**3. Affinity maturation strategies for recombinant antibodies**

Recombinant antibodies obtained via combinatorial library technology from naïve or synthetic libraries have the advantage of increased diversity as a result of the large repertoire of the antibody genes. Antibodies isolated from combinatorial libraries against their respective targets sometimes may not exhibit the desired specificity and affinity. The increased affinity of an antibody is important to enhance its pharmacokinetics, efficacy and safety profile by enhancing the binding strength and function of an antibody [75]. Such optimizations can be achieved either by *in vitro* or *in vivo* affinity maturation strategies.

#### **3.1.** *In vitro* **approaches toward affinity maturation of antibodies**

library are able to achieve picomolar range when tested on different target molecules [63]. The latest optimized version, HuCAL PLATINUM® has a more advantageous design focusing on the optimization of CDR3 sequences in the modular sequence in order to yield antibodies with improved folding and enhanced binding [70]. The optimization includes avoiding N-glycosylation sites and unproductive sequences to maximize the sequence space and availability. In addition, the library is improved to enhance antibody expression in both bacterial and mammalian expression systems. Sequence optimization on nucleotides has been extensively carried out during library construction, therefore Fab fragments and IgG formats can be expressed optimally in both bacterial and mammalian systems, respectively. The resulting

The principle of the n-CoDeR® library is based on the recombination of a single framework with multiple CDRs from non-immunized donors to generate functional diversity [71]. This approach allows the retrieval of CDR loops from immunoglobulin genes from different

while maintaining reactivity and functionality of the antibody fragments [72]. The underlying concept of constructing the n-CoDeR® library is the amplification of desired CDR loops from immunoglobulin cDNA with overlap extension and assembly being performed to place the CDRs into the single framework [73]. The use of CDR loops originating from the human immune system is said to be remarkable as the sequences obtained have undergone *in vivo* processing, thus such sequences are said to have undergone proof-reading and the functionality has been confirmed [74]. The resulting genetic diversity of this library is remarkably enormous

), and has the potential to yield diversities equaling the human immune system [56].

This library appears to be a suitable candidate for therapeutic and diagnostic applications as it can generate functional antibody fragments against many types of antigens. Initially the approach of using a single framework to present various types of CDR loops was seemed risky due to the limitation in capacity. It was later proved to be successful with the isolation of antibodies specific to various types of antigens reaching affinities in the sub-nanomolar range. Another benefit afforded by this approach was the ability to select a single framework that can customize desirable characteristics and properties, as well as ensuring that antibodies can be generated which can be produced and folded in good condition [56]. Antibodies harnessed from the n-CoDeR® library are potentially advantageous for therapeutic purpose as they demonstrated a lower number of T-cell epitopes than normal antibodies. It indicates that self-reactivity is therefore circumvented and immunoge-

**3. Affinity maturation strategies for recombinant antibodies**

Recombinant antibodies obtained via combinatorial library technology from naïve or synthetic libraries have the advantage of increased diversity as a result of the large repertoire of

scaffold,

germline origins. All CDR loops are successfully recombined into one single VH-V<sup>L</sup>

library offers higher diversity than the HuCAL GOLD® library [64, 70].

*2.3.2. Case study of synthetic antibody libraries: n-CoDeR®*

(2 × 109

26 Antibody Engineering

nicity issues are reduced [72].

There are several strategies that have been used to perform *in vitro* affinity maturation to improve recombinant antibody molecules. Mutagenesis is widely employed to introduce mutations into antibody sequences. Sequences of antibody are diversified by random mutations via methods such as error-prone PCR or through site-directed mutations, where mutations are assigned to specific positions in CDRs or framework regions as well as mutational hot spots by using PCR and degenerate primers [76]. In addition, *de novo* synthesis of DNA offers the most straightforward modification procedure to further diversify the antibody sequences as a whole.

**Random mutagenesis** is a non-systematic mutagenesis method that can be performed in the absence of information regarding the importance of structures and residues that contribute to antigen-antibody binding as well as affinity maturation of antibody [77]. The method introduces point mutations into antibody genes in a random fashion. The mechanisms involves: (1) transitions, where a purine or pyrimidine is substituted by another purine or pyrimidine, (2) transversions, where a purine is substituted by a pyrimidine, or *vice versa*, (3) deletions of one or more nucleotides from the gene sequence, (4) insertions of one or more nucleotides into a gene sequence, (5) inversions where double-stranded DNA segments of two base pairs or longer is rotated at 180° [78].

Error-prone PCR is a universal method used for the introduction of random mutations by capitalizing on the natural error rate of a low fidelity DNA polymerase, for example *Taq* polymerase that lacks 3′ to 5′ proofreading activity. Several parameters during PCR amplification govern the error rate of DNA polymerases in order to create ideal mismatches in the amplified product. The manipulation of the enzyme's fidelity can be performed by varying several parameters like: (1) concentration of *Taq* DNA polymerase, (2) concentration of divalent cations (Mn2+ and Mg2+), (3) concentration of deoxyribonucleoside triphosphates (dNTPs), (4) polymerase extension time and the (5) number of PCR cycles [79, 80]. Upon amplification, the product must be ligated to a suitable plasmid and an additional step is required to recover the transformants that consist of the mutations. Error-prone PCR is a robust technique, whereby it can only introduce limited amount of base substitutions into the gene sequence. Therefore it is very useful to identify amino acid positions that are associated with function, affinity and specificity of antibodies for the method to be applied on [1]. The resulting libraries consist of a large amount of A to G and T to C transitions, thus causing high GC content amplification bias. This limitation can be circumvented by the addition of unbalanced ratios of nucleotides to reduce the amplification bias. A commercial DNA polymerase, Mutazyme® was introduced for error-prone PCR with reduced mutational bias which overcomes the issue of preferential nucleotide base selection by *Taq* DNA polymerase during amplification [78]. Error-prone PCR has been performed across the entire coding region to promote enhanced binders by the introduction of additional interacting residues between ligands and targets, altering the three dimensional structure of the target contact regions or promoting the thermal stability of ligands [53]. This method is suitable for use in ribosome, mRNA, and DNA displays whereby PCR amplification step is required after each round of selection. Additional mutations can be introduced to potential binders during this stage and can be characterized in the following round of selection. This approach was successfully used in combination with DNA shuffling for the selection and affinity maturation of an anti-fluorescein scFv which achieve an affinity of 100 fM from a 10<sup>7</sup> yeast display library [81]. Another variant of error-prone PCR applies isothermal rolling circle amplification for gene diversification. It amplifies a circular DNA template by rolling circle mechanism, generating single-stranded DNA comprising of multiple tandem repeats [82]. To generate a randomly mutated sequence library, a wild-type sequence can be introduced into a plasmid followed by isothermal rolling circle amplification under error-prone conditions [78, 83].

polymerase I and is ligated with T4 DNA ligase. The resulting combination of mutant and wild-type DNA is produced when the heteroduplex DNA is transfected into competent *E. coli* [85]. Kunkel mutagenesis uses a circular, single-stranded DNA (ssDNA) template that incorporates uracil as template. The ssDNA is then annealed to the mutagenic primers to generate double-stranded DNA (dsDNA) that consists of the mutation. The dsDNA is then transfected into *E. coli* where the bacterial repair mechanisms will remove the parent strand

High Affinity Maturated Human Antibodies from Naïve and Synthetic Antibody Repertoires

http://dx.doi.org/10.5772/intechopen.71664

29

Mutagenesis on a single-stranded DNA template (ssDNA) is labor intensive because the template requires subcloning and ssDNA rescue. Therefore, several commercial kits are available that utilizes double-stranded DNA (dsDNA) as template for site-directed mutagenesis with mutagenic primers. The QuickChange™ system (Stratagene) uses a pair of complementary oligonucleotides (forward and reverse) that consists of the desired mutations to amplify the whole plasmid with high fidelity polymerase, followed by removal of the parental DNA using *Dpn 1* endonuclease. The GeneTailor™ system (Invitrogen) is somewhat similar to QuickChange™, however it requires DNA methylase to methylate the DNA template prior to amplification. The GeneEditor™ system requires multiple transformations and ampicillin resistance cloning vectors for selecting mutants that has undergone mutagenesis [88, 90]. A major convenience for the GeneTailor™ and QuickChange™ system is the ability to carry out the mutagenesis without requiring additional vectors, host strains,

Another variant of site-directed mutagenesis is a PCR-driven method termed as overlap extension PCR. This technique employs PCR to generate modified genes from cloned DNA with just a few simple steps. The segments of a target gene are amplified from a DNA template using two flanking master primers and two internal primers. The internal primers consist of the desired mutation and overlapping nucleotide sequences. Two rounds of PCR are carried out, first by amplifying the target genes with their respective pair of primers to create two gene fragments that share some overlapping sequences at the 3′ end. Subsequently, these double-stranded duplexes are denatured and annealed, resulting in two heteroduplexes with each strand consisting of the mutated site. Then DNA polymerase functions to extend the overlapping ends of each heteroduplexes. A second PCR is done with the use of two flanking master primers to amplify the entire modified gene [90, 91]. This method was recently employed by Kitzman et al. [92] to create massive single amino acid mutagenesis in a parallel fashion coupled with microarray-based DNA synthesis technology. This is particularly useful

The increased understanding of molecular biology and specific functions of molecular biology enzymes has allowed the introduction of different approaches for mutagenesis. The combination of the different function of various enzymes has been utilized successfully to carry out directed evolution of antibody genes. Lambda exonuclease in nature functions to assist the repair of dsDNA breaks of viral DNA. It is a highly processive 5′→3′ dsDNA exonuclease which selectively degrades the phosphorylated chain of a duplex DNA to yield mononucleotides

(uracil-strand) while the recombinant clones predominate and propagate [88, 89].

or restriction sites.

for assessing and screening of variants in libraries.

Recombination provides another approach for gene modification and diversification. Mutational rearrangements are highly advantageous to identify and obtain beneficial mutational combinations otherwise absent in nature. Chain shuffling is a process that serves as a "mix and match" system to increase gene repertoires. Chain shuffling allows for one of the two antibody chains (heavy or light chains) to be paired with a repertoire of partner chains to generate a secondary library in order to produce higher affinity antibodies. The domain shuffling is a useful affinity maturation tool for antibodies as it mimics the *in vivo* SHM process [84]. While DNA shuffling generates chimeric libraries through random fragmentation of a pool of similar genes, reassembly of the fragments will result in template switching giving rise to sequence diversity. Application of DNA shuffling to different antibody genes leads to exchange of CDRs and frameworks. Affinity maturation can be achieved by using a single variable heavy chain domain or light chain domain from a known binder and mixing it with an array of diverse different heavy chain or light chain domains for an improved affinity.

**Site-directed mutagenesis** involves *in vitro* gene modifications that are targeted at a specific genetic locus or a segment of DNA sequence to study the sequence-structure-function of a gene candidate [85]. However, site-saturating mutagenesis will substitute specific sites against all possible amino acid residues. Hence, the importance of a specific amino acid residue towards the function of an antibody can be elucidated through this focused mutagenesis method [78, 86]. This can be applied for stability engineering of antibodies by determining the influence of different amino acids at strategic positions along the antibody structure. Sitedirected mutagenesis can be performed through several different approaches. The availability of restriction nucleases and DNA ligases allows easy incorporation of mutagenic sequences into templates to construct recombinant DNAs [87]. The rapid development of oligonucleotide synthesis has also contributed to oligonucleotide-mediated mutagenesis method. Such approach is designed to consist of internal mismatches that complement the template DNA for directing point mutations or multiple mutations. For instance, a mutagenic primer anneals to a single-stranded DNA template, followed by extension with Klenow fragment of DNA polymerase I and is ligated with T4 DNA ligase. The resulting combination of mutant and wild-type DNA is produced when the heteroduplex DNA is transfected into competent *E. coli* [85]. Kunkel mutagenesis uses a circular, single-stranded DNA (ssDNA) template that incorporates uracil as template. The ssDNA is then annealed to the mutagenic primers to generate double-stranded DNA (dsDNA) that consists of the mutation. The dsDNA is then transfected into *E. coli* where the bacterial repair mechanisms will remove the parent strand (uracil-strand) while the recombinant clones predominate and propagate [88, 89].

PCR has been performed across the entire coding region to promote enhanced binders by the introduction of additional interacting residues between ligands and targets, altering the three dimensional structure of the target contact regions or promoting the thermal stability of ligands [53]. This method is suitable for use in ribosome, mRNA, and DNA displays whereby PCR amplification step is required after each round of selection. Additional mutations can be introduced to potential binders during this stage and can be characterized in the following round of selection. This approach was successfully used in combination with DNA shuffling for the selection and affinity maturation of an anti-fluorescein scFv which achieve

applies isothermal rolling circle amplification for gene diversification. It amplifies a circular DNA template by rolling circle mechanism, generating single-stranded DNA comprising of multiple tandem repeats [82]. To generate a randomly mutated sequence library, a wild-type sequence can be introduced into a plasmid followed by isothermal rolling circle amplification

Recombination provides another approach for gene modification and diversification. Mutational rearrangements are highly advantageous to identify and obtain beneficial mutational combinations otherwise absent in nature. Chain shuffling is a process that serves as a "mix and match" system to increase gene repertoires. Chain shuffling allows for one of the two antibody chains (heavy or light chains) to be paired with a repertoire of partner chains to generate a secondary library in order to produce higher affinity antibodies. The domain shuffling is a useful affinity maturation tool for antibodies as it mimics the *in vivo* SHM process [84]. While DNA shuffling generates chimeric libraries through random fragmentation of a pool of similar genes, reassembly of the fragments will result in template switching giving rise to sequence diversity. Application of DNA shuffling to different antibody genes leads to exchange of CDRs and frameworks. Affinity maturation can be achieved by using a single variable heavy chain domain or light chain domain from a known binder and mixing it with an array of

**Site-directed mutagenesis** involves *in vitro* gene modifications that are targeted at a specific genetic locus or a segment of DNA sequence to study the sequence-structure-function of a gene candidate [85]. However, site-saturating mutagenesis will substitute specific sites against all possible amino acid residues. Hence, the importance of a specific amino acid residue towards the function of an antibody can be elucidated through this focused mutagenesis method [78, 86]. This can be applied for stability engineering of antibodies by determining the influence of different amino acids at strategic positions along the antibody structure. Sitedirected mutagenesis can be performed through several different approaches. The availability of restriction nucleases and DNA ligases allows easy incorporation of mutagenic sequences into templates to construct recombinant DNAs [87]. The rapid development of oligonucleotide synthesis has also contributed to oligonucleotide-mediated mutagenesis method. Such approach is designed to consist of internal mismatches that complement the template DNA for directing point mutations or multiple mutations. For instance, a mutagenic primer anneals to a single-stranded DNA template, followed by extension with Klenow fragment of DNA

diverse different heavy chain or light chain domains for an improved affinity.

yeast display library [81]. Another variant of error-prone PCR

an affinity of 100 fM from a 10<sup>7</sup>

28 Antibody Engineering

under error-prone conditions [78, 83].

Mutagenesis on a single-stranded DNA template (ssDNA) is labor intensive because the template requires subcloning and ssDNA rescue. Therefore, several commercial kits are available that utilizes double-stranded DNA (dsDNA) as template for site-directed mutagenesis with mutagenic primers. The QuickChange™ system (Stratagene) uses a pair of complementary oligonucleotides (forward and reverse) that consists of the desired mutations to amplify the whole plasmid with high fidelity polymerase, followed by removal of the parental DNA using *Dpn 1* endonuclease. The GeneTailor™ system (Invitrogen) is somewhat similar to QuickChange™, however it requires DNA methylase to methylate the DNA template prior to amplification. The GeneEditor™ system requires multiple transformations and ampicillin resistance cloning vectors for selecting mutants that has undergone mutagenesis [88, 90]. A major convenience for the GeneTailor™ and QuickChange™ system is the ability to carry out the mutagenesis without requiring additional vectors, host strains, or restriction sites.

Another variant of site-directed mutagenesis is a PCR-driven method termed as overlap extension PCR. This technique employs PCR to generate modified genes from cloned DNA with just a few simple steps. The segments of a target gene are amplified from a DNA template using two flanking master primers and two internal primers. The internal primers consist of the desired mutation and overlapping nucleotide sequences. Two rounds of PCR are carried out, first by amplifying the target genes with their respective pair of primers to create two gene fragments that share some overlapping sequences at the 3′ end. Subsequently, these double-stranded duplexes are denatured and annealed, resulting in two heteroduplexes with each strand consisting of the mutated site. Then DNA polymerase functions to extend the overlapping ends of each heteroduplexes. A second PCR is done with the use of two flanking master primers to amplify the entire modified gene [90, 91]. This method was recently employed by Kitzman et al. [92] to create massive single amino acid mutagenesis in a parallel fashion coupled with microarray-based DNA synthesis technology. This is particularly useful for assessing and screening of variants in libraries.

The increased understanding of molecular biology and specific functions of molecular biology enzymes has allowed the introduction of different approaches for mutagenesis. The combination of the different function of various enzymes has been utilized successfully to carry out directed evolution of antibody genes. Lambda exonuclease in nature functions to assist the repair of dsDNA breaks of viral DNA. It is a highly processive 5′→3′ dsDNA exonuclease which selectively degrades the phosphorylated chain of a duplex DNA to yield mononucleotides and ssDNA. A strategy that takes advantage of this feature of lambda exonuclease was applied for antibody gene mutagenesis. The formed ssDNA template will function as the template for *in vitro* antibody gene recombination. The ssDNA template is then hybridized with degenerate oligonucleotides and treated with Klenow Fragment to generate dsDNA templates of the hybridized products. This will result in a final dsDNA template that has incorporated the diversification introduced by the degenerate oligos at a specific site of the antibody gene. The method was successfully applied to carry out chain shuffling and mutagenesis of antibody clones [93]. However, a major bottleneck with these methods is their inability to allow directed mutagenesis with codon specificity.

molecular technologies has allowed the introduction of various approaches for gene modification. The design of the framework regions in the antibody gene also plays a contributing role in the improvement of the antibody affinity. This is due to the influence of the framework genes on the stability, solubility and affinity of the antibody [104]. The framework regions mainly in the neighboring regions of the CDRs have been known to also contribute to the binding characteris-

High Affinity Maturated Human Antibodies from Naïve and Synthetic Antibody Repertoires

http://dx.doi.org/10.5772/intechopen.71664

31

Direct gene synthesis of modified sequences or *de novo* synthesis of DNA is ideal to create desired gene sequences based on iterative and comprehensive analyses with the aid of high throughput sequencing technology. There are few reasons why *de novo* synthesis of DNA is preferred in many instances. Firstly, engineering new functions normally requires great modifications to the genetic sequences therefore *de novo* synthesis is more preferred. Secondly, *de novo* gene synthesis allows specific design of DNA constructs. *De novo* synthesis enables to study the influence of new designed sequences on particular functions of corresponding expressed recombinant antibodies. The aim is to improve or modify phenotypic features of

Lastly, targeted sequences from natural constructs are sometimes hard to access, therefore synthesis provides a more efficient alternative to retrieve the targeted sequences [105]. Currently, oligos are generated or synthesized automatically, employing solid-phase phosphoramidite chemistry. The principle behind phosphoramidite-based oligo synthesis encompasses a total of four key steps (deprotection, coupling, capping and oxidation) to add one base at a time to

Synthesis takes place individually in small columns. The purified oligos are then subjected to quality assessments. The automated process can generate up to 100 nmol of oligos at a time

Besides conventional gene synthesis from oligo fragments using column-based synthesized oligos, array-based oligos can be used for gene synthesis as well. An array-based synthesis has the advantage of high throughput synthesis. The polymer array support by Affymetrix is synthesized chemically comprising photolabile protecting groups and photolithography. The photolithographic mask is able to direct UV light over the solid substrate and selectively deprotect and activate the 5′-hydroxyl group in the growing chain, in order for free nucleotides to be incorporated into the chain. The mask is designed for exposing targeted sites on the microarray, where incorporation of nucleotides occurs while masking other non-targeted sites. The oligo fragments are directly synthesized on the support surface, and can be recovered as a heterogenous pool of sequences. Today, several technologies have surpassed the need to use the masking technique. An ink-jet-based printing developed by Agilent allows picolitres of free nucleotides and activator to be spotted on targeted sites on one array. NimbleGen Systems uses the programmed automated micromirror device to activate specific sites on the array. Furthermore, CustomArray (CombiMatrix) utilizes semiconductor-based

with low error rates in the region of one base error in 200 nucleotides [105, 106].

electrochemical acid production to deprotect desired nucleosides [107, 108].

tic of antibody clones [46].

antibodies.

**3.2.** *De novo* **synthesis of antibody genes**

a growing oligo chain attached to a solid support.

The diversification of the antibody repertoire can also be realized by *in vitro* somatic hypermutation using the AID enzyme [94]. The AID enzyme is classified in the APOBEC family of cytidine deaminases that is able to catalyze the deamination of cytidine residues to uridine residues *in vitro* only on ssDNA, giving rise to thymine residues at the end of the replication events [95].

Typically, the cytidines are targeted at the mutational hotspot motif RGYW and AGY (R =A/G, Y = C/T, W = A/T). This motif is also the preferred region for mutations during *in vivo* somatic hypermutation [96, 97]. Reports revealed that *in vitro* cytidine deamination occurs in an orientation-dependent fashion, relying on the transcription to gain access to both template and non-template strands of DNA. Nevertheless, it is capable of introducing mutations into DNA and therefore applicable for gene diversification [98, 99]. AID-mediated mutagenesis serves as a useful method to enhance antibody affinities through sequence diversity by introducing point mutations, such as single amino acid substitutions or indels (insertions, deletions) specifically on the antibody CDR sequences. Nucleotide transversions and duplications are among the most complicated to design into a library but possible with the AID-mediated mutagenesis approach [34]. The AID enzyme is capable of generating indels which are localized in CDR regions, while affinity maturation through somatic mutation further improves the antibody binding and specificity [100]. *In vitro* expression of the AID enzyme is sufficient to initiate indels, hypermutations inside the CDRs and clonal expansion that is comparable to *in vivo* events for antibody evolution.

Studies have been carried out to analyze the amino acid diversities in the germline and mature antibody sequences. It was found that the number of germline hotspots decreases in high affinity antibodies, suggesting that hotspot-based somatic mutations occurred via *in vivo* affinity maturation [101]. Through the *in vitro* randomization of these short CDR regions that somewhat mimics the natural *in vivo* SHM sequences diversity is generated and results in *in vitro* affinity maturation. These hotspots are embedded in the codons of amino acids that are directly and indirectly involved in interactions with antigens. They can serve as the mutation targets in the human genome allowing for various mutagenesis to occur with the aid of the general mutator candidates being AID enzymes or other trans-acting hypermutation factors [102].

The diversity associated with the utilization of various sequences either in the CDR or framework is directly related to the affinity of the clones generated [5, 103]. The continuous development in molecular technologies has allowed the introduction of various approaches for gene modification. The design of the framework regions in the antibody gene also plays a contributing role in the improvement of the antibody affinity. This is due to the influence of the framework genes on the stability, solubility and affinity of the antibody [104]. The framework regions mainly in the neighboring regions of the CDRs have been known to also contribute to the binding characteristic of antibody clones [46].

#### **3.2.** *De novo* **synthesis of antibody genes**

and ssDNA. A strategy that takes advantage of this feature of lambda exonuclease was applied for antibody gene mutagenesis. The formed ssDNA template will function as the template for *in vitro* antibody gene recombination. The ssDNA template is then hybridized with degenerate oligonucleotides and treated with Klenow Fragment to generate dsDNA templates of the hybridized products. This will result in a final dsDNA template that has incorporated the diversification introduced by the degenerate oligos at a specific site of the antibody gene. The method was successfully applied to carry out chain shuffling and mutagenesis of antibody clones [93]. However, a major bottleneck with these methods is their inability to allow directed

The diversification of the antibody repertoire can also be realized by *in vitro* somatic hypermutation using the AID enzyme [94]. The AID enzyme is classified in the APOBEC family of cytidine deaminases that is able to catalyze the deamination of cytidine residues to uridine residues *in vitro* only on ssDNA, giving rise to thymine residues at the end of the replication

Typically, the cytidines are targeted at the mutational hotspot motif RGYW and AGY (R =A/G, Y = C/T, W = A/T). This motif is also the preferred region for mutations during *in vivo* somatic hypermutation [96, 97]. Reports revealed that *in vitro* cytidine deamination occurs in an orientation-dependent fashion, relying on the transcription to gain access to both template and non-template strands of DNA. Nevertheless, it is capable of introducing mutations into DNA and therefore applicable for gene diversification [98, 99]. AID-mediated mutagenesis serves as a useful method to enhance antibody affinities through sequence diversity by introducing point mutations, such as single amino acid substitutions or indels (insertions, deletions) specifically on the antibody CDR sequences. Nucleotide transversions and duplications are among the most complicated to design into a library but possible with the AID-mediated mutagenesis approach [34]. The AID enzyme is capable of generating indels which are localized in CDR regions, while affinity maturation through somatic mutation further improves the antibody binding and specificity [100]. *In vitro* expression of the AID enzyme is sufficient to initiate indels, hypermutations inside the CDRs and clonal expansion that is comparable to

Studies have been carried out to analyze the amino acid diversities in the germline and mature antibody sequences. It was found that the number of germline hotspots decreases in high affinity antibodies, suggesting that hotspot-based somatic mutations occurred via *in vivo* affinity maturation [101]. Through the *in vitro* randomization of these short CDR regions that somewhat mimics the natural *in vivo* SHM sequences diversity is generated and results in *in vitro* affinity maturation. These hotspots are embedded in the codons of amino acids that are directly and indirectly involved in interactions with antigens. They can serve as the mutation targets in the human genome allowing for various mutagenesis to occur with the aid of the general mutator candidates being AID enzymes or other trans-acting hypermuta-

The diversity associated with the utilization of various sequences either in the CDR or framework is directly related to the affinity of the clones generated [5, 103]. The continuous development in

mutagenesis with codon specificity.

*in vivo* events for antibody evolution.

tion factors [102].

events [95].

30 Antibody Engineering

Direct gene synthesis of modified sequences or *de novo* synthesis of DNA is ideal to create desired gene sequences based on iterative and comprehensive analyses with the aid of high throughput sequencing technology. There are few reasons why *de novo* synthesis of DNA is preferred in many instances. Firstly, engineering new functions normally requires great modifications to the genetic sequences therefore *de novo* synthesis is more preferred. Secondly, *de novo* gene synthesis allows specific design of DNA constructs. *De novo* synthesis enables to study the influence of new designed sequences on particular functions of corresponding expressed recombinant antibodies. The aim is to improve or modify phenotypic features of antibodies.

Lastly, targeted sequences from natural constructs are sometimes hard to access, therefore synthesis provides a more efficient alternative to retrieve the targeted sequences [105]. Currently, oligos are generated or synthesized automatically, employing solid-phase phosphoramidite chemistry. The principle behind phosphoramidite-based oligo synthesis encompasses a total of four key steps (deprotection, coupling, capping and oxidation) to add one base at a time to a growing oligo chain attached to a solid support.

Synthesis takes place individually in small columns. The purified oligos are then subjected to quality assessments. The automated process can generate up to 100 nmol of oligos at a time with low error rates in the region of one base error in 200 nucleotides [105, 106].

Besides conventional gene synthesis from oligo fragments using column-based synthesized oligos, array-based oligos can be used for gene synthesis as well. An array-based synthesis has the advantage of high throughput synthesis. The polymer array support by Affymetrix is synthesized chemically comprising photolabile protecting groups and photolithography. The photolithographic mask is able to direct UV light over the solid substrate and selectively deprotect and activate the 5′-hydroxyl group in the growing chain, in order for free nucleotides to be incorporated into the chain. The mask is designed for exposing targeted sites on the microarray, where incorporation of nucleotides occurs while masking other non-targeted sites. The oligo fragments are directly synthesized on the support surface, and can be recovered as a heterogenous pool of sequences. Today, several technologies have surpassed the need to use the masking technique. An ink-jet-based printing developed by Agilent allows picolitres of free nucleotides and activator to be spotted on targeted sites on one array. NimbleGen Systems uses the programmed automated micromirror device to activate specific sites on the array. Furthermore, CustomArray (CombiMatrix) utilizes semiconductor-based electrochemical acid production to deprotect desired nucleosides [107, 108].

Nevertheless, the NimbleGen and CustomArray oligo synthesis techniques suffer high error rates when trying to generate longer and multiple oligo strands in parallel. This is due to the side reactions such as depurination and inefficient addition of nucleotides that results in unwanted substitution and indels (insertion/deletion) errors, which greatly affects the overall quality of the synthesized product. Therefore purification steps utilizing polyacrylamide gel electrophoresis and high performance liquid chromatography are essential to remove erroneous sequences upon generating the intended DNA sequences [109].

that undergo controlled synthesis allowing various applications for antibody and protein engineering. The main application focuses specifically on gene construction and editing. gBlocks are constructed using gene fragment libraries (pools of short DNA fragments that comprise 18 consecutive N bases or K (G,T) bases). The synthesized product is then subjected to various quality control tests such as capillary electrophoresis (fragment length) and mass spectrometry (sequence composition) to verify the final product and reduce potential errors. For gene editing, gBlocks can introduce modifications such as deletion or insertion on relatively short stretches of DNA fragments. The primers are designed to target the region of the gene that is to be edited. Subsequently, the region will be cleaved and replaced by the gBlock [109]. This method has allowed the design and generation of

High Affinity Maturated Human Antibodies from Naïve and Synthetic Antibody Repertoires

http://dx.doi.org/10.5772/intechopen.71664

33

Bacterial mutator strains, such as *Escherichia coli* mutants are shown to introduce random mutations, such as single-base substitutions with higher rates than wild type strains. The mutant strains are characterized by the absence of several DNA repair pathways, resulting in a high rate of mutations [78]. Affinity maturation via this approach involves two key steps; firstly, antibody genes are transformed and replicated in *E. coli* mutants to introduce random mutations. Next, screening of mutated antibody clones to identify highest affinity binders is done using display technologies. The affinity maturation process requires several rounds of mutation, selection and amplification in order to obtain high affinity mutants. Phage display technology is best coupled with *E. coli* mutator cells for *in vivo* mutation of antibody fragments due to the ease of application with phage and phagemid vectors [119]. The *E. coli* mutator cell such as *E. coli* mutD5-FIT consists of mutD mutation, F′ factor for Fd phage transfection and *supE* mutation. This mutant is able to express phage displayed antibody fragments, where antibody genes are fused to the N-terminus of gene III protein and are subsequently packaged to form a mature virus particle. Alternatively, another *E. coli* mutator strain, XLIRed carries mutD, mutL and mutS, while it is devoid of the F′ episome. This does not allow the cell to be applied for phage infection. However, this F′ deficient mutator cell can be converted to F′ mutator strain by mating with other *E. coli* strains with F′ episome [120, 121]. The choice of bacterial mutator strains are largely governed by downstream selection strategies that requires rational considerations. A human antibody fragment that targets the hapten 2-phenyl-5-oxazolone (phOx) was affinity matured by a factor 100 fold via *E. coli* mutD5 strain, whereby the mutations are extensively located in CDR loops and less in framework regions, which improve the binding affinity of the anti-

*In vivo* affinity maturation can also be performed via AID-mediated mutagenesis. It fits quite well with the robust mammalian cell display techniques for selection and affinity maturation of functional antibody clones. As an example, the cDNA of an anti-human complement protein C5 antibody is transfected into HEK293 cells together with the AID enzyme expression plasmid to initiate *in vitro* SHM. The antibody clones are identified

antibody libraries [117, 118].

body to the target [122].

**3.3.** *In vivo* **approaches towards affinity maturation of antibodies**

The generated oligo fragments obtained after conventional or area-based synthesis are then used as raw substrates to construct larger synthetic fragments (usually few hundreds of base pairs), also known as gene synthesis. Using a ligation-based approach, the complementary overlapping fragments are joined enzymatically by the thermostable DNA ligases, producing larger DNA fragments under high stringency [110]. Another approach, known as polymerase cycling assembly (PCA)-based method, utilizes polymerase to elongate the originated overlapping oligo fragments into double-stranded fragments [111]. Ligation-based synthesis offer higher stringency, therefore error in sequences is less likely to be assembled, but the oligo synthesis are costly due to synthesis of longer fragments. The longer oligonucleotides will allow for better annealing and less steps in comparison to shorter oligonucleotdies. The final step would sometimes involve an additional Polymerase chain reaction (PCR) amplification step to yield more material for cloning. On the contrary, PCA-based methods are more cost-effective as it relies on overlapping short oligo fragments (15–25 nt) per gene synthesis. However, this approach promotes higher error rates due to the lack of error elimination during hybridization [112]. Also, target diversity can be introduced at the regions where the overlapping oligo fragments hybridizes [109].

Despite the fact that concentrations of individual oligos on the array are quite low and insufficient for priming, as well as the error rates of the oligo pools are higher as the column-based methods, there are successful examples that overcome these challenges. This is done with the use of programmable DNA microchips with an array of oligonucleotides and their selection [113]. To increase the concentration of oligonucleotides for gene synthesis, amplification of oligo fragments before assembly is required. Sequence errors can be detected via hybridization of the synthetized cleaved oligonucleotides to complementary oligonucleotides spanned on a second area. Lastly error-free fragments will be assembled into full-length sequences. However, this method is not feasible for assembling a large pool of oligos because of the risk of cross-hybridization based on the huge diversity. Another approach used selective oligonucleotide pool amplification directed by predesigned barcodes to generate and assemble particular DNA fragments that are required to make a full gene before the barcodes are digested prior to full gene assembly [114]. Recently, this approach was applied to construct few scFv gene libraries with degenerate oligonucleotides synthesized on two DNA microchips in parallel [115]. The humanized anti-ErbB2 antibody (HuA21) was targeted to diversify the CDR regions via a small perturbation mutagenesis method and was validated using deep sequencing by the Illumina platform. Finally, the mutant candidates were screened by phage display to select for high affinity binders [115, 116].

gBlocks gene fragments are readily usable short-to-medium length synthesized DNA fragments that contains particular desired gene modifications. gBlocks are dsDNA blocks that undergo controlled synthesis allowing various applications for antibody and protein engineering. The main application focuses specifically on gene construction and editing. gBlocks are constructed using gene fragment libraries (pools of short DNA fragments that comprise 18 consecutive N bases or K (G,T) bases). The synthesized product is then subjected to various quality control tests such as capillary electrophoresis (fragment length) and mass spectrometry (sequence composition) to verify the final product and reduce potential errors. For gene editing, gBlocks can introduce modifications such as deletion or insertion on relatively short stretches of DNA fragments. The primers are designed to target the region of the gene that is to be edited. Subsequently, the region will be cleaved and replaced by the gBlock [109]. This method has allowed the design and generation of antibody libraries [117, 118].

#### **3.3.** *In vivo* **approaches towards affinity maturation of antibodies**

Nevertheless, the NimbleGen and CustomArray oligo synthesis techniques suffer high error rates when trying to generate longer and multiple oligo strands in parallel. This is due to the side reactions such as depurination and inefficient addition of nucleotides that results in unwanted substitution and indels (insertion/deletion) errors, which greatly affects the overall quality of the synthesized product. Therefore purification steps utilizing polyacrylamide gel electrophoresis and high performance liquid chromatography are essential to remove errone-

The generated oligo fragments obtained after conventional or area-based synthesis are then used as raw substrates to construct larger synthetic fragments (usually few hundreds of base pairs), also known as gene synthesis. Using a ligation-based approach, the complementary overlapping fragments are joined enzymatically by the thermostable DNA ligases, producing larger DNA fragments under high stringency [110]. Another approach, known as polymerase cycling assembly (PCA)-based method, utilizes polymerase to elongate the originated overlapping oligo fragments into double-stranded fragments [111]. Ligation-based synthesis offer higher stringency, therefore error in sequences is less likely to be assembled, but the oligo synthesis are costly due to synthesis of longer fragments. The longer oligonucleotides will allow for better annealing and less steps in comparison to shorter oligonucleotdies. The final step would sometimes involve an additional Polymerase chain reaction (PCR) amplification step to yield more material for cloning. On the contrary, PCA-based methods are more cost-effective as it relies on overlapping short oligo fragments (15–25 nt) per gene synthesis. However, this approach promotes higher error rates due to the lack of error elimination during hybridization [112]. Also, target diversity can be introduced at the regions where the overlapping oligo fragments hybridizes [109].

Despite the fact that concentrations of individual oligos on the array are quite low and insufficient for priming, as well as the error rates of the oligo pools are higher as the column-based methods, there are successful examples that overcome these challenges. This is done with the use of programmable DNA microchips with an array of oligonucleotides and their selection [113]. To increase the concentration of oligonucleotides for gene synthesis, amplification of oligo fragments before assembly is required. Sequence errors can be detected via hybridization of the synthetized cleaved oligonucleotides to complementary oligonucleotides spanned on a second area. Lastly error-free fragments will be assembled into full-length sequences. However, this method is not feasible for assembling a large pool of oligos because of the risk of cross-hybridization based on the huge diversity. Another approach used selective oligonucleotide pool amplification directed by predesigned barcodes to generate and assemble particular DNA fragments that are required to make a full gene before the barcodes are digested prior to full gene assembly [114]. Recently, this approach was applied to construct few scFv gene libraries with degenerate oligonucleotides synthesized on two DNA microchips in parallel [115]. The humanized anti-ErbB2 antibody (HuA21) was targeted to diversify the CDR regions via a small perturbation mutagenesis method and was validated using deep sequencing by the Illumina platform. Finally, the mutant candidates were screened by phage display to select for high affinity binders [115, 116].

gBlocks gene fragments are readily usable short-to-medium length synthesized DNA fragments that contains particular desired gene modifications. gBlocks are dsDNA blocks

ous sequences upon generating the intended DNA sequences [109].

32 Antibody Engineering

Bacterial mutator strains, such as *Escherichia coli* mutants are shown to introduce random mutations, such as single-base substitutions with higher rates than wild type strains. The mutant strains are characterized by the absence of several DNA repair pathways, resulting in a high rate of mutations [78]. Affinity maturation via this approach involves two key steps; firstly, antibody genes are transformed and replicated in *E. coli* mutants to introduce random mutations. Next, screening of mutated antibody clones to identify highest affinity binders is done using display technologies. The affinity maturation process requires several rounds of mutation, selection and amplification in order to obtain high affinity mutants. Phage display technology is best coupled with *E. coli* mutator cells for *in vivo* mutation of antibody fragments due to the ease of application with phage and phagemid vectors [119]. The *E. coli* mutator cell such as *E. coli* mutD5-FIT consists of mutD mutation, F′ factor for Fd phage transfection and *supE* mutation. This mutant is able to express phage displayed antibody fragments, where antibody genes are fused to the N-terminus of gene III protein and are subsequently packaged to form a mature virus particle. Alternatively, another *E. coli* mutator strain, XLIRed carries mutD, mutL and mutS, while it is devoid of the F′ episome. This does not allow the cell to be applied for phage infection. However, this F′ deficient mutator cell can be converted to F′ mutator strain by mating with other *E. coli* strains with F′ episome [120, 121]. The choice of bacterial mutator strains are largely governed by downstream selection strategies that requires rational considerations. A human antibody fragment that targets the hapten 2-phenyl-5-oxazolone (phOx) was affinity matured by a factor 100 fold via *E. coli* mutD5 strain, whereby the mutations are extensively located in CDR loops and less in framework regions, which improve the binding affinity of the antibody to the target [122].

*In vivo* affinity maturation can also be performed via AID-mediated mutagenesis. It fits quite well with the robust mammalian cell display techniques for selection and affinity maturation of functional antibody clones. As an example, the cDNA of an anti-human complement protein C5 antibody is transfected into HEK293 cells together with the AID enzyme expression plasmid to initiate *in vitro* SHM. The antibody clones are identified and isolated by fluorescence-activated cell sorting (FACS) with fluorescence labeled antigen followed by iterative rounds of SHM to yield high affinity antibodies [34]. In mammalian, yeast and bacterial cell surface display it is essential to select and isolate the cells that are able to produce functional antibodies. Cell-based sorting such as high throughput flow cytometry, allows high throughput screening of cells per minute, analyzing the cells according to size, granularity and binding to fluorescence labeled antigens. The utmost reason for antibody engineering is the production of human monoclonal antibody in large scale, hence, it is crucial to implement a selection towards antibody expressing mammalian cells. Mammalian display allows for screening of eukaryotes-expressed full length antibodies with correct glycosylation [18, 123].

resulting in lower binding efficiency towards DNA targets. Although TALEs require only one nucleotide for binding towards target, however the synthesis for novel TALEs is costly due to their repetitive sequences [125]. Nevertheless, these enzymes are constructed in customizable fashion to cater for the need of genome editing, as well as programming the enzymes for multiplex gene targeting [54]. Some model organisms were tested with the genome editing technologies, such as zebrafish, rats, mice, *Drosophila*, *C. elegans*. Some delivery methods of introducing these programmed enzymes into organisms are microinjections of stem cells with mRNA encoding the enzymes or direct transfection of an plasmid consisting of the enzyme

High Affinity Maturated Human Antibodies from Naïve and Synthetic Antibody Repertoires

http://dx.doi.org/10.5772/intechopen.71664

35

Naïve and synthetic human antibody repertoires are a very valuable source for the selection of antibodies against nearly any antigen. The role display technologies play in the quest to generate monoclonal antibodies from these libraries is obvious with the increasing number of

Affinity maturation of selected binders is now possible by expressing for example the AID enzyme during selection of antibodies using antibody mammalian cell surface display or by using a pool of microchip-synthesized CDRs incorporated into an antibody framework. Selection of naïve and synthetic recombinant antibodies combined with *in vitro* and *in vivo* affinity maturation techniques will have a profound effect on the generation of high affinity

The authors would like to acknowledge the support from the Malaysian Ministry of Higher Education under the Higher Institution Centre of Excellence (HICoE) Grant (Grant no. 311/

and Theam Soon Lim1,2\*

1 Institute for Research in Molecular Medicine, Universiti Sains Malaysia, Minden, Penang,

2 Analytical Biochemistry Research Centre, Universiti Sains Malaysia, Minden, Penang,

cDNA into HEK293 cells [128, 131].

antibody lead candidates going into clinical trials.

diagnostic and therapeutic human antibodies.

, Yee Siew Choong1

\*Address all correspondence to: theamsoon@usm.my

**Acknowledgements**

CIPPM/44001005).

**Author details**

Chia Chiu Lim1

Malaysia

Malaysia

**4. Conclusion**

The advancement of genome editing technologies offers a new approach to create sequence diversity. In fact, cells can repair DNA damages intrinsically by joining two ends together or filling the gap with similar sequences. However, cells can also repair the break by using a new piece of DNA that has the desired mutation. This is the basis of *in vivo* genome editing technologies today [124]. Extensive functional genomics studies helps to provide the insights required that targeted DNA double-stranded breaks (DSBs). The DSBs can induce genome editing via homologous recombination (HR) in the presence of exogenous homology repair template, as well as error-prone non-homologous end-joining repair (NHEJ) pathway in the absence of repair template. These two pathways are versatile to allow precise genome modification [125]. To date, there are four major classes of engineered DNA binding proteins to target DSBs: meganucleases, zinc finger (ZF) nucleases, transcription activator-like effectors (TALEs) and RNA-guided DNA endonuclease Cas9 [124].

Meganucleases are derived from microbial mobile genetic elements that integrate nuclease and DNA binding domains. ZF nucleases (Cys<sup>2</sup> His2 bound to a single atom of zinc) are eukaryotic transcription factors that contain the DNA binding domain and is similar to a set of three fingers, with each finger contacting with 3 nucleotides of DNA [126]. TALEs are produced naturally by *Xanthomonas* sp. that consist of DNA binding domains with 30–35 tandem repeats, with each domain recognizing a single nucleotide of DNA. Specificity of TALEs is governed by the two amino acids that are known as the repeat-hypervariable diresidues [127]. Both ZF nucleases and TALEs require *Fok*I nuclease to direct the nucleolytic activity towards the genome locus for modifications. Recently, RNA-guided DNA endonuclease Cas9 is given much more attention due to its simplicity and versatile approach towards *in vivo* genome engineering. It is derived from type II bacterial adaptive immune system. The CRISPR-Cas9 mediated immunity is explicitly explained in Yang et al. [128]. The hallmark of this system is the short RNA guide that recognizes the target DNA through base pairing, forming a RNA-DNA complex, subsequently Cas9 creates DSB on the target sequence [129, 130] and a designed DNA fragment can be specifically incorporated.

While other approaches have their own limitations, the robustness of the CRISPR-Cas9 system sheds some light on direct endogenous genome editing on virtually any organism of choice. Meganuclease lacks target specificity, which is why it is not widely employed. However, ZF domains have a tendency to crosslink with neighboring protein -domains or -complexes resulting in lower binding efficiency towards DNA targets. Although TALEs require only one nucleotide for binding towards target, however the synthesis for novel TALEs is costly due to their repetitive sequences [125]. Nevertheless, these enzymes are constructed in customizable fashion to cater for the need of genome editing, as well as programming the enzymes for multiplex gene targeting [54]. Some model organisms were tested with the genome editing technologies, such as zebrafish, rats, mice, *Drosophila*, *C. elegans*. Some delivery methods of introducing these programmed enzymes into organisms are microinjections of stem cells with mRNA encoding the enzymes or direct transfection of an plasmid consisting of the enzyme cDNA into HEK293 cells [128, 131].
