNA degradation under replication stress conditions in human cells. Lsm1-7-Pat1 has been shown to bind preferentially mRNAs carrying U-tracts in human cells, and oligoadenylated over polyadenylated mRNAs in yeast [85, 86]. Upon DNA replication arrest, histone mRNAs in human cells suffer an oligouridylation process ac‐ quiring a terminal oligo U-tract required for an efficient degradation by this complex [87]. Uridylation of mRNA has not been detected to date in *S. cerevisiae* but there is a re‐ cent report showing that the average length of the poly(A) tail of the yeast H2B histone mRNA is quite short compared to other transcripts. The length of this poly(A) is cell cy‐ cle-dependent and seems to decrease as cells exit G1 and progress through S-phase up to G2, when some of the transcripts completely lack a poly(A) tail [88]. This difference in length opens a possible explanation as to how the Lsm1-7 yeast complex preferentially recognizes yeast histone mRNAs over other transcripts to degrade them at the end of Sphase.

#### **3.3. Last frontier of histone regulation: controling protein levels**

In addition to the tight regulation of histone mRNA levels, a mechanism able to control histone protein levels was described some years ago [89]. To date, this pathway has on‐ ly been described in the yeast *S. cerevisiae* and involves the action of the yeast homo‐ logue of CHK2, Rad53. Rad53 plays an important role in the DNA Damage Response and has been shown to be essential upon DNA damage or replication stress [1, 90]. His‐ tone degradation involves the direct action of Rad53 along with the E2 ubiquitin ligases (UL) Ubc4 and Ubc5 and the E3 UL Tom1 [91]. This complex is able to degrade histo‐ nes in a mechanism that involves tyrosine phosphorylation and poly-ubiquitylation, be‐ fore their proteolysis by the proteasome. Histone degradation is independent on the central DNA damage checkpoint signal, since it does not depend on other kinases in‐ volved in the DDR like Mec1 (ATM) or Tel1 (ATR). Further studies in higher eukaryotes need to be done to confirm if this pathway is conserved in all eukaryotes.

**Figure 3. Regulation of histone levels in** *S.cerevisiae* **and** *H.sapiens.* Different mechanisms able to control histone levels in *S.cerevisiae* and *H.sapiens*. Arrows normally indicate a positive effect on the pathway and straight lines a neg‐ ative One. Interrogation marks are used when the protein/s involved in such process remain unknown or when the pathway has not been directly demonstrated. The big interrogation mark shown for post-translational regulation in the *H.sapiens* column, remarks that this pathway has not been demonstrated to date in human cells.

### **4. Histones: Enough to pack but not too much**

SLBP is artificially present at constitutive levels at the end of S-phase [78] or when DNA rep‐ lication is inhibited [82], indicating that although this protein has a major contribution to

Canonical histone mRNAs in lower eukaryotes and plants are polyadenylated. The fact that these transcripts lack any known specific structure at their 3´end and have a short half-life has lead to the conclusion that regulation in these organisms mostly takes place at a transcriptional level. Nevertheless, there is quite a lot of recent evidence that strong‐ ly suggests the importance of the post-transcriptional regulation of histone mRNAs in *S. cerevisiae*. Several reports implicate some of the components of the exosome in the specif‐ ic degradation of the H2B transcripts [83]. One year ago, Herrero and Moreno revealed the importance of the SM-like protein Lsm1 in histone mRNA degradation. Mutants lack‐ ing Lsm1 are sensitive to DNA damaging drugs and histone over-expression, and show a defect in histone mRNA degradation under replication stress conditions [84]. This pro‐ tein is part of the Lsm1-7-Pat1 mRNA degradation complex, which has an important role in histone mRNA degradation under replication stress conditions in human cells. Lsm1-7-Pat1 has been shown to bind preferentially mRNAs carrying U-tracts in human cells, and oligoadenylated over polyadenylated mRNAs in yeast [85, 86]. Upon DNA replication arrest, histone mRNAs in human cells suffer an oligouridylation process ac‐ quiring a terminal oligo U-tract required for an efficient degradation by this complex [87]. Uridylation of mRNA has not been detected to date in *S. cerevisiae* but there is a re‐ cent report showing that the average length of the poly(A) tail of the yeast H2B histone mRNA is quite short compared to other transcripts. The length of this poly(A) is cell cy‐ cle-dependent and seems to decrease as cells exit G1 and progress through S-phase up to G2, when some of the transcripts completely lack a poly(A) tail [88]. This difference in length opens a possible explanation as to how the Lsm1-7 yeast complex preferentially recognizes yeast histone mRNAs over other transcripts to degrade them at the end of S-

histone mRNA stability, it is not able to prevent degradation itself.

386 The Mechanisms of DNA Replication

**3.3. Last frontier of histone regulation: controling protein levels**

need to be done to confirm if this pathway is conserved in all eukaryotes.

In addition to the tight regulation of histone mRNA levels, a mechanism able to control histone protein levels was described some years ago [89]. To date, this pathway has on‐ ly been described in the yeast *S. cerevisiae* and involves the action of the yeast homo‐ logue of CHK2, Rad53. Rad53 plays an important role in the DNA Damage Response and has been shown to be essential upon DNA damage or replication stress [1, 90]. His‐ tone degradation involves the direct action of Rad53 along with the E2 ubiquitin ligases (UL) Ubc4 and Ubc5 and the E3 UL Tom1 [91]. This complex is able to degrade histo‐ nes in a mechanism that involves tyrosine phosphorylation and poly-ubiquitylation, be‐ fore their proteolysis by the proteasome. Histone degradation is independent on the central DNA damage checkpoint signal, since it does not depend on other kinases in‐ volved in the DDR like Mec1 (ATM) or Tel1 (ATR). Further studies in higher eukaryotes

phase.

Histone levels are regulated as soon as transcription of its mRNA starts. On top of the nor‐ mal cell cycle regulation, additional mechanisms are able to block histone production when replication slows down or is completely blocked. Eukaryotic cells are unable to live without histones [92] and inhibition of histone deposition behind the replication fork blocks DNA synthesis and activates the DNA Damage Response (DDR) [35]. Eukaryotes seem to have evolved to a situation in which histones must not be free in the cell and DNA must not be free of histones. In this last part of the chapter, we will focus on how cells cope with situa‐ tions that break this bidirectional relationship.

#### **4.1. Harmful effects of free histones**

Histones are basic proteins that can bind-specifically to negatively charged molecules. Re‐ constitution experiments show that a slight excess of histones over DNA is sufficient to pro‐ mote chromatin aggregation, probably due to the neutralization of negative charged DNA. In yeast, high levels of histones increase chromosome loss and enhance DNA damage sensi‐ tivity [57, 89]. Defects in histone degradation during replication stress or DNA damage se‐ verely decreases cell viability [84]. Free histones show electrostatic interactions with some cellular macromolecules carrying the opposite charge such as RNA molecules. Additionally, an excess of free histones can saturate and inhibit the activity of some histone modifying en‐ zymes, and change the expression pattern of almost 240 genes [93]. Two different studies in the yeasts *S. cerevisiae* and *Schizosaccharomyces pombe* have demonstrated the importance of a correct balance between histone H3 and the centromeric variant Cse4 (CENPA) for efficient chromosome segregation. H3 can compete with Cse4 in the assembly of centromeric chro‐ matin and this competition largely depends on a correct balance between levels of H3 and H4 [52, 53]. Cells must therefore not only prevent the accumulation of free histones but also ensure a correct homeostasis between canonical and other histone variants. Once cells have decided to initiate replication, any problem that unbalances replication fork progression with histone levels can potentially lead to an increase in the abundance of free histones. In order to prevent this, there is an additional pathway linked to the DDR able to block histone synthesis under DNA replication stress conditions or replication fork arrest.

#### **4.2. The DNA Damage Response (DDR): Coupling DNA and histone synthesis**

DDR is probably one of the most well characterized checkpoints in the cell and is normally activated whenever a cell senses DNA damage. Activation leads to the sequential action of a cascade of kinases that block or delay cell cycle progression to allow the cell to correct the damage. If damage cannot be repaired, human cells enter the apoptosis program and die [94]. Proper functioning of this pathway is essential for genome integrity and mutations in most of the branches of this path are linked to cancer and other diseases. DDR is able to block cells at G1, S and G2/M [95]. In human cells, two kinases ATM and ATR (Tel1 and Mec1 respectively in *S. cerevisiae*) play a major role in the activation of the DDR. ATM has been directly involved in the activation of a mechanism that ultimately leads to repression of histone expression.

In human cells, activation of histone gene transcription requires NPAT phosphorylation by the cyclin E-CDK2 complex at the beginning of S-Phase. Activation of NPAT is essential for S-phase progression and histone expression. Repression of histone synthesis upon DNA damage requires the activation of ATM, which leads to the sequential activation of p53 and then p21. p21 is able to block the activity of the cyclin E-CDK2 complex. Inhibition of this complex leads to a progressive dephosphorylation of NPAT, which no longer localizes to histone clusters to activate transcription [96]. One interesting hypothesis that remains to be tested is if this cascade could also lead to histone repression by a change in the activity or location of HIRA, the human homologue of the HIR complex, at histone promoters (see pre‐ vious paragraph about transcriptional regulation of histones). DNA damage also promotes post-transcriptional degradation of histone mRNAs. Treatment of cells with hydroxyurea (HU) increases oligouridylation of histone mRNAs in a process that depends on Upf1. Upf1 binds SLBP and helps to recruit a 3' Terminal Uridylyl Transferase (TUT-ase) required for oligourydilation. These 3′ oligo(U) tails are recognized by the Lsm1–7 complex that triggers mRNA degradation through the exosome and Xrn1 [87]. How Upf1 is recruited to histone mRNAs upon DNA damage remains unknown.

evolved to a situation in which histones must not be free in the cell and DNA must not be free of histones. In this last part of the chapter, we will focus on how cells cope with situa‐

Histones are basic proteins that can bind-specifically to negatively charged molecules. Re‐ constitution experiments show that a slight excess of histones over DNA is sufficient to pro‐ mote chromatin aggregation, probably due to the neutralization of negative charged DNA. In yeast, high levels of histones increase chromosome loss and enhance DNA damage sensi‐ tivity [57, 89]. Defects in histone degradation during replication stress or DNA damage se‐ verely decreases cell viability [84]. Free histones show electrostatic interactions with some cellular macromolecules carrying the opposite charge such as RNA molecules. Additionally, an excess of free histones can saturate and inhibit the activity of some histone modifying en‐ zymes, and change the expression pattern of almost 240 genes [93]. Two different studies in the yeasts *S. cerevisiae* and *Schizosaccharomyces pombe* have demonstrated the importance of a correct balance between histone H3 and the centromeric variant Cse4 (CENPA) for efficient chromosome segregation. H3 can compete with Cse4 in the assembly of centromeric chro‐ matin and this competition largely depends on a correct balance between levels of H3 and H4 [52, 53]. Cells must therefore not only prevent the accumulation of free histones but also ensure a correct homeostasis between canonical and other histone variants. Once cells have decided to initiate replication, any problem that unbalances replication fork progression with histone levels can potentially lead to an increase in the abundance of free histones. In order to prevent this, there is an additional pathway linked to the DDR able to block histone

synthesis under DNA replication stress conditions or replication fork arrest.

**4.2. The DNA Damage Response (DDR): Coupling DNA and histone synthesis**

DDR is probably one of the most well characterized checkpoints in the cell and is normally activated whenever a cell senses DNA damage. Activation leads to the sequential action of a cascade of kinases that block or delay cell cycle progression to allow the cell to correct the damage. If damage cannot be repaired, human cells enter the apoptosis program and die [94]. Proper functioning of this pathway is essential for genome integrity and mutations in most of the branches of this path are linked to cancer and other diseases. DDR is able to block cells at G1, S and G2/M [95]. In human cells, two kinases ATM and ATR (Tel1 and Mec1 respectively in *S. cerevisiae*) play a major role in the activation of the DDR. ATM has been directly involved in the activation of a mechanism that ultimately leads to repression

In human cells, activation of histone gene transcription requires NPAT phosphorylation by the cyclin E-CDK2 complex at the beginning of S-Phase. Activation of NPAT is essential for S-phase progression and histone expression. Repression of histone synthesis upon DNA damage requires the activation of ATM, which leads to the sequential activation of p53 and then p21. p21 is able to block the activity of the cyclin E-CDK2 complex. Inhibition of this complex leads to a progressive dephosphorylation of NPAT, which no longer localizes to

tions that break this bidirectional relationship.

**4.1. Harmful effects of free histones**

388 The Mechanisms of DNA Replication

of histone expression.

Regulation of histone levels upon DNA damage in *S. cerevisiae* shows some common regula‐ tory elements with human cells, and suggests the existence of a conserved mechanism. Posttranscriptional regulation also depends on the Lsm1-7 complex. It is not clear how this complex recognizes histone mRNAs but it could be related to the poly (A) tail-length (see post-transcriptional regulation of histones). Post-translational regulation by the Rad53 his‐ tone degradation pathway has not been directly addressed during the DDR, but taking into account the role of this protein in both pathways, it is reasonable to think that Rad53 could be important to destroy the population of translated histones when replication is halted. There are no NPAT homologues described in yeast and negative regulation during the DDR depends on the integrity of the HIR complex. The repressive structure formed to block tran‐ scription on histone promoters also requires Asf1 and Rtt106 among others. Although there is a lot of information about the formation of the repressive structure created at the promot‐ er [97], the first steps by which DNA damage triggers histone repression remain largely un‐ known. There is some data nevertheless that suggest that Asf1 and Rad53 could play a role in this process.

Asf1 is able to form a very stable complex with Rad53. Upon activation of the DNA damage response, Mec1 phosphorylates Rad53 and this phosphorylation dissociates the stable Asf1- Rad53 complex. This mechanism has been linked to a possible connection between check‐ point activation and DNA repair since Asf1 plays a role in chromatin remodeling during DNA repair [98]. Rad53 can also be found in a hypophosphorylated form in normal condi‐ tions during G1, G2 and M, stages at which histone transcription is repressed. This phos‐ phorylation seems to depend on Cdc28, the yeast functional homologue of human CDK1 and CDK2. Asf1 is able to co-immunoprecipitate with all the subunits of the HIR complex. This complex has been related to replication-independent nucleosome assembly and *in vitro* data prove that it is able to assembly nucleosomes to a DNA template [99]. Mutants lacking Asf1 have higher levels of histone mRNA and show defects in S-phase progression [100]. We have recently seen in our lab that mutants lacking the kinase activity of Rad53 also have enhanced levels of these mRNAs (unpublished results). Taking into account the close rela‐ tionship that Asf1 plays with both Rad53 and the HIR complex, it is possible to think that the dissociation of Rad53 and Asf1 during DNA damage could be important for the efficient repression of histone transcription.

#### **4.3. Generation of free histones in the cell**

How can free histones be generated during a normal cell cycle? Taking into account the tight regulation of histone levels, such situations may seem unlikely. There are two scenarios in which it is possible to think that histone supply and DNA replication can be unbalanced during a normal cell cycle. In the first one, this situation could arise from differences be‐ tween the rate of DNA synthesis and histone supply during replication. Early S-phase cells use more replication forks than late S-phase cells [101, 102] and lesions in DNA or replica‐ tion stress also affect the speed of the replication fork [103-106]. The second scenario in which a cell can encounter free histones would take place during the G2 stage of the cell cy‐ cle. Given the importance of a balanced ratio between histone H3 and CENPA in chromo‐ some segregation, once cells have finished replication, all free histones that are not positioned should be quickly degraded. It is possible to think that an imbalance between these two types of histones could sometimes take place in actively replicating cells and opens a simple explanation to why most cancer cells have a high incidence rate of chromo‐ some loss [107].

#### **4.4. Transcription as a source of free histones**

Transcription of a chromatin template also requires nucleosomes to be disassembled and re‐ assembled after the passage of RNA polymerase II (RNA Pol II). Outside of the S-phase, transcribed chromatin is probably the major potential source of free histones. These free his‐ tones could arise due to minor imbalances between histone supply and demand during chromatin reassembly. One very well described essential factor involved in RNA pol II tran‐ scription is the FACT complex [108, 109]. This complex is able to stimulate RNA Pol II-de‐ pendent transcription elongation through chromatin *in vitro* [110, 111] and also *in vivo* [112-114]. FACT is able to bind H3/H4 tetramers and H2A/H2B dimers [115, 116] and it has been shown that the integrity of at least one of its subunits, Spt16, is important for an effi‐ cient reassembly of the original H3 and H4 histones evicted during transcription [117]. Our group, in collaboration with others, demonstrated two years ago that dysfunction in chro‐ matin reassembly during transcription due to defects in the Spt16 protein generates an accu‐ mulation of free histones in yeast. Combination of this mutant with a kinase dead version of Rad53 *(rad53K227A)*, unable to efficiently degrade histones, increases the accumulation of free histones and greatly impairs viability of this mutant in a checkpoint-independent way [118]. Deletion of one of the two-histone clusters for H2A-H2B expression is able to partially sup‐ press the growth defect of this mutant and increasing H2A-H2B expression has the opposite effect. There is a strong correlation between histone levels and viability defects of the chro‐ matin reassembly mutant of Spt16. This defect is not exclusive for Spt16, since Spt6, another chromatin remodeling factor involved in H3-H4 repositioning during transcription, also has a strong negative interaction with *rad53K227A*. Chromatin reassembly defects can lead to the generation of free histones evicted from chromatin during transcription, a new source of his‐ tones potentially toxic for the cell. Rad53 negatively interacts with many different proteins involved in chromatin-related processes and could have an important function in maintain‐ ing chromatin structure in yeast [119]. Some of these interactions are with factors that have only been exclusively involved to date in chromatin related processes during transcription [118, 119] suggesting that Rad53 could play an important role in the degradation of histones when chromatin is not correctly reassembled during transcription.

**4.3. Generation of free histones in the cell**

390 The Mechanisms of DNA Replication

**4.4. Transcription as a source of free histones**

some loss [107].

How can free histones be generated during a normal cell cycle? Taking into account the tight regulation of histone levels, such situations may seem unlikely. There are two scenarios in which it is possible to think that histone supply and DNA replication can be unbalanced during a normal cell cycle. In the first one, this situation could arise from differences be‐ tween the rate of DNA synthesis and histone supply during replication. Early S-phase cells use more replication forks than late S-phase cells [101, 102] and lesions in DNA or replica‐ tion stress also affect the speed of the replication fork [103-106]. The second scenario in which a cell can encounter free histones would take place during the G2 stage of the cell cy‐ cle. Given the importance of a balanced ratio between histone H3 and CENPA in chromo‐ some segregation, once cells have finished replication, all free histones that are not positioned should be quickly degraded. It is possible to think that an imbalance between these two types of histones could sometimes take place in actively replicating cells and opens a simple explanation to why most cancer cells have a high incidence rate of chromo‐

Transcription of a chromatin template also requires nucleosomes to be disassembled and re‐ assembled after the passage of RNA polymerase II (RNA Pol II). Outside of the S-phase, transcribed chromatin is probably the major potential source of free histones. These free his‐ tones could arise due to minor imbalances between histone supply and demand during chromatin reassembly. One very well described essential factor involved in RNA pol II tran‐ scription is the FACT complex [108, 109]. This complex is able to stimulate RNA Pol II-de‐ pendent transcription elongation through chromatin *in vitro* [110, 111] and also *in vivo* [112-114]. FACT is able to bind H3/H4 tetramers and H2A/H2B dimers [115, 116] and it has been shown that the integrity of at least one of its subunits, Spt16, is important for an effi‐ cient reassembly of the original H3 and H4 histones evicted during transcription [117]. Our group, in collaboration with others, demonstrated two years ago that dysfunction in chro‐ matin reassembly during transcription due to defects in the Spt16 protein generates an accu‐ mulation of free histones in yeast. Combination of this mutant with a kinase dead version of Rad53 *(rad53K227A)*, unable to efficiently degrade histones, increases the accumulation of free histones and greatly impairs viability of this mutant in a checkpoint-independent way [118]. Deletion of one of the two-histone clusters for H2A-H2B expression is able to partially sup‐ press the growth defect of this mutant and increasing H2A-H2B expression has the opposite effect. There is a strong correlation between histone levels and viability defects of the chro‐ matin reassembly mutant of Spt16. This defect is not exclusive for Spt16, since Spt6, another chromatin remodeling factor involved in H3-H4 repositioning during transcription, also has a strong negative interaction with *rad53K227A*. Chromatin reassembly defects can lead to the generation of free histones evicted from chromatin during transcription, a new source of his‐ tones potentially toxic for the cell. Rad53 negatively interacts with many different proteins involved in chromatin-related processes and could have an important function in maintain‐ ing chromatin structure in yeast [119]. Some of these interactions are with factors that have

#### **4.5. Can the state of chromatin influence the decision of cells to initiate DNA replication?**

Histones are able to affect DNA replication right from the beginning; the state of the chro‐ matin influences the timing and organization of origin firing. Replication fork progression also depends on the correct histone deposition behind the replication machinery, since de‐ fects in CAF1 lead to checkpoint activation and block cells in S-phase. The state of chromatin is able therefore to influence DNA replication. Work done in our lab, suggests that chroma‐ tin state might also influence the decision of cells to enter or not replication during the G1/S transition in the *S. cerevisiae*.

The commitment to a new round of cell division takes place towards the end of the G1 phase of the cell cycle in a process called START in yeast, and Restriction Point in mammals [120]. In yeast, this is the main regulatory event of the G1 phase of the cell cycle and in‐ volves an extensive transcriptional program driven by transcription factors SBF (Swi4-Swi6) and MBF (Mbp1-Swi6) [121, 122]. MBF and SBF activation depends on the cyclin/cyclin-de‐ pendent-kinase (CDK) complex Cln3-Cdc28. This complex phosphorylates Whi5, the nega‐ tive regulator of START, thus promoting its release from SBF (Swi4-Swi6). Activation of MBF-dependent transcription by Cln3-Cdc28 acts through a mechanism independent of Whi5, involving the phosphorylation of Mbp1 [123]. Activation of these two complexes re‐ sults in the accumulation of G1 (Cln1 and Cln2) and the early S-phase cyclins (Clb5 and Clb6), which promote in last term S-phase entry [124]. The kinase activity of Cln1,2-Cdc28 triggers the degradation of cyclin-dependent kinase inhibitor Sic1 which no longer inhibits the S phase-promoting complex Clb5,6-Cdc28 [125, 126].

FACT plays a role in maintaining the integrity of the chromatin structure during transcrip‐ tion [127-129] but has also been related to a G1/S cell cycle defect in yeast in a genetic screen to identify *cdc* (cell division cycle) mutants. This cell cycle defect had been linked initially to a general transcription defect of the three G1 cyclins Cln1-Cln3 [130] and later, to a possible important role of FACT in the transcription of *CLN1* and *CLN2* [131]. We recently described that this G1 defect is also due to a transcriptional downregulation of the cyclin Cln3 at the promoter level. Surprisingly, FACT seems not to be directly involved in the transcriptional regulation of this cyclin, since it is not recruited to the promoter at START when *CLN3* levels are maximal (D. Stillman unpublished results). One rather unexpected but interesting result is that this cell cycle defect shows a direct correlation with histone levels. Decreasing the H2A-H2B histone pool diminishes the cell cycle accumulation of this mutant while blocking the efficient degradation of histones has an additive effect. This defect is not exclusive for FACT mutants, since an Spt6 mutant also shows cell cycle defects at the G1/S transition [118]. Moreover, in yeast, a structural mutant of histone H4 in a region important for the in‐ teraction between the H3-H4 tetramer and the two H2A-H2B dimers completely mimics the cell cycle defects of the Spt16 mutant [132]. Defects in the chromatin structure seem to be connected somehow to the G1/S transition. Our group has speculated that cells might be able to sense the chromatin state before entering a new round of replication. This mecha‐ nism would act at least in part through a transcriptional repression of the cyclin Cln3 mRNA. Although our first results pointed out that free histones could be the signal that trig‐ gers this G1/S transition defect, new results obtained by our lab show that this regulation could be more complex and also involve the chromatin structure itself (unpublished results).

### **5. Conclusion**

In eukaryotes and also some archaebacteria, DNA forms a nucleoprotein complex called chromatin, which allows extensive compaction of genomic DNA in the limited space of the nucleus. This traditional view of chromatin as simple building-bricks has substantially changed since the nucleosome hypothesis was proposed [133, 134]. Cells have evolved a unique and complex machinery to cope with the fact that most of the processes involving DNA are going to need to interact with and probably modify chromatin first. Chromatin acts as a new step of regulation and carries an epigenetic specific code that in some cases can be as important for the cell as the one contained on DNA. In addition, cells must also care‐ fully balance the levels of histones during chromatin formation to avoid the generation of free histones, in order to prevent their deleterious effects.

### **Acknowledgments**

We thank Akash Gunjan´s and Vincent Geli´s labs for our fruitful collaboration. We would also like to thank people in the lab for such a wonderful work environment and to all those people who are always there to make things easier. The Spanish Ministry of Education and Science (grant BFU2007-67575-C03-02/BMC), the Andalusian Government (grant P07- CVI02623) and the European Union (FEDER) have supported this work. D.M. was covered by a F.P.I. fellowship from the Regional Andalusian Government, and M. M-H. and L.D-R., by fellowships from the Spanish Ministry of Education and Science.

#### **Author details**

Douglas Maya\* , Macarena Morillo-Huesca, Lidia Delgado Ramos, Sebastián Chávez and Mari-Cruz Muñoz-Centeno

\*Address all correspondence to: dmaya@us.es

Department of Genetics, Faculty of Biology, University of Seville, Spain

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able to sense the chromatin state before entering a new round of replication. This mecha‐ nism would act at least in part through a transcriptional repression of the cyclin Cln3 mRNA. Although our first results pointed out that free histones could be the signal that trig‐ gers this G1/S transition defect, new results obtained by our lab show that this regulation could be more complex and also involve the chromatin structure itself (unpublished results).

In eukaryotes and also some archaebacteria, DNA forms a nucleoprotein complex called chromatin, which allows extensive compaction of genomic DNA in the limited space of the nucleus. This traditional view of chromatin as simple building-bricks has substantially changed since the nucleosome hypothesis was proposed [133, 134]. Cells have evolved a unique and complex machinery to cope with the fact that most of the processes involving DNA are going to need to interact with and probably modify chromatin first. Chromatin acts as a new step of regulation and carries an epigenetic specific code that in some cases can be as important for the cell as the one contained on DNA. In addition, cells must also care‐ fully balance the levels of histones during chromatin formation to avoid the generation of

We thank Akash Gunjan´s and Vincent Geli´s labs for our fruitful collaboration. We would also like to thank people in the lab for such a wonderful work environment and to all those people who are always there to make things easier. The Spanish Ministry of Education and Science (grant BFU2007-67575-C03-02/BMC), the Andalusian Government (grant P07- CVI02623) and the European Union (FEDER) have supported this work. D.M. was covered by a F.P.I. fellowship from the Regional Andalusian Government, and M. M-H. and L.D-R.,

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by fellowships from the Spanish Ministry of Education and Science.

Department of Genetics, Faculty of Biology, University of Seville, Spain

**5. Conclusion**

392 The Mechanisms of DNA Replication

**Acknowledgments**

**Author details**

Mari-Cruz Muñoz-Centeno

\*Address all correspondence to: dmaya@us.es

Douglas Maya\*


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