Synthetic antibodies: Concepts, potential and practical considerations

Methods 57 (2012) 486–498

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Methods journal homepage: www.elsevier.com/locate/ymeth

Review Article

Synthetic antibodies: Concepts, potential and practical considerations S. Miersch ⇑, S.S. Sidhu Banting and Best Department of Medical Research, University of Toronto, Toronto, Ontario, Canada

a r t i c l e

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Article history: Available online 27 June 2012 Keywords: Phage display Combinatorial libraries Synthetic antibodies Protein engineering

a b s t r a c t The last 100 years of enquiry into the fundamental basis of humoral immunity has resulted in the identification of antibodies as key molecular sentinels responsible for the in vivo surveillance, neutralization and clearance of foreign substances. Intense efforts aimed at understanding and exploiting their exquisite molecular specificity have positioned antibodies as a cornerstone supporting basic research, diagnostics and therapeutic applications [1]. More recently, efforts have aimed to circumvent the limitations of developing antibodies in animals by developing wholly in vitro techniques for designing antibodies of tailored specificity. This has been realized with the advent of synthetic antibody libraries that possess diversity outside the scope of natural immune repertoires and are thus capable of yielding specificities not otherwise attainable. This review examines the convergence of technologies that have contributed to the development of combinatorial phage-displayed antibody libraries. It further explores the practical concepts that underlie phage display, antibody diversity and the methods used in the generation of and selection from phage-displayed synthetic antibody libraries, highlighting specific applications in which design approaches gave rise to specificities that could not easily be obtained with libraries based upon natural immune repertories. Ó 2012 Elsevier Inc. All rights reserved.

1. Introduction The past 20 years have witnessed tremendous growth in the development and application of affinity reagents including RNA and DNA aptamers, peptides, and a variety of alternative protein scaffolds. This progress has been spurred by the maturation of display technologies [2–5], which enable the in vitro evolution and precise engineering of binding properties. Nevertheless, the original affinity agent, the antibody, still remains at the forefront of versatility and applicability. In 1975, biological research was revolutionized by the landmark discovery that the fusion of B cells with myeloma cells yields an immortalized ‘‘hybridoma’’ cell that continuously produces monoclonal immunoglobulin [6]. As a key tool in the biochemists armamentarium, a cornerstone of clinical diagnostics, and an increasingly viable alternative for treatment in the hands of clinicians, antibodies are arguably now the most important biomolecules in modern medical sciences. Despite this, commercially available antibodies are often plagued by a lack of performance and specificity [7,8], and consequently, their use is often considered to be an experiment unto itself. Unfortunately, this diverts valuable resources away from fundamental research efforts.

⇑ Corresponding author. E-mail address: [email protected] (S. Miersch). 1046-2023/$ - see front matter Ó 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ymeth.2012.06.012

Fortunately, critical advances on several fronts have enabled competitive, recombinant technologies for isolation and engineering of fully human antibodies, without the use of animals and hybridomas. As the molecular understanding of the filamentous bacteriophage lifecycle and structure evolved, it became clear that the simplicity of its single-stranded DNA (ssDNA) genome and the close coupling of viral genotype with phenotype offered means for manipulating and presenting proteins for binding selections [9,10]. Ultimately, the advent of display technologies would be met with recombinant and combinatorial gene techniques [11,12] that would merge into a powerful technology for displaying diverse libraries in a manner that allows for unprecedented exploration of protein structure and function. Simultaneously, intense enquiry aimed at determining how antibodies are capable of interacting with such a broad array of molecular entities led to insights regarding the generation [13,14], structure [15,16], composition [17,18], and molecular interactions [19] of the antigen-binding site and have informed library construction and optimization [20–23]. By combining knowledge regarding the structural and physicochemical determinants of binding with efforts to optimize vectors and display formats for the diversification and expression of antibodies [24], high quality antibody libraries that possess molecular diversity paralleling natural diversity have been made possible [25–27]. These libraries have yielded an exquisite set of molecular tools that enable the detailed characterization and modulation of

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biological function. Extending beyond conventional antibody applications (blotting, immunostaining, affinity purification, etc.), synthetic antibodies with tailored functions have been designed to be viable tools for intracellular targeting (intrabodies) [28–30], misfolded protein recognition [31], sensing protein conformation [29,32,33], in vivo homing [34,35] and even therapeutics applications. Given the vast array of potential applications for antibodies, techniques for generating recombinant antibodies with desired specificities and affinities have an enormous potential for impact on biological research. This review provides context for the development and current state of phage-displayed antibody libraries with an overview of the theoretical and practical considerations when designing and constructing libraries. It will assume a basic familiarity with the structure, function and in vivo generation of natural antibodies; while elaborating on how knowledge of the complementarity-determining regions (CDRs) is used to construct libraries of wholly synthetic antibodies. Throughout, we attempt to offer valuable insights for the successful application of phage display by emphasizing critical aspects, performance and future prospects. 2. Hybridoma technologies Hybridoma technologies were first realized in 1975, when Köhler and Milstein described a method for fusing B-lymphocytes from an immunized animal to immortal myeloma cells. Populations of fused hybrid cells could be separated to isolate cells of a single clonal specificity and used to continually produce monoclonal immunoglobulin [6]. This discovery revolutionized biochemical research and contributed to the ascendancy of antibodies to the central position they currently occupy in both basic and applied sciences. In the clinical sciences, the power of hybridoma antibody technology is evident from the growing number of therapeutics initially derived from mouse hybridomas and now approved for clinical application [1], and from the fact that hybridoma technology continues to generate valuable therapeutic leads. However, bottlenecks in the immunization process, intense manual liquid handling and characterization of antibody specificity have slowed the pace at which antibodies can be developed. Further, from a therapeutic standpoint monoclonal antibodies derived from mouse hybridoma pose specific challenges insofar as they bear the potential for immunogenicity in humans occasionally triggering hypersensitivity reactions and the generation of human anti-mouse antibodies (HAMAs). These antibodies can affect clinical outcomes by neutralizing therapeutic antibodies, increasing rates of clearance, and ultimately hampering efficacy [36–38]. Several approaches aimed at obviating this particular challenge have been devised including the development of transgenic mice that yield fully human antibodies [39], panning techniques that enable comprehensive probing of antibody specificity from peripheral blood plasma cells [40–42] and of course, in vitro technologies for humanization aimed at reducing immunogenicity [40–46]. Nevertheless, cellular stability, storage costs, the uncontrolled environment in which antigen challenge occurs, the lack of direct access to antibody genes and the inability to immunize with toxic protein targets are inherent shortfalls of hybridoma technology, which continue to provide impetus for the development of alternative strategies. As we will see, the ease of automation and storage, exquisite control over selection conditions and direct access to immunoglobulin genes (due to phenotype–genotype linkage in filamentous phage) offer unique advantages to in vitro technologies that can compete with and complement hybridoma and human antibody approaches, offering access to antibody specificities that would be otherwise unavailable by conventional methods [24,47].

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3. Phage display technologies 3.1. Phage structure and biology Although many virions (including k, T4, and T7) have been successfully employed for phage display, the non-lytic filamentous bacteriophages (M13, fd, F1) have become most commonly used for in vitro selection of interacting peptides and proteins. These viruses do not kill their host cell upon infection and thus allow for the continued propagation and growth of both host and phage such that very high viral titres are possible. These features and a detailed molecular understanding of phage structure and biology paved the way for its development as a display tool. A thorough appreciation of these details can provide important insights into the variables that influence the quality of antibody libraries, affect the success of selections, and inform troubleshooting efforts. Filamentous bacteriophages are pencil-shaped viruses of the Inovirus genus, approximately 900 nm long and 6–7 nm in diameter [48]. The viral coat, comprised of five coat proteins (pIII, pVI, pVII, PVIII, PIX), houses a single-stranded genome of 6400 base pairs, containing a total of 11 genes [49]. The phage particle is capped at one end by 3–4 copies each of the minor capsid proteins pVII and PIX [50], and at the other end, by 4–5 copies of pIII [51] and pVI [50]. The length of the phage is comprised of 2700 copies of the major coat protein pVIII, which forms a protein sheath for encapsulation and protection of the genetic material [52]. Although the molecular mechanism of phage infection remains largely elusive, pIII has been intensely studied and is known to mediate infective processes [53–56]. Structural studies on pIII suggest a domain structure comprised of exposed amino-terminal N1 and N2 domains intervened by a flexible glycine-rich linker [55], followed by another linker region and the carboxy-terminal (CT) domain. The CT domain is thought to anchor the minor coat protein to the viral capsid and to mediate post-infection release of the phage genome [53]. The critical first interaction of infection is mediated by binding of the N2 domain on pIII to the F-pilus on a host cell [54]. Retraction of the pilus enables subsequent binding between the N1 domain and the bacterial periplasmic TolA receptor [56] followed by injection of the viral genetic cargo and insertion of viral coat proteins into the bacterial inner membrane. Notably, it has been determined that even small amounts of pIII are capable of disrupting inner membrane structure and the ability of the F-pili to mediate further viral entry [57]. This observation has important practical consequences for in vitro selections during infection and amplification of interacting affinity clones, because careful controls are necessary to ensure the absence of pIII to prevent resistance to infection. Inside the cell, ssDNA is converted to double-stranded DNA (dsDNA) by host DNA polymerases and viral replication begins (Fig. 1). Upon production, all viral coat proteins localize to the bacterial membrane as integral membrane-spanning proteins prior to viral assembly [58]. Approximately 60 min post-infection, viral pV protein binding to replicating DNA blocks formation of the complementary strand, thereby switching synthesis toward the generation of progeny ssDNA whereupon new particles are packaged at the membrane prior to virion progeny release [59]. In general, filamentous phage replication is initiated within 10 min following infection and new virus produced at an estimated rate of approximately 200–300 phage particles per cell doubling time (up to 1000 per hour) [60], although others have estimated rates as low as 100 [61] or as high as 2000 per cell doubling time [59]. The rapid replication and high attainable virus titres have contributed to the appeal and widespread use of the M13 bacteriophage protein display system in biotechnology. In 1985, George Smith provided a pivotal first demonstration that antigenic polypeptides could be successfully displayed in a

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functional form as fusions to the pIII viral coat protein on the surface of phage particles [10]. Shortly thereafter, several groups showed that antibody fragments could also be displayed as fusions with viral coat proteins [12,62,63] and led the way for the development of phage-displayed antibody libraries. Since then, all five phage coat proteins have been used for display [32,64–66] but pIII and pVIII are most commonly used and best developed for display of proteins or peptides, respectively [67]. Given the limitations in expressing larger proteins as pVIII fusion [68] and its general lack of utility in expressing antibodies and antibody fragments [67], we will restrict discussion to antibody fusions with pIII. 3.2. Phage display of antibodies Early attempts to display polypeptides on the exposed surfaces of pIII incorporated genes of exogenous proteins directly into the viral genome [9,10,69]. However, these efforts revealed the limitations of certain approaches to expressing proteins fragments as fusions to phage coat proteins and led to the critical recognition that the insertion of a protein into the structural components of a viral particle can affect the balance of the phage–host system or the structural stability of the phage particle. This prompted the development of approaches that were well tolerated and enabled routine, high-efficiency display of proteins fused to pIII with

minimal perturbation of infectivity and propagation [69–71]. In particular, it was recognized that display of proteins could hamper phage infectivity, production and stability, and this led to the development of phagemid systems in which the heterologous protein is fused to an additional copy of pIII and phage production is accomplished by the use of helper phage containing wild-type pIII (wt-pIII) (discussed further under Section 3.4). In 1990, McCafferty et al. demonstrated the viability of expressing antibody variable domains as scFv fused to pIII on virus that exhibited specificity identical to the parent antibody in enzyme-linked immunosorbent assay (ELISA) and could be isolated from mixtures in which the specific clone represented less than one in 106 phage [12]. Since this initial demonstration, antibody variable domains have been displayed in a variety of formats including Fab [11,21,63,72], F(ab)2 [26,73], and scFv [74–76] (containing linked VH and VL domains), autonomous VH domains [20,22,77] (see Fig. 2) and even full-length IgG’s [78,79]. Insofar as Fabs are stable, functionally folded, well displayed, compatible with established immunological methods and now reliably converted to bivalent IgG molecules, we will focus our subsequent discussion primarily on approaches used for obtaining robust display and high diversity libraries based upon Fab fragments. It is worth noting however, that only through progress made on various fronts, has the true potential of this technology been

Fig. 1. The lifecycle of filamentous phage. Phage particles initiate contact with bacteria bearing the F-pilus. Additional interactions between pIII domains and bacterial receptors occur with pilus retraction, followed by injection of the phage genome into the cell interior. ssDNA is converted to the dsDNA replicative form, which enables the synthesis of viral proteins and the generation of new ssDNA required for replication. Nascent viral coat proteins take up residence in the bacterial inner membrane in preparation for viral assembly, and pV pre-packages the ssDNA viral genome for assembly and extrusion. Viral assembly and export occurs through a membrane pore without cell lysis.

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Fig. 2. The full-length IgG molecule and antibody fragments. The crystal structure (PDB entry 1HZH) shows that the IgG molecule is comprised of two heavy chains and two light chains, which associate to form a heterodimer possessing two identical antigen-binding sites at the distal end of each Fab. Antigen interactions are mediated collectively by hypervariable loops in both light and heavy chains, termed complementarity-determining regions (shown in red). Monovalent Fabs can fold independently of the whole IgG structure via the association of the N-terminal (VH–CH1) domains of the heavy chain and the entire light chain (VL–CL). Efforts to reduce the antigen-binding unit have resulted in the generation of single-chain variable fragments (scFv), comprised of the variable domains of the heavy and light chains linked through a polypeptide linkers. Further minimization has resulted in the expression of autonomous heavy-chain variable (VH) domains.

realized. Major advances include framework optimization [43], control of display valency [80–82], chain shuffling [83], cloning of antibody variable domains [84], production of phagemid systems [85], and improved library designs. As a maturing technology, synthetic antibody systems that display precisely designed antibodies can now be used to display repertoires with diversities in excess of 1010 unique clones [27], and these repertoires routinely yield antibodies of high affinity and specificity to a broad range of antigens. Although selection strategies have proven highly successful for obtaining desired antibody selectivity, there are a host of elements that exert a major influence over the selection process itself and these are discussed below. 3.3. Display of antibody–pIII fusion protein The critical roles that each domain of pIII plays in early infection mandate careful insertion of exogenous antibody genes to minimize disruption of viral replication and stability. The first attempts to fuse proteins to pIII inserted foreign genes into the inter-domain regions and although they were successful in this aim, defects in viral infectivity and propagation were noted [10]. In general, N-terminal fusions of either full-length or N-terminally truncated pIII variants are now employed for display of antibody fragments [9,12,70,86,87]. Importantly, when pIII domains critical for infectivity and viral release are omitted or compromised via insertion of exogenous genes, wt-pIII must be supplemented in order to maintain viable phage propagation required for clone selection from libraries (discussed further under Section 3.4). Both heavy [88,89] and light chains [90,91] have been successfully fused to pIII, although it is not clear whether either provides substantial advantage in the context of high diversity libraries given the apparent absence of systematic studies comparing approaches. In order to direct heavy and light chains to the site of viral assembly, leader sequences are added to each, resulting in their direction to the bacterial periplasm where they can associate and be incorporated into

a nascent viral particle. Functional antigen-binding sites are formed by the non-covalent association of heavy and light chains, and thus, display vector designs need to consider the stoichiometry of chain association. Vectors for Fab expression can be either bicistronic or monocistronic and enable expression under the same promoter from a single mRNA or different promoters from separate mRNAs, respectively. Systematic comparison of the two systems suggest that, although there is no apparent effect on phage titre, bicistronic vectors may improve folding of functional Fabs in comparison with monocistronic vectors [92]. Others have employed separate vectors to deliver heavy and light chains via co-transfection into the same bacterial cell [93]. Similarly, in vivo recombination events have been used to generate antibody fragments from two separate vectors encoding VH or VL domains, respectively, in either Cre recombinase-mediated [81,83,94] reaction or other site-specific recombinase systems [95]. Both approaches seek to enhance diversity of phage-displayed antibody libraries by exploiting random association of VH and VL domains. 3.4. Phage and phagemid display systems As with any technology based upon the presentation of functional proteins for investigation, the efficiency of display is paramount to the success of the application. In the case of phagedisplayed antibodies, display efficiency also has consequences regarding modes of interaction and the attributes of isolated binding clones and should be considered when using antibody libraries for selection of species that interact with an antigen of interest. Early phage display vectors were constructed by fusing the gene encoding the heterologous protein to be displayed directly to the pIII gene within the viral genome [12,63]. By substituting pIII fusion for wt-pIII, multivalent display is ensured since the only source of pIII is from the fusion product [10,69,96] (Fig. 3A). This has the advantage of enhancing functional affinity through avidity effects and enabling the selection of lower affinity antibodies that

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may be of interest [97]. However, it has been noted that the transformation efficiency of large phage vectors is reduced and multivalent display may limit the isolation of higher affinity clones, promote the generation of ‘polyphage’ due to aggregation [10,69,96], and result in complex effects when selections are performed on cellular systems. Variations on the early phage vectors thus sought to circumvent these issues and reduce the valency of antibody display and did so by incorporating a second, wt copy of the phage coat protein being used for fusion (referred to as hybrid phage vectors) [10,98]. This has the effect of producing both fusion and wt coat protein, which will compete for incorporation into the virion particle resulting in oligovalent display (ranging from zero to 5 copies per phage), although most proteins are displayed monovalently [9,71,99] (Fig. 3B). This approach effectively reduces the number of fusion proteins displayed such that aggregation is avoided and, without avidity effects the isolation of high-affinity antibodies is promoted. However, similar limitations in transformation efficiency are encountered and once produced, many virions may indeed possess no displayed antibody fusion (perhaps due to proteolysis of the fusion protein in the periplasm), which can reduce the effective size of the library and limit the potential for successful selection due to the presence of background phage particles. With the aim of improving transformation efficiency while controlling the number of copies of proteins displayed on the phage

A

B

surface, investigators have devised systems in which both fusion pIII and wt-pIII are supplied in trans to reduce vector sizes and enable heterologous display on phage particles [9,70,100,101] (Fig. 3C). It is estimated that, by supplying viral components in trans, transfection efficiencies can be increased by two to three orders of magnitude, and this has been accomplished by the design of phagemid systems [63]. Phagemids are designed to possess a protein-encoding gene of interest on a double stranded plasmid that however lacks most of the critical elements for the formation of functional phage. By virtue of their smaller size, increased transformation efficiencies can be achieved. They also possess an antibiotic selectable marker, an origin of replication that allows for plasmid propagation in Escherichia coli, and an f1 origin that enables the production of ssDNA from the replicative double strand and, when co-infected with helper phage, packaging of ssDNA into phage particles (Fig. 3C and D). Helper phage generally possesses a resistance marker distinct from the phagemid and contributes all components required for viral assembly and replication, but has been engineered with weakened packaging signals to enable preferential packaging of phagemid ssDNA [82,102,103]. When super-infected into cells already transformed with phagemid, helper phage enables the production of viral particles containing phagemid DNA and displaying the phagemid-encoded fusion protein [63], thus establishing the genotype–phenotype linkage that enables selection. Cells that have been super-infected can be selected

C

D

Fig. 3. Phage and phagemid vectors designed for display of antibody fragments. (A) Phage vectors with fusion-pIII as the sole source of pIII generate phage with multivalent display of antibody fragments. (B) Phage vectors encoding for both fusion-pIII and wt-pIII generate phage with a range of valencies. (C) Phagemid systems expressing fusionpIII supplemented with helper phage contributing wt-pIII generate phage particles with a range of valencies. (D) pIII-deleted helper phage rely upon fusion-pIII from phagemid as the sole source of pIII and express multivalent display protein similar to phage vectors that do not express wt-pIII.

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for by cultivation in media containing antibiotics for resistance markers on both the phagemid and helper phage vectors or isolated and identified by cellular infection and plating on the appropriate selection media. Many variations on the phagemid system have been developed in attempts to maximize functional antibody display [80– 82,85,101]. Studies have shown that wt-pIII supplied from helper phage is preferentially incorporated into nascent virus particles [12,101,104,105] resulting in the production of a substantial proportion of virus particles that possess no fusion protein whatsoever and a severe curtailing of the quality of specific clones obtainable from the library. One approach aimed at circumventing this problem was to eliminate wt-pIII expression from the helper phage, thus ensuring that during phagemid amplification, the pIII for viral assembly comes primarily from fusion pIII encoded in the phagemid [80–82,85] (Fig. 3D). However, in light of the critical role that pIII plays in infectivity and propagation of phage, the various approaches ranging from complete deletion [81,82,106] to partial truncations of helper pIII [85,106], continue to require supplementation with exogenous wt-pIII. To avoid its inhibitory effects on cellular infectivity, various sources of pIII have been employed, each of which differentially affect helper phage titre and purity. In some cases, helper phage are amplified in bacteria supplemented with exogenous pIII via transformation with pIII expression plasmids [81,85,106]. However, significant contamination of helper phage with pIII-encoding plasmid has been observed, (despite compromised phage assembly signals) [82,85], raising concerns about adverse effects not only on purity but also on infectivity and amplification, which were reflected in low titres. Alternatively, investigators have explored the expression of wt-pIII from genes either expressed from a complementing plasmid that is transcriptionally activated exclusively by phage infection [82] or integrated directly into the bacterial genome of host cells [85]. Other approaches designed to improve display levels include conditional pIII deletions that produce wt-pIII in non-infected cells, but not in phagemid-infected cells [80,107], and ‘pIII-sufficient’ helper phage that exploit the leakiness of amber stops in some host suppressor strains to provide limited expression of wt-pIII and counter preferential incorporation of wt-pIII [101]. Others have sought to eliminate the need for helper phage altogether by transforming bacteria with helper plasmids that lack packaging signals but supply all the packing components that support the formation of functional phage upon infection with phagemid [102]. The major advantage of this approach is that it eliminates the necessity of supplying helper phage at specific stages in the phage growth cycle such that phagemid can be propagated by simple infection into cells transformed with helper plasmid. In light of the complex interplay between phage, host, and phagemids, further optimization of display is likely forthcoming from molecular engineering of phagemids, phage vectors and bacterial cells used for library construction and amplification. 3.5. Natural immune repertoires and their display In humans, germline immunoglobulin gene segments (56 variable (V), 23 diversity (D) and 6 joining (J)) in pro- and pre-B cells undergo re-arrangement in the bone marrow in a tightly regulated process to generate diversity of the natural immune repertoire. The usage of VJ and VDJ gene segments in functional light and heavy chains of antibodies is not however uniform and some segments appear far more often than others. This has been attributed in part to recombinational frequencies that arise from variation in recombination signal sequences and their accessibility [108] and has led to the observation that some gene segments are also over-represented in particular disease states [109,110]. Further diversity in the natural somatic immune repertoire is generated from a variety

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of sources including junctional flexibility at coding joints and the addition of P- and N-nucleotides during B cell maturation. Subsequent exposure of B cells to antigen results in further diversification that takes place during activation in secondary tissues by the processes of somatic hypermutation and gene conversion which adds additional diversity to previously rearranged immunoglobulin genes. In 1991, Clackson and colleagues first demonstrated that antibody variable domain genes from mice could be amplified and cloned as a pIII fusion product into a phage display vector such that nearly the entire diversity of the mouse immune repertoire was represented, and from these repertoires, antibody fragments with specific binding properties could be obtained [62]. Since this demonstration, phage-displayed libraries of natural human immune repertoires have been used to isolate antibodies that bind specifically to a wide range of target proteins [76,81,111,112]. Although B cell immunoglobulin genes are most easily obtained via extraction from peripheral blood lymphocytes sampled from circulation, many tissues can be used to extract diversity including bone marrow [76], splenocytes [62], tonsils, lymph nodes, foetal liver [62], [113] and may yield libraries of varying diversity and clones of varying quality. Insofar as B cells in secondary lymphoid tissues may represent an activated population of cells that have encountered antigen, diversity extracted from these tissues may represent a bias in the repertoire arising from the in vivo process of affinity maturation, whereas diversity extracted from bone marrow or circulating B cells, would more closely reflect germline diversity following immunoglobulin gene rearrangement. Libraries based upon natural immune repertoires are useful insofar as pre-determined knowledge of antibody variable domain sequences are not required in order to generate functional antibodies and standardized methodology for generation of libraries is now widely available [114]. Although natural repertoires offer high functional diversity and have been successfully exploited for the construction of and selection from phage-displayed antibody libraries [111,112], there are key limitations that restrict the range of targets to which binding clones can be successfully isolated. One of the primary limitations that have confounded investigators and antibody engineers stems from functioning immune mechanisms that tolerize or delete clones to self-antigens, so as to prevent the development of autoimmune disease. This, of course, may impede the isolation of antibodies specific to these targets from libraries derived from natural immune repertories. For this and other reasons, synthetic or semi-synthetic sources of antibody diversity are increasingly being employed, thus avoiding the inherent limitations of natural immune repertoires. 3.6. Synthetic antibody repertoires The ex vivo production of antibodies has been envisioned since the now distant discovery of their material substance [115]. However, only within the last 20 years has this aim been fully realized by the advent of synthetic diversity libraries that provide a ‘chemical solution’ [11,116–119] to the biological constraints on natural repertoire diversity. Synthetic approaches are particularly attractive to those seeking to understand the fundamentals of antibody–antigen interactions or to engineer desired antibody properties, because they enable precise control over the composition of diversity incorporated into antigen-binding sites. Undoubtedly, this capability will offer the key to optimization of future libraries as we unlock a greater understanding of how sequences within antibody variable domains contribute to antibody stability, binding and function, and we develop our ability to engineer affinity agents with desired specificity. Indeed, the fundamental difference between natural and synthetic libraries can be highlighted by the source of their diversity whether obtained from biological

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(natural) or chemical (synthetic) sources. Whereas natural libraries extract and clone immunoglobulin variable domain sequences from immune tissues and do not generally introduce de novo diversity, synthetic libraries introduce wholly novel combinatorial oligonucleotide sequences into an antibody framework. Herein lies one of the fundamental strengths of synthetic libraries, in that they bear the potential to reflect diversity alien to natural immune repertoires, without inherent biases, and thus enable recognition of targets outside the natural scope. Some of the earliest successful attempts at showing the feasibility of obtaining specific binding clones from libraries generated from non-natural sources employed semi-synthetic approaches to generating library diversity [11,76,120,121]. These studies employed natural human repertoires, but incorporated synthetic elements in various ways including PCR-based rearrangement of germline VH domains [121], shuffling of naïve VH and VL domains obtained from peripheral blood lymphocytes [76], or introduction of CDR randomization using chemically synthesized random oligonucleotides [11,120]. Advances made over the subsequent 20 years have embraced primarily either cassette-based methods (which rely upon restriction and ligation of inserts) [25,114] or oligonucleotide-directed mutagenesis and whole plasmid amplification to incorporate diversity into synthetic antibody libraries [26,122,123], and we will focus our attention on the latter approach. 3.7. Incorporating synthetic diversity into antibody CDRs The chemical synthesis of nucleotide sequences for incorporation into antibody libraries has become a cornerstone of phage display techniques. Commercial availability of high-quality synthetic DNA and facile incorporation into antibody frameworks have helped contribute to their widespread use in the construction of phage-displayed libraries from which novel binding specificities are now routinely isolated [25–27,47]. ‘Hard-randomized’ oligonucleotides are synthesized using equimolar mixtures of the four phosphoramidite precursors resulting in the generation of degenerate codons encoding for all 20 amino acids. Theoretically, the diversity of a library can be determined by the number of options at a particular position (i.e. how many amino acids are encoded for) to the power of the number of positions to be randomized. In a library in which 8 positions of a CDR have been randomized with NNN codons, there are 2.6  1010 (208) theoretical unique clones. Given a practical diversity ceiling of approximately 1010 clones (limited by transformation efficiency of E. coli in which libraries are generated), it becomes apparent that only 6–7 fully diversified residues can be completely represented in a library before the theoretical diversity eclipses the practical achievable diversity [119]. Similarly, the theoretical diversity of a library often exceeds its functional diversity (i.e. the total repertoire of clones that are well-folded and capable of binding) due to the presence of nonfunctional clones that do not fold properly and sequence bias that arises from the unequal representation of amino acids by codons. Collectively, these can contribute to a significant loss of diversity and the generation of misfolded antibody fragments that can give rise to non-specific binders. Thus, synthetic techniques have aimed at generating diversity while minimizing the incorporation of codons that adversely effect display and library quality. In short, the practice of restricting random diversity to exclude non-functional sequence (thus maximizing functional diversity) is of benefit and has been a guiding principle in the design of synthetic libraries [47]. A simple and superior approach to reducing non-functional diversity, while maintaining complete randomization, is achieved by restricting nucleotide incorporation in the third codon position to either G/C (NNK) or G/T (NNS) such that only 32 codons are

generated, including only one of the three stop codons but encoding for all 20 amino acids [11,117]. This limits the potential for incomplete polypeptides that can hamper the quality of an antibody library and reduces (but does not eliminate) codon bias [11,26,27,124,125]. This approach bears the additional advantage of a singly represented amber stop codon to further eliminate truncated mutants by expression in suppressor E. coli strains. Alternatively, the development of trinucleotide phosphoramidite cassette technology has enabled the elimination of codon biases that can negatively alter the random distribution of amino acids in synthetic antibody libraries [126,127]. Insofar as each three-nucleotide cassette is chosen and designed to encode for only a single amino acid, mixtures of trinucleotides further allow for either diversity tailored to a subset of amino acids incorporated at desired proportions or for a truly equal distribution of all amino acids. In concert, this represents a substantial increase in positional control during synthesis of mutagenic oligonucleotides. Oligonucleotides generated in this manner are increasingly being used in the construction of synthetic libraries for selection of antibodies to diverse targets [25,128–130]. Both tailored and trinucleotide-based mutagenic oligonucleotide methods of generating libraries have been used to capitalize on the analysis of structural and sequence data derived from natural antibody repertoires [17,131] and to tailor diversity to regions that are more likely to play roles in antigen binding [27,132,133]. Computational analysis of available antibody data has revealed the presence of CDR positions with low positional diversity [27,132,133] or low solvent exposure [27], and this knowledge has been used to restrict diversity at these sites using the methods discussed above, with the aim of enhancing the representation of diversity at sites more critical to target interaction. Control of the positional composition of CDRs raises the question of what constitutes a highly functional library. In the absence of other reduced diversity models, mimicry of the natural immune repertoire is an intuitive and effective approach to generating synthetic diversity [26,27]. But does this simply recapitulate the very same limitations by which natural antibodies are constrained? With only a paucity of CDR binding sequences obtained from synthetic repertoires, it is unknown whether the positional frequency would parallel that of natural antibodies. With an increasing number of groups now using synthetic antibody libraries and a growing portfolio of high affinity binding clones obtained from synthetic libraries, the opportunity will arise to compare the observed positional frequencies between natural and synthetic libraries. It will be interesting to determine whether the antibodies selected from synthetic repertoires ultimately come to reflect positional frequencies of natural repertoires or whether they reflect the unique sequences of the synthetic space. With this question in mind, the limits of the synthetic sequence space have been explored using libraries based upon drastically reduced diversity [21,117,134,135] and by characterizing the spectrum of diversity tolerated by libraries generated in an autonomous VH domain [21]. The construction of phage-displayed antibody libraries using minimalist diversity repertoires revealed that high specificity could be obtained with as few as only two amino acids in the CDRs [136], and that certain residues are more effective than others in contributing to binding specificity and affinity [135]. Alternately, studies suggest that the range of diversity tolerated by antibody frameworks may exceed that which is observed in nature. Synthetic diversity introduced into autonomous VH domains and followed by selection for functional folding, enabled the construction of a database of diversity upon which positional analysis could be conducted. Comparison of diversity revealed that significantly higher diversity could be tolerated in synthetic VH domains than their natural counterparts [23]. These results suggest that upper limits of natural diversity do not arise from structural constraints inherent

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to the antibody framework, but more likely from other mechanisms and that in vitro evolution of synthetic antibodies may offer a broader scope of potential diversities. These findings advance our understanding of antigen–antibody interactions and the potential limitations of the natural immune repertoire. Moreover, they provide practical clues to viable engineering techniques for library construction and obtaining novel binding specificities. 3.8. Library construction To construct synthetic antibody libraries, diversity is incorporated from synthetic oligonucleotides into the CDRs of antibodies in vectors designed for phage display. Mutagenic oligonucleotides are generally incorporated into antibody variable domain frameworks using either standard cassette-based methodology with restriction-based cloning [132,137] or via site-specific annealing of degenerate oligonucleotides to ssDNA and whole plasmid synthesis [26,27]. Synthetic diversity in cassette form can be obtained by (1) PCR amplification of individual rearranged CDR segments from isolated B cells and random reconstruction of variable domains by overlapping PCR extension of diverse CDR pools with framework segments to generate full-length variable domains [137], (2) chemical synthesis of overlapping degenerate nucleotides and PCR-based amplification of variable domain cassettes [25], or (3) a combination of amplified natural immune repertories combined with synthetic oligonucleotides by overlapping PCR resulting in semi-synthetic diversity [138,139]. Cassette-based methods employing degenerate oligonucleotides rely upon PCR amplification in order to assemble multiple sites of degeneracy that cannot be synthesized in a single oligonucleotide due to length limitations for synthetic DNA synthesis. An alternate approach, requiring no prior assembly of overlapping fragments, entails the use of discrete degenerate oligonucleotides flanked by sequence complementary to framework regions surrounding CDRs. Synthetic oligonucleotides are annealed directly to ssDNA vectors and incorporated into replicative dsDNA plasmids using optimized site-directed mutagenesis techniques [123,140]. This methodology exploits the f1 origin in phage genomes that signals for and enables conversion of dsDNA to ssDNA. ssDNA is isolated from purified phage containing template DNA, and once annealed to the ssDNA template, oligonucleotides initiate polymerase-dependent nucleotide extension according to the template sequence of the phage vector, which enables whole vector synthesis, ligation and the formation of covalently closed, circular dsDNA. One of the major advantages of this approach is the ability to simultaneously mutate multiple CDRs in a single mutagenesis reaction without the need for restriction sites. Further, the incorporation of length diversity into CDRs is facilitated by the use of pools of degenerate oligonucleotides of varying lengths. A successful mutagenesis reaction will result in the incorporation of mutagenic oligonucleotides into heteroduplex dsDNA and the complete conversion of ssDNA to dsDNA. The resultant plasmid pool reflects combinatorial diversity design and is capable of transforming E. coli with a high mutation frequency (80%). Introduction of the plasmid pool into E. coli cells by electroporation offers the most efficient means of transformation but requires cells containing the F0 episome that enables subsequent helper phage infection required to amplify libraries. Infection of electrocompetent cells with helper phage provides the necessary proteins and viral machinery required for the synthesis of mature phage particles displaying antibody fragments encoded in phagemid vectors, and standardized methodology are available to accomplish this [123]. Libraries generated in this manner routinely contain >1010 unique members. As discussed under Section 3.4, transformation efficiency is a critical determinant of library size. Electroporation

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efficiencies vary amongst bacterial strains [103,141] and are dependent upon the phase of growth [142,143], cell density [144] and vary linearly as a function of DNA concentration over 6 orders of magnitude [143]. Efforts to improve upon bacterial strains used in propagation of phage libraries have resulted in increased transformation efficiencies [71] and the careful testing of any strain is warranted prior to use for construction, amplification or propagation of phage-displayed antibodies libraries. Following electroporation, cells are rescued and titration on selective media corresponding to phagemid resistance enables the estimation of library diversity assuming that each transformant represents a single unique clone. Growth of rescued cells results in library amplification and the generation of multiple copies of each phage displaying a unique clone depending on library size and final phage concentration. In general, overnight amplification can result in phage concentrations of 1011–12 phage/mL representing 10–100 instances of each clone per mL for a library of 1010 diversity. Phage preparations can be concentrated to a maximum of 1013 phage/mL using crowding agents such as concentrated solutions of polyethylene glycol (PEG) and NaCl, but at higher concentrations, changes in viscosity may alter phage diffusion. Following concentration, phage can be resuspended to a desired concentration in an appropriate buffer and used directly in selections against target proteins as described below. 3.9. Selection strategies Although library size is a major determinant of the availability of highly functional antibody clones, the selection process is the point at which the in vitro conditions that determine displayed target epitopes are established, the diversity of the library is sampled and binding events occur that determine the clones that will be recovered and amplified. Properly folded and displayed antigen is critical to obtaining clones that bind functional protein not only in vitro but also in a cellular context, and ambient conditions can influence both the conformation of the antigen and the affinity of selected binding clones. By manipulating the in vitro selection environment, we capitalize on the opportunity to tailor conditions that promote desired molecular target configurations and binding events, which in turn enhances the chances of obtaining clones with desired binding properties. Published studies confirm that carefully established in vitro conditions during selection enable the isolation of antibody clones that exhibit desirable properties including exquisite specificity, ultra-high affinity, conformational specificity, thermostability, and species cross-reactivity. In our opinion, the thoughtful and imaginative development of selection strategies and conditions leaves few boundaries to the specific properties that can be engineered. The following section will survey techniques for antigen immobilization, epitope targeting, affinity enhancement, and elution and expansion. 3.9.1. Protein immobilization Next to the quality of the phage-displayed antibody library, the most critical determinant of selection success is perhaps the immobilization of the target protein in a functional conformation. Most frequently, proteins are passively adsorbed from solutions onto polystyrene microplate or tube surfaces where they are held by non-specific interactions in random orientation. Due to the physicochemical nature of the surfaces and the forces that stabilize protein conformations, it is not uncommon for protein adsorption to result in some denaturation, which can reduce the proportion of correctly folded antigen [134,145]. In addition, the random nature of immobilization impedes the uniform presentation of epitopes and increases the likelihood that, at least in some target molecules,

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some epitopes may be sterically inaccessible. Nevertheless, passive immobilization is appealing due to the simplicity of the method and the frequent success of selections that yield clones that bind functional protein both in vitro and in cells. Specific capture methods have been widely explored and provide alternatives when adsorption methods are inadequate. Many recombinant protein libraries incorporate affinity tags to facilitate purification and these can be exploited for capture using functionalized immobilization surfaces [146,147]. Affinity tags provide options for tethering proteins in a relatively uniform orientation, with minimal disruption of native conformation. There are numerous approaches, including metal ion- [148,149], antibody- [150,151], and biotin-capture [69,152–154], as well as newer protein–ligand capture technologies [155,156]. Widely used, nickel-based metal affinity supports often require high-densities of functionalized surface to provide sufficient sites for metal–histidine co-ordination, but can increase non-specific interactions [157]. However, the ongoing development of novel nitrilotriacetic acid-based capture methods will likely circumvent these issues in the near future [134,158]. Alternately, the antibody-based capture of affinity tags (such as anti-hexahistidine, anti-glutathione-S-transferase (GST) or anti-maltose binding protein (MBP)) relies critically on the high affinity of the capture antibody. Insufficient affinity results in dissociative loss of immobilized target and ultimately hampers enrichment of desired phage populations during selection. Chemical biotinylation of proteins is a proven technique when used with care to optimize and avoid excess labelling that can alter surface properties, protein stability and solubility [134,158]. Site-specific enzymatic biotinylation with BirA ligase [153] provides an alternative for singly labelling target proteins, but it requires an additional cloning step to add the substrate sequence recognized by the ligase. To enable affinity-independent release, protein immobilization methods have incorporated linkers possessing cleavage sites that enable specific release of the target protein under mild conditions (discussed further under Section 3.9.2). Although it is unlikely that any one approach will work for all protein targets, there are nevertheless ample choices available to

A

ensure that sufficiently optimal conditions can be achieved for successful selection. 3.9.2. Clonal selection and elution Once suitably immobilized, non-specific sites on the proteincovered surface are blocked and ready for exposure to phage libraries. A critical consideration here is to ensure adequate coverage of the diversity of the library. Phage preparations can generally be concentrated to a maximum of 1013 phage/mL before viscosity adversely affects selection. Ideally, the phage count will exceed diversity of the library by 100–1000 fold to ensure numerous copies of each clone and to facilitate the selection of rare binding clones. For a library possessing a diversity of 1010, a 1-mL volume of phage provides 1000-fold coverage at maximum concentration. However, microplates in which selections are conducted generally have a capacity of a few hundred microlitres and thus in the first round of panning, multiple wells are coated and exposed to phage and in order to completely cover all diversity in a 1-mL volume, 10 wells would be required. To avoid the propagation of non-specific clones, it is customary to pre-expose phage libraries to an adsorbed irrelevant protein or purified version of the affinity tag used to immobilize target protein. Following pre-incubation with irrelevant protein, depleted phage solution is transferred to immobilized target protein to allow for specific interaction and binding. Elution techniques also vary widely depending upon the means by which antigen is immobilized, the surface upon which selections were conducted and the level of simplicity and/or specificity desired by the investigator. The most commonly employed means of elution aims to disrupt the molecular interactions between bound phage-displayed antibody and immobilized antigen by drastic alteration of pH. Elutions with either low or high pH are effective and most often employ either HCl or triethylamine [12], respectively. Despite their widespread use, these methods risk the possibility of incompletely eluting phage, thus hampering recovery and the effectiveness of selection. Where the target antigen is known, competitive elution with excess antigen is a gentle and highly specific means of releasing

B

Fig. 4. The phage display selection cycle. (A) Phage pools exposed to target protein in multiple rounds of selection are gradually enriched for target-specific binding clones. In each round, negative selections are conducted to remove phage clones that bind to undesired species and promote isolation of clones that interact specifically with target. Non-binding phage are washed away and remaining phage eluted and amplified. (B) The enrichment process can be monitored at each selection round by comparing the titres of phage eluted from target protein or non-specific protein. Increasing titres in panning against target versus non-specific protein indicate enrichment of specific binding clones.

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phage clones for propagation that can limit the propagation of nonspecific clones [62,196,197]. To obviate concerns about incomplete release of tight binding clones that may be lost during elution, affinity independent methods have been developed and, in general, entail cleavage of a unique proteolytic site [90,91,198], target DNA sequence [199], or disulfide bond [81]. Investigators have demonstrated that phage elution can be dispensed with completely by direct infection of E. coli cells added to antigen-bound phage in selection wells. Antigen-bound phages maintain infectivity and specific Fab-phage are propagated following incubation [200– 202]. Although no apparent differences in output titres yielded by direct-infection versus chemical elution are noted, biases in clone affinity and/or antibody display may arise. Following library exposure to target, unbound Fab-phage clones are washed away and target-immobilized Fab-phage is eluted for amplification / isolation of binding clones (Fig. 4A). To monitor the selection process, it is advisable to titrate input and output titres, to determine enrichment over background for each round (Fig. 4B). Enrichment can be assessed by the inclusion of a well of irrelevant protein to which the phage library is exposed in parallel to target proteins for comparison of post-elution titres (Fig. 4B). Alternately, binding of phage eluted from both target and irrelevant protein can be determined by pooled phage enzyme-linked immunosorbent assays (ELISAs) to target protein. In pools that exhibit enrichment, antibody–phage clones can then be isolated by plating and picking individual colonies to identify those that exhibit specific binding. Isolated antibody fragments can then be expressed by conversion of antibody–phage clones to expression clones via insertion of a stop codon between the genes encoding antibody fragments and fused pIII coat protein, or where libraries are engineered to posses an intervening ‘amber stop’, simply transferred to a non-suppressor strain of bacteria, in both cases completely avoiding the use of animals at all stages. It is generally recognized that there exists correlation of biological activity with antibody affinity [159–162] and antibodies isolated from initial rounds of selection may require additional optimization of affinity, activity, thermostability or any other property affecting its utility. In vitro selections provide ideal opportunities for optimization by tailoring panning conditions to support the isolation of desired antibody attributes in a process referred to as ‘directed evolution’. 3.9.3. In vitro directed evolution Combinatorial gene synthesis and recombinant expression techniques have enabled the creation of diverse libraries of functionally distinct species including antibodies, enzymes, ligands etc. Following initial selection and isolation of binding clones, the directed evolution of these molecular species can be routinely accomplished via repeated rounds of selections from libraries in which selective pressure is applied (including competitive ligand, decreasing target concentrations, increased washing, negative selections, increased temperature) that enable the optimization of lead molecular species, ultimately yielding desirable properties perhaps not found in nature. Where the sequence space is known (i.e. where a binding clone has been identified), or where an investigator has insight into the nature of mutations that may be tolerated at a particular position, mutagenic oligonucleotides with restricted diversity are highly useful for isolating optimal antibody variants and exploring sequence-structure relationships [133,136]. The coupling of genotype and phenotype and the ability to selectively randomize discrete sites within antibody-binding interfaces have made possible the engineering of desirable properties by evolving the sequence and structure of the antibody-binding site. Tailored diversity is particularly valuable for the in vitro evolution of binding clones isolated from phage-displayed antibody libraries to scan for sequences that optimize binding properties while

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avoiding deleterious mutations. When restricted/tailored diversity is coupled with the application of evolutionary pressures during selection, it becomes possible to isolate variants with desired attributes (e.g. high affinity, particular conformation, thermostability etc.) and general strategies now exist for the directed enhancement of affinity, specificity and thermostability. Several investigators describe and provide evidence for the effectiveness of techniques aimed at increasing affinity by enhanced on-rates [163], and/or slowed off-rates simply by adjusting binding and washing times and/or temperatures. Immobilization of progressively decreasing target protein concentrations during consecutive rounds of selections can further promote the isolation of higher affinity clones [164,165]. Additional selection pressure can also be applied to high affinity antibody clones by panning in the presence of soluble target protein in tandem with extended wash steps [74,165–167]. This tactic effectively stratifies clones on the basis of off-rate by capturing and removing clones having higher off-rates with soluble target and retaining clones with low off-rates bound to immobilized target for elution and isolation. This technique has been proven viable for enhancing affinity to the sub-nanomolar or even picomolar range [74]. The versatility of in vitro selections has spurred the development of techniques for isolating antibodies exhibiting conformational specificity [32,168], enhanced thermostability [169], species cross-reactivity [170–172], bispecificity [173,174] or specificity for non-immunodominant epitopes [175]. When coupled with soft-randomized mutagenic oligonucleotides (described below) based upon CDR sequences for existing specificity clones, these approaches become powerful techniques for the directed evolution of antibodies that recognize complex epitopes and make possible the generation of antibodies to targets that would not otherwise be accessible with conventional methods. Soft-randomization of a sequence is accomplished by ‘spiking’ nucleotides during each coupling cycle of a chemical synthesis with defined quantities of the other three ‘aberrant’ nucleotides. This results in the random mis-incorporation of each, such that a position will possess a defined proportion of wt sequence, the remainder of which varies over the entire set of 20 amino acids [133,176]. Phage-displayed libraries of antibodies constructed with soft-randomized oligonucleotides are biased toward wt sequence but enable a relatively broad exploration of the sequence space [71]. Alternately, a more restricted scan can be achieved via synthesis of degenerate nucleotides that encode for only a subset of amino acid residues that are physicochemically similar or ‘homologous’ to the wt [117,177,178]. Homology scanning provides a less drastic alternative to hard or soft randomization techniques that can avoid the incorporation of residues known to destabilize binding loops or can help to isolate conserved antibody variants with optimal complementarity. Alanine scanning can be an invaluable tool when coupled with phage display for mapping and quantifying energetic contacts between residues in the antibody combining site and target antigen [177,178]. This information then provides a rational basis for subsequent randomization schemes that precisely direct randomization efforts toward critical residues. 3.9.4. Alternate selection strategies In addition to selections on purified protein, in vitro panning of phage-displayed antibody libraries on complex surfaces and samples has also been successfully conducted on normal [179,180] and tumourigenic cells [179–182], engineered bacterial [183], yeast [168,181] and mammalian cells [184–186], proteoliposomes [187] and tumour histological samples [180,183,185,188,189]. Recent studies have even demonstrated the feasibility of obtaining clones to a rare, single cell in a heterogeneous population via microselection [190,191]. Engineered cell lines are attractive as a platform for presenting protein to antibody libraries insofar as they

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offer a means of displaying proteins such as insoluble membrane proteins [192,193] or proteins that require a native environment and would otherwise be impossible to target by means of simple adsorption. Alternately, selection strategies can be devised to identify cell-specific surface markers with no a priori knowledge of the particular target [128,194] or to isolate antibodies specific to active conformers [168,195]. These strategies often employ subtractive techniques aimed at depleting libraries of clones specific for surface antigens shared between cell lines and then panning against the cell line of interest to isolate clones binding to distinctive biomarkers. 3.9.5. High throughput methods for antibody selection Adoption of a high throughout approach to phage selection and engineering possesses the potential to transform this effort to an industrial scale operation where simultaneous selection against hundreds or even thousands of targets is possible. The in vitro selection process and accessibility of genome sequences make this technology wholly amenable to integration with liquid handling systems, and high-throughput pipelines for cloning, sequencing, expression and characterization of specific antibody clones. Indeed, one of the major advantages of generating synthetic libraries in a single framework is versatility, insofar as it enables sub-cloning of binders into alternate expression platforms for rapid characterization and comparison. Multi-well plate selection is an obvious point of departure [203], but alternate selection platforms including 96 solid-support, pin arrays [204], magnetic particles [205] and electrophoretically immobilized antigen [206] have also been explored and are finding impressive success [206,207]. Indeed the establishment of high-throughput surface plasmon resonance methodology for obtaining kinetic constants from 96 Fab fragments in a single experiment [208] provides the opportunity to automate every single step of the selection process from antigen challenge through clone isolation and sequencing, expression conversion and affinity characterization. 4. Conclusions Phage-displayed combinatorial libraries of antibody fragments have demonstrated wide-ranging utility in the selection and engineering of high affinity binding clones with properties desirable for a variety of purposes [1,209]. Phage-displayed antibodies containing synthetic diversity have been pivotal in acquiring a deeper understanding of antibody structure and function, have yielded antibodies of exquisite specificity and high affinity, and are now poised to make valuable contributions to basic research and drug development. We believe that continued advances in our understanding of diversity within antibody CDRs, coupled with the development of novel approaches for creating synthetic diversity in antibody frameworks, will continue to drive the construction of high-quality libraries that routinely yield specific, tight-binding clones to diverse antigens beyond the scope of conventional hybridoma methodology. By embracing an imaginative approach to devising novel in vitro selection techniques, it is possible to expand the versatility of in vitro selections to obtain rare clones with valuable bioactivity to virtually any molecular species. In light of the versatility, robustness and proven capabilities of the technology, we anticipate that phage-displayed antibody libraries will be a cornerstone of modern biotechnology and drug development for the foreseeable future. References [1] A.L. Nelson, E. Dhimolea, J.M. Reichert, Nat. Rev. Drug Discov. 9 (2010) 767– 774.

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