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. Author manuscript; available in PMC: 2023 Feb 13.
Published in final edited form as: Methods Mol Biol. 2022;2446:269–298. doi: 10.1007/978-1-0716-2075-5_13

Engineering pH-Sensitive Single Domain Antibodies

Tosha M Laughlin 1, James R Horn 1,*
PMCID: PMC9924069  NIHMSID: NIHMS1860082  PMID: 35157278

ii. Summary/Abstract

There is increasing interest in expanding an antibody’s properties beyond high affinity and specificity. One such feature is custom regulation of the binding event, such as pH-dependent control. Here, we provide the methodology for generating single domain antibodies (sdAbs) that bind their antigen in a pH-dependent fashion. As each sdAb is unique, we start by providing the conceptual framework for designing a combinatorial histidine scanning library within an sdAb antigen binding interface. Methods are provided to create a phage display library, containing up to 1×1010 unique members where each permutation of histidine substitution is sampled within the confines of the specified interface region(s). Finally, we describe phage display protocols for the selection and analysis of unique pH-dependent sdAb clones.

Keywords: VHH, phage display, combinatorial histidine library, nanobody, pH switch, linked-protonation, linked-equilibria

1. Introduction

Antibodies are used in a wide range of applications, such as therapeutics, diagnostics, and chromatography. Perhaps not surprisingly, these applications have expanded beyond using conventional IgG antibodies (150 kDa) to include smaller antibody fragments, such as single domain antibodies (sdAbs). The variable domain from camelid heavy-chain-only antibodies (termed VHH) (1) is perhaps the best known example of a sdAb. Due to their small size and modular nature, sdAbs are typically more soluble/easier to express (2), are able to access distinct antigen regions (3), and are easily manipulated into novel constructs (4, 5) as compared to conventional antibodies. Furthermore, the single domain architecture has facilitated their use in intracellular targeting applications (6), as well as potentially as oral therapeutics, capable of maintaining function within a gastrointestinal tract environment (7, 8). Consequently, sdAbs serve as attractive, modular building blocks across a range of therapeutic and diagnostic applications (9).

Antibody engineering is often primarily focused on enhanced binding affinity, which is desirable for most applications. However, protein interactions in biology often possess function beyond high affinity, including binding events regulated by environmental conditions. One example of environmental regulation is linked-protonation events that influence a protein’s binding at different pH (10). The ability to alter the strength of a protein interaction with changes in pH opens opportunities in both therapeutic and in vitro antibody applications. For example, there is evidence that generating pH-dependent therapeutic antibodies can lead to desirable antibody recycling, as opposed to antibody degradation in acidic lysosomes (11, 12). In addition, pH-sensitive antibodies, which do not require extreme elution conditions, are ideal for immunoaffinity chromatography (13, 14).

The origins of pH-dependent protein binding stem from protonation events that are linked to binding. This typically involves one or more ionizable side chains exhibiting a change in environmental conditions between the free and bound states. The simplest approach to create pH-dependent binding is to introduce an ionizable residue, such as histidine (His), into the binding interface using site-directed mutagenesis (15). The limitation with this method is that substituting a wild-type (Wt) residue with His may not always produce a linked event and may frequently result in unintended penalties to protein stability and/or binding at the permissive pH. Here, a combinatorial library approach is described, which is based on general phage display library preparations by Sidhu and coworkers (16, 17). This method, originally developed and tested using an anti-RNase A VHH sdAb model system (18), allows the user to generate a phage display library where all possible combination of His or Wt residue are sampled at multiple positions in the binding interface. Subsequent phage selection allows the user to identify clones that retain strong binding at the permissive pH, while displaying weakened binding at lower pH. This combinatorial approach increases the likelihood of identifying highly sensitive pH-dependent clones, due to sampling simultaneous His substitutions, without major penalties to binding at the permissive pH.

2. Materials

All solutions should be prepared using ultrapure water (18 MΩ·cm at 25°C). All culture media are autoclaved. Where noted, buffers/solutions should be autoclaved or filter-sterilized using 0.2 μm filters. Buffers and reagents are used at room temperature, unless otherwise noted. Aerosol barrier pipette tips should be used for all sample transfers containing phage particles to minimize the potential for contamination. Similarly, when possible, use plastic disposable labware for phage samples. Glassware or plasticware exposed to phage should be subjected to a multi-step decontamination process, starting with a 24–48-hour incubation in a detergent/bleach bucket wash (e.g., 1% SDS / 2% bleach), followed by a steam sterilization in an autoclave using a tray filled with a small amount of water, and finally subjecting the labware to a high temperature dishwasher run. Labware can then be subjected to a dry autoclave run to prepare for storage until future use.

2.1. Conceptual framework for combinatorial library and design of degenerate and stop codon oligonucleotides

  1. Coding sequence of sdAb inserted as a fusion with gene 3 within the “drop-out” phagemid vector designed by Kay and coworkers (19).

  2. Knowledge of likely antigen binding interface residues (e.g, amino acid sequence alignment to identify hypervariable regions or crystal structure of the sdAb/antigen complex).

2.2. Small scale preparation of uracil-containing single strand DNA wildtype template

  1. SOC: 20 g/L tryptone, 5g/L yeast Extract, 0.584 g/L NaCl, 2.5 mM KCl, 10 mM MgCl2, 5 mM MgSO4, 20 mM glucose.

  2. Phagemid DNA sample containing sdAb from Subheadings 2.1/3.1.

  3. 1 mm gap electroporation cuvette.

  4. Electroporator.

  5. 50 mg/mL carbenicillin (Carb) stock in water, filter sterilize (0.2 μm).

  6. 34 mg/mL chloramphenicol (Cam) stock in 95% ethanol.

  7. 25 mg/mL kanamycin (Kan) stock in water, filter sterilize (0.2 μm).

  8. M13KO7 helper phage (NEB, Ipswich, MA, USA).

  9. Escherichia coli CJ236, dut− /ung− electrocompetent cells (Lucigen Corporation, Middleton, WI, USA)

  10. 14 mL Falcon culture tubes.

  11. 50 mL conical tubes.

  12. 2×YT broth and agar plates: 16 g/L tryptone, 10 g/L yeast extract, 5 g/L NaCl. Add 15 g/L agar for plates.

  13. 2×YT-Carb/Cam broth and agar plates: Supplement media from item 12 with 50 μg/mL Carb and 5 μg/mL Cam.

  14. 2×YT-Carb/Kan broth: Supplement media from item 12 with 50 μg/mL Carb and 25 μg/mL Kan.

  15. Polyethylene glycol (PEG)/NaCl: 20% PEG 8000 kDa, 2.5 M NaCl.

  16. Phosphate-buffered saline (PBS)-A: 20 mM Na2PO4, 150 mM NaCl, pH 7.4. Autoclave.

  17. QIAprep Spin Miniprep Kit with QIAprep 2.0 spin columns, PE and EB buffer (Qiagen, Valencia, CA, USA) (see Note 1).

  18. MP: 5.7 mM citric acid shaken at 200 rpm for 5 minutes, filter sterilized (0.2 μm).

  19. MLB: 1 M sodium perchlorate in 30% v/v isopropanol.

  20. Shaker incubator.

  21. Spectrophotometer and cuvettes.

  22. NanoDrop UV-Vis spectrophotometer.

2.3. Stop codon insertion using small-scale Kunkel mutagenesis

  1. 10 mM ATP: freshly prepared, kept on ice.

  2. 100 mM dithiothreitol (DTT): freshly prepared, kept on ice.

  3. Custom oligonucleotide(s): see Subheading 3.1.

  4. 10× TM: 0.5 M Tris, pH 7.5, 0.l M MgCl2, filter sterilized (0.2 μm).

  5. T4 polynucleotide kinase (NEB, 10 U/μL).

  6. T4 DNA ligase (NEB, 400 U/μL).

  7. T7 DNA Polymerase (NEB, 50 U/μL).

  8. dNTP mixture: 25 mM each of dATP, dCTP, dGTP and dTTP.

  9. Wizard PCR Cleanup kit (Promega, Madison, WI, USA).

  10. Wizard Miniprep Kit (Promega).

  11. E. coli XL-1 Blue chemically competent cells (Agilent, Santa Clara, CA, USA).

  12. 14 mL Falcon culture tubes.

  13. SOC: see Subheading 2.2.

  14. LB-Carb broth and plates: 10 g/L tryptone, 5 g/L yeast extract, 5 g/L NaCl. Add 15 g/L agar for plates. Supplement with 50 μg/mL Carb.

  15. Heating block and water baths.

  16. Nuclease-free water.

2.4. Large scale preparation of uracil-containing single strand DNA stop codon template

  1. Phagemid DNA sample encoding sequence-verified stop codon sdAb template from Subheadings 2.3/3.3.

  2. All materials from Subheading 2.2.

2.5. Large scale in vitro synthesis of heteroduplex degenerate dsDNA

  1. Uracil-containing single strand DNA stop codon template from Subheadings 2.4/3.4.

  2. All materials from Subheading 2.3.

2.6. Phage display library generation

  1. BTX ECM-600 electroporation system.

  2. Heteroduplex degenerate dsDNA from Subheadings 2.5/3.5.

  3. 2 mm electroporation cuvettes.

  4. 250 mL baffled culture flasks.

  5. 2×YT-Carb broth and agar plates: see Subheading 2.2. Supplement with 50 μg/mL Carb.

  6. 50 mL conical tubes

  7. SOC: see Subheading 2.2.

  8. E. coli SS320 electrocompetent cells (Lucigen Corporation, Middleton, WI, USA).

  9. M13KO7 helper phage (≥ 1x1012 phage/mL).

  10. 250 mL and 4 L baffled flasks.

  11. Sterile 96-well plate.

  12. Shaker incubator.

  13. PEG/NaCl: see Subheading 2.2.

  14. 500 mL centrifuge bottles.

  15. NanoDrop UV-Vis spectrophotometer.

  16. PBS-A: see Subheading 2.2.

2.7. Phage titering

  1. E. coli XL-1 Blue chemically competent cells.

  2. 5 mg/mL tetracycline (Tet) stock in 70% ethanol.

  3. 2×YT-Tet broth: see Subheading 2.2. Supplement with 10 μg/mL Tet.

  4. 2×YT-Carb agar plates: see Subheading 2.2. Supplement with 50 μg/mL Carb.

  5. Sterile 96 well plate.

  6. PBS-A: see Subheading 2.2.

  7. 14 mL Falcon culture tubes.

  8. Spectrophotometer and cuvettes.

  9. Sterile multi-channel pipette basin.

2.8. Biotinylation of antigen

  1. 100 mM sodium phosphate, 150 mM NaCl, pH 7.2.

  2. EZ-Link Sulfo-NHS-SS-Biotin (Thermo Fisher Scientific, Waltham, MA, USA).

  3. Dialysis tubing or cassettes.

  4. PBS-A: see Subheading 2.2.

2.9. Phage display panning with double selection

  • 1.

    SAV Beads (Streptavidin MagneShpere Paramagnetic Particles, Promega).

  • 2.

    Microcentrifuge tubes.

  • 3.

    Combinatorial phage library: prepare in Subheadings 3.6 and 3.7. An ideal titer is 1012 phage per mL.

  • 4.

    Magnetic stand capable of holding microcentrifuge tubes.

  • 5.

    KingFisher magnetic bead handler.

  • 6.

    Tris-buffered saline (TBS): 50 mM Tris Base, 150 mM NaCl, pH 7.4.

  • 7.

    TBS containing 20 μM bovine serum albumin (BSA).

  • 8.

    Biotinylated antigen in TBS.

  • 9.

    TBS containing 0.5% BSA and 0.1% Tween-20 (TBS-T/BSA): 50 mM Tris, 150 mM NaCl, 0.1% (v/v) Tween-20, 0.5% (w/v) BSA, pH 7.4.

  • 10.

    50 mM sodium acetate, 150 mM NaCl, pH 4.0.

  • 11.

    50 mM sodium acetate, 150 mM NaCl, pH 4.5.

  • 12.

    2×YT-Tet broth: see Subheading 2.7.

  • 13.

    2×YT-Carb broth and plates: see Subheading 2.6.

  • 17.

    E. coli XL-1 Blue chemically competent cells.

  • 14.

    M13 helper phage (≥ 1x1012 phage/mL).

  • 15.

    PEG/NaCl: see Subheading 2.2.

  • 16.

    50 mL conical tubes.

2.10. Assessment of single clones using phage ELISA

  1. 96-well deep well plate (2 mL wells).

  2. 2×YT-Carb broth: see Subheading 2.6.

  3. Shaker incubator.

  4. M13 helper phage (≥ 1x1012 phage/mL).

  5. Freshly selected E. coli XL-1 blue cells containing individual phagemid clones.

  6. E. coli XL-1 blue cells containing Wt sdAb phagemid.

  7. 96-well microtiter plate.

  8. Carbonate buffer: 50 mM sodium carbonate, 150 mM NaCl, pH 9.4.

  9. 2 μg/mL antigen in carbonate buffer.

  10. 2 μg/mL BSA in carbonate buffer.

  11. PBS-B: 50 mM Sodium Phosphate, 150 mM NaCl, pH 7.4

  12. PBS-B containing 0.5% BSA and 0.1% Tween-20 (PBS-T/BSA): 50 mM sodium phosphate, 150 mM NaCl, 0.1% Tween, 0.5% BSA, pH 7.4.

  13. Acetate-T/BSA: 50 mM sodium acetate, 150 mM NaCl, 0.1% Tween, 0.5% BSA, pH 4.0.

  14. Anti-M13 antibody conjugated to horseradish peroxidase (HRP) (Sino Biological, Beijing, China).

  15. Tetramethylbenzidine (TMB) substrate.

  16. 1 M sulfuric acid.

  17. Plate reader.

3. Methods

The method to generate pH-dependent sdAbs is a multi-step process allowing the introduction of new protonation events that are linked to antigen binding. Initially, residues within the antigen binding interface are identified, whether through simply identifying hypervariable loop residues or through structural data. Oligonucleotides are then designed and used to generate a combinatorial His/Wt phage library (containing up to 1010 unique members) that samples ionizable His residues within the antigen binding interface. A two-step selection strategy is used to identify variants that maintain strong binding at neutral pH yet display reduced binding affinity at low pH. Variants are then assessed for pH-dependent binding using phage ELISA screening.

3.1. Conceptual framework for combinatorial library and design of degenerate and stop codon oligonucleotides

  1. The combinatorial library is constructed using oligonucleotide-directed mutagenesis (Figure 1) (20). To guide oligonucleotide design, the first step is to identify the sdAb residues that are most likely to form the antigen binding interface. This can be accomplished through sequence alignment to identify residues in the hypervariable loop regions or through structural knowledge of the sdAb/antigen complex, if available. The following steps will help determine how many residues may be targeted in the combinatorial library.

  2. Residue-specific degeneracy. The power of a combinatorial His library is that it includes all possible theoretical combinations of His substitutions within the confines of the library limit. If His is not tolerated at a specific residue position, the Wt residue can be maintained during selection. To generate a His/Wt combinatorial library, degenerate nucleotides are used to produce the minimal codon diversity covering both the original Wt residue and His at a given interface position. The diversity limit for phage display is approximately 1010, consequently, this is the maximum number of unique clones that can be sampled (see Note 2). As a tractable example, for an interface where two tyrosine residues will be targeted to introduce His, the library’s total (codon) diversity would be four using a 50/50 mix of C and T at the first codon position (Figure 2; see Note 3).

  3. Degeneracy considerations. A single oligo can typically cover an entire CDR loop. This means a sdAb combinatorial His library will generally consist of one to three oligos. Figure 3 provides a hypothetical example of His/Wt degeneracy design for a single CDR loop within a sdAb. The codon diversity of this region is 1.6×104, which is calculated from the product of each position’s codon degeneracy. Continuing this example, if two additional degenerate oligos were designed for the two remaining CDRs possessing diversities of 1×103 and 1×102, the total library diversity would be the product of each region’s diversity (i.e., [1.6 × 104][1 × 103][1 × 102]= 1.6 × 109). Notably, this example falls approximately six-fold below the 1×1010 phage display library size limit.

  4. Degenerate oligo design. Once the residue-specific degeneracy is determined, oligonucleotides are designed and synthesized. For each of the potential oligonucleotides, complementary bases at the 5’ and 3’ ends should extend 12-15 bases from the closest degenerate codon. The 3’ end should end in a G/C pairing if possible. An example degenerate oligonucleotide for a combinatorial sdAb CDR1 is illustrated in Figure 4a.

  5. If the diversity of the proposed library exceeds the maximum 1010 diversity limit, the library can easily be refined through reducing the number of degenerate residues. This can include eliminating His/Wt degeneracy at amino acids that are less likely to accommodate His residues, such as proline or glycine, or leave out residues requiring many codon position changes, such as tryptophan (see Note 4).

  6. Once the degenerate oligonucleotide(s) are designed to create the combinatorial His/Wt library, additional oligonucleotide(s) must be designed to introduce stop codons within the stretch of residues subjected to His/Wt screening. The resulting “stop-codon” sdAb serves an important role as an inactive sdAb template when generating the combinatorial library (see Note 5). Using the His/Wt oligos (designed above) as a template, stop codons (TAA) should be introduced at the most 5’ and 3’ His/Wt degeneracy positions, along with every two to three residues in between. This design should ensure the absence of any large regions of complementary bases in between stop codons (see Note 6). As in the design of the degenerate oligonucleotides, complimentary bases at the 5’ and 3’ ends should extend 12-15 bases from the terminal stop codon change and the 3’ end should end in a G/C pairing. An example stop codon insertion oligonucleotide for the CDR1 region is illustrated in Figure 4b.

Figure 1: Overview of oligonucleotide-directed (Kunkel) mutagenesis for combinatorial His scanning library generation.

Figure 1:

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Degenerate oligonucleotide primers, with mutated stretches shown in red, are annealed to a uracil-containing ssDNA template strand (generated from a dut− /ung− bacterial strain). A fill-in reaction is performed using T7 Polymerase, along with a mixture of dNTPs and T4 DNA ligase. This creates a double-stranded DNA product with inserted degenerate mutations on one of the two strands. The double-stranded DNA is then transformed into a dut+ /ung+ E. coli strain (e.g., XL-1 blue), which repairs the DNA to produce either double-stranded DNA containing the mutations of interest or wild-type DNA.

Figure 2: Example of a small-scale combinatorial His scanning library.

Figure 2:

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Assuming two wild-type tyrosine residues are identified as positions to sample His residues, the use of the degenerate base “Y” (i.e., a 50/50 mixture of cytosine and thymine) in the first codon position allows both histidine and wild-type tyrosine to be produced. The total library diversity is calculated by multiplying each position’s codon degeneracy. In this case, 2 × 2 = 4. Image generated using PyMol (23).

Figure 3: Example of combinatorial His scanning library of a sdAb CDR1.

Figure 3:

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For residues 26-31 within CDR1, base degeneracy is designed to incorporate both the wild-type and His residues using base changes that result in the minimum number of codons represented. The higher the codon diversity for a given position, the higher the final diversity (see Note 3). Image generated using PyMol (23).

Figure 4: Example oligonucleotides for producing combinatorial His scanning library.

Figure 4:

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A) Degenerate codons are used with oligonucleotide-directed Kunkel mutagenesis to create a combinatorial library. Each degenerate oligonucleotide should have 12 to 15 complimentary bases on both the 3′ and 5′ ends. B) An inactive sdAb template is created by incorporating multiple TAA stop codons across the sequence where degeneracy is to be introduced. The first and last stop codons should correspond to the first and last degenerate codon positions. Stop codons are inserted every two to three residues to prevent a primer from partially annealing. Image generated using SnapGene® software (Insightful Science; available at snapgene.com).

3.2. Small-scale preparation of uracil-containing single strand DNA Wt template

  1. Warm 960 μL of SOC to room temperature 30 minutes before electroporation

  2. Prepare phagemid vector DNA (containing the sdAb-gene 3 genetic fusion; we use the “drop-out” phagemid vector designed by Kay and coworkers (19)) from a 5 mL overnight culture using a Wizard Miniprep Kit. Incubate a sample of phagemid DNA, a 1 mm gap electroporation cuvette, and a frozen aliquot of CJ236 electrocompetent cells (40 μL) on ice.

  3. Add 2 μL of 150-300 ng/uL phagemid DNA to the bottom/middle of the electroporation cuvette. Then, add the CJ236 cells directly on top of the DNA. Gently tap cuvette to ensure settling and then briefly incubate on ice for 2 minutes.

  4. Set the electroporator’s discharge voltage to 1.83 kV, resistance to 2.5 kV, and resistance timing to 129 ohms.

  5. Wipe off moisture from the outside of the electroporation cuvette with a Kimwipe and insert the cuvette into the electroporator, making sure the cuvette is capped and the protective guard is closed. Electroporate the cells and remove the cuvette when it is safe to open.

  6. Immediately add 960 μL of room temperature SOC medium into the cuvette and slowly pipet the medium up and down to recover the cells. Transfer the recovered solution into a 14 mL Falcon tube and incubate with shaking (235 rpm) at 37°C for 1 hour.

  7. Plate 20 μL and 200 μL of recovered cells on two different 2×YT-Carb/Cam plates. Incubate overnight at 37°C.

  8. The next morning, pick a single colony from the plate using a sterile pipet tip and inoculate 2 mL of 2×YT-Carb/Cam broth. Grow culture at 37°C with shaking until mid-log phase is reached (optical density between 0.5 and 0.8), which should take approximately 6 hours (see Note 7).

  9. When the cells reach mid-log phase, infect the culture with M13K07 helper phage at a final phage concentration of 1010 phage/mL. Incubate the culture at 37°C for 15 minutes with shaking.

  10. Use the 2 mL culture to inoculate 30 mL of 2×YT-Carb/Kan broth. Grow overnight at 37°C with shaking at 235 rpm.

  11. The next morning, transfer the culture to a 50 mL conical tube and centrifuge at 7,000 rpm using a JA-14 rotor with a tube insert (7,500 g) for 10 minutes (4°C). Collect the supernatant in a new centrifuge bottle. To precipitate the phage, add 1/5 of the culture volume of PEG/NaCl solution (6 mL) and mix by inverting tube. Incubate on ice for 1 hour.

  12. Centrifuge the solution at 15,000 rpm using a JA-20 rotor (27,000 g) for 20 minutes (4°C) to pellet the phage. Carefully discard the supernatant without disrupting the pellet. A second, brief 2,000 g centrifugation step should be performed to pool any remaining PEG/NaCl solution, which can then be removed through aspiration with a pipette (see Note 8).

  13. Resuspend the phage pellet with 0.5 mL of PBS-A. Centrifuge sample in a refrigerated tabletop centrifuge at 14,000 rpm (20,800 g) for 10 minutes (4°C) to pellet any insoluble material.

  14. Collect the supernatant into a 1.5 mL microcentrifuge tube and add 7 μL of MP and mix. Allow the solution to incubate for 15 minutes at room temperature to precipitate the phage.

  15. Place a QIAprep 2.0 spin column into a collection tube. Add the precipitated phage sample to the column and centrifuge for 15 seconds at 8,000 rpm (5,900 g) in a tabletop microcentrifuge.

  16. Add 700 μL of MLB to the spin column. Centrifuge at 8,000 rpm (5,900 g) for 15 seconds and discard the flowthrough.

  17. Add 700 μL of MLB again and incubate for about 5 minutes at room temperature (phage lysis). Centrifuge at 8,000 rpm (5,900 g) for 15 seconds and discard the flowthrough. The ssDNA remains adsorbed to the column.

  18. Add 700 μL of PE and centrifuge at 8,000 rpm (5,900 g) for 15 seconds to wash the column (see Note 9). Discard the flowthrough. Repeat the wash step one additional time.

  19. Centrifuge the column again, with no added solution, at 8,000 rpm (5,900 g) for 30 seconds to remove remnants of PE.

  20. Remove the spin column from the collection tube and place in a 1.5 mL microcentrifuge tube. Add 30 μL of EB to the spin column membrane and incubate for 15 minutes. Centrifuge at 8,000 rpm (5,900 g) for 30 seconds to elute the ssDNA. Measure the absorbance at 260 nm to determine the concentration of the ssDNA (Abs260 of 1.0 = 33 ng/μL).

3.3. Stop codon insertion using small-scale Kunkel mutagenesis

  1. Equilibrate a temperature block to 90°C and water baths to 37°C and 50°C.

  2. Resuspend lyophilized primer(s) using nuclease free water to create 1 μg/μL stock solutions. Make a 100 μL working stock of each primer at a final concentration of 330 ng/μL. Store the original stock solution at −20°C.

  3. Phosphorylate each primer individually in microcentrifuge tube(s) by adding 2 μL of 330 ng/μL primer, 2 μL of 10× TM, 2 μL of 10 mM ATP, 1 μL of 100 mM DTT, and nuclease free water to a total volume of 20 μL. Add 1 μL of T4 polynucleotide kinase (20 units) to the microcentrifuge tube. Incubate the solution at 37°C for 1 hour and then immediately move on to the next step.

  4. Anneal oligo(s)/template. In a separate microcentrifuge tube, add the following items in the order listed: 1.0 μg of uracil-containing ssDNA, 2.0 μL of each phosphorylated primer, 2.5 μL of 10× TM, and nuclease free water to a total volume of 25 μL (see Note 10). Incubate the solution at 90°C for 1 minute. Immediately transfer the microcentrifuge tube to the 50°C water bath and incubate for 3 minutes. Finally, transfer the microcentrifuge tube to an ice bucket and incubate for 5 minutes.

  5. Fill-in reaction. Add the following in the order listed to the annealed ssDNA/oligo solution: 1.0 μL of 10 mM ATP, 1.0 μL of 25 mM dNTP mixture, 1.5 μL of 100 mM DTT, 0.6 μL of T4 DNA ligase and 0.3 μL of T7 DNA polymerase. Incubate at 20°C overnight (see Note 11).

  6. Use the Wizard PCR Cleanup kit to purify/desalt the dsDNA product. First, add 250 μL of the Membrane Binding Solution to the reaction and mix (see Note 12).

  7. Insert an SV Minicolumn into a collection tube. Transfer the sample to the SV Minicolumn and incubate at room temperature for 1 minute.

  8. Centrifuge at 16,000 g in a tabletop microcentrifuge for 1 minute and discard the flowthrough.

  9. Add 700 μL of the Membrane Wash Solution to the column and centrifuge at 16,000 g for 1 minute. Discard the flow through.

  10. Add 500 μL of the Membrane Wash Solution to the column and centrifuge at 16,000 g for 5 minutes. Discard the flowthrough.

  11. Without any added Membrane Wash Solution, centrifuge spin column for 1 minute at low speed (10,000 g) with the lid open/off to allow excess residual ethanol to evaporate.

  12. Transfer the SV Minicolumn to a 1.5 mL microcentrifuge tube.

  13. Add 35 μL of nuclease-free water directly to the center of the membrane. Incubate at room temperature for 1 minute and then centrifuge at 16,000 g for 1 minute.

  14. Determine the concentration of the final dsDNA product by measuring its absorbance at 260 nm. The DNA sample should be stored at −20°C.

  15. Transform E. coli XL-1 Blue competent cells with the mutagenic DNA. Add 2 μL of the Kunkel product from step 6 to the bottom of a 14 mL round-bottom Falcon culture tube. Add 40 μL of E. coli XL-1 Blue chemically competent cells (that have been thawed in an ice/water slurry) directly on top of the DNA. Incubate on ice for 30 minutes. During this time, ensure SOC medium is at room temperature.

  16. Hold the culture tube in a 42°C water bath for 45 seconds before immediately transferring the culture tube back to the ice bucket. Incubate on ice for 2 minutes.

  17. Add 300 μL of room temperature SOC medium and incubate at 37°C with shaking (235 rpm) for 1 hour.

  18. Plate 20 μL and 200 μL of the transformed cells on two different LB-Carb plates and incubate overnight at 37°C.

  19. The next morning, ensure single colonies are visible on the plates and then store at 4°C. In the late afternoon, using individual colonies, start several 5 mL cultures in LB-Carb broth. Depending on whether one, two, or three oligonucleotides were used in the Kunkel reaction, start 4, 10, or 15 cultures, respectively. Grow cultures overnight at 37°C with shaking.

  20. The next morning, perform a plasmid purification from all overnight cultures using a Wizard Miniprep Kit following the manufacturer’s instructions. It is recommended to use 30 μL of DNase-free water to elute the DNA. Submit samples of DNA for Sanger sequencing using an appropriate sequencing primer.

  21. Examine sequencing results to identify sample(s) that contain all of the designed stop codon insertions. This is often facilitated by performing a multiple sequence alignment. Use the identified stop codon containing dsDNA to produce the ssDNA template as described in Subheading 3.4.

3.4. Large-scale preparation of uracil-containing single strand DNA stop codon template

Starting with the sequence-verified dsDNA stop codon template (in place of the phagemid vector), follow the procedure from subsection 3.2, steps 1 through 9. Then, complete the preparation by following described below.

  1. Use the 2 mL culture to inoculate 200 mL of 2×YT-Carb/Kan broth. Grow overnight at 37°C with shaking at 235 rpm (see Note 13).

  2. The next morning, spin down the bacteria at 7,000 rpm using a JA-14 rotor (7,500 g) for 10 minutes at 4°C. Collect the supernatant in a new centrifuge bottle. To precipitate the phage, add 1/5 of the culture volume of PEG/NaCl solution (i.e., 40 mL). Incubate on ice for 1 hour.

  3. Spin the solution at 13,000 rpm using a JA-14 rotor (25,000 g) for 20 minutes at 4°C to pellet the phage. Before discarding the supernatant, ensure that a faint phage pellet is visible. A second, brief 2,000 g centrifugation step should be performed to pool any remaining PEG/NaCl solution, which can then be removed through aspiration with a pipette (see Note 8).

  4. Resuspend the phage pellet with 6 mL of PBS-A, transfer to multiple microcentrifuge tubes, and spin down at 14,000 rpm (20,800 g) for 10 minutes at 4°C using a refrigerated tabletop centrifuge to pellet any insoluble material.

  5. Collect the supernatant into a 15 mL Falcon Tube. Add 60 μL of MP and mix to precipitate the phage particles. Incubate at room temperature for 15 minutes.

  6. Place two QIAprep 2.0 spin columns into two collection tubes. Add 700 μL of the sample to each column and centrifuge for 15 seconds at 8,000 rpm (5,900 g). Discard the flow-through and repeat until all the sample has been loaded and run through the columns (see Note 14).

  7. Perform steps 16 through 19 as described in subsection 3.2.

  8. Remove the spin columns from the collection tube and place in microcentrifuge tubes. To each, add 100 μL of nuclease free water directly to the spin column matrix and incubate for 15 minutes. Centrifuge at 8,000 rpm (5,900 g) for 30 seconds to elute the ssDNA. Measure the absorbance at 260 nm to determine the concentration of the ssDNA (Abs260 of 1.0 = 33 ng/μL).

3.5. Large-scale in vitro synthesis of heteroduplex degenerate dsDNA

Start by repeating the procedure from subsection 3.3 steps 1 through 3 using the degenerate His/Wt library primers (designed in subsection 3.1 step 4). Then, complete the preparation by following the steps described below.

  1. Anneal oligo(s)/template. In a separate microcentrifuge tube, add the following items in the order listed: 20.0 μg of uracil-containing stop-codon template ssDNA, 20 μL of each phosphorylated primer, 25 μL of 10× TM, and nuclease-free water to a final volume of 250 μL (see Note 10). Incubate the solution at 90°C for 1 minute. Immediately transfer the microcentrifuge tube to the 50°C water bath and incubate for 3 minutes. Finally, transfer the microcentrifuge tube to an ice bucket and incubate for 5 minutes.

  2. Fill-in reaction. Add the following in the order listed to the annealed ssDNA/oligo solution: 10 μL of 10 mM ATP, 10 μL of 25 mM dNTP mixture, 15 μL of 100 mM DTT, 6.0 μL of T4 DNA ligase and 3.0 μL of T7 DNA polymerase. Incubate at 20°C overnight (see Note 11).

  3. Use the Wizard PCR Cleanup kit to purify/desalt the dsDNA product as described in subsection 3.3 steps 6-13 with the following modifications. Initially, add 1 mL of the Membrane Binding Solution to the large-scale Kunkel reaction. Then, split and load the solution across two SV Minicolumns. Carryout the protocol as described in steps subsection 3.3 steps 7-14 for the two samples in parallel. Pool final purified dsDNA samples (see Note 15).

  4. Perform steps 15 through 18 as described in subsection 3.3 with the exception that 1 μL of the Kunkel product DNA can be used in the XL-1 Blue transformation. Then, follow the steps below to characterize the success of heteroduplex degenerate dsDNA synthesis.

  5. In addition to plating the transformed cells, inoculate 5 mL of LB-Carb broth with 10 μL of the transformed cells and incubate overnight at 37°C with shaking (235 rpm). This represents the mixed population sample.

  6. The next morning, ensure single colonies are visible on the plates and then store at 4°C. In addition, isolate plasmid DNA from the overnight 5 mL mixed population culture using a Wizard Miniprep kit. Elute the mixed population DNA with 30 μL of nuclease-free water and store at −20°C.

  7. In the late afternoon, identify easy to pick individual colonies and start several 5 mL LB-Carb cultures. Depending on whether one, two, or three oligonucleotides were used in the Kunkel reaction, start at least 4, 10, or 15 cultures, respectively. Grow cultures overnight at 37°C with shaking.

  8. The next morning, isolate plasmid DNA from the overnight cultures using a Wizard Miniprep Kit following the manufacturer’s instructions. It is recommended to use 30 μL of DNase-free water to elute the DNA. Submit samples of DNA from the individual clones, as well as the mixed population sample, for sequencing using an appropriate sequencing primer.

  9. Evaluation of mutagenesis success through sequencing. The sequences of both the mixed population and individual clones provide insight into mutagenesis efficiency. For the mixed population, the sequencing chromatogram (which shows profiles for each base G, C, A, and T) can be used for a crude, yet base pair specific evaluation of base frequency at each site of degenerate incorporation. For example, for a position where degenerate base “Y” is inserted, the chromatogram should reveal an approximate 50/50 appearance of both bases T and C (see Note 16). For the individual clones, determine the approximate success rate of full degenerate base pair incorporation. For multi-oligonucleotide libraries, calculate by counting the number of clones that suggest all primers incorporated library substitutions, thus removing all stop codons, and dividing by the total number of individual colonies sequenced (see Note 17).

  10. Determine the maximum diversity coverage of the library, based on the expected phage display maximum diversity, 1×1010, which represents a practical upper limit for unique electroporation/phage particle production. Multiply the fraction of library that was successfully mutated (calculated above) by the diversity limit, 1×1010. For instance, if 30% of your library was estimated to contain the designed coverage, then 3×109 is the maximum diversity that can be covered assuming 1×1010 total diversity can be reached during the electroporation/phage particle generation. If the maximum diversity coverage of the generated library is larger than the theoretical diversity of the combinatorial library (calculated in section 3.1), then proceed to phage generation (see Note 18).

3.6. Phage display library generation

The last step in producing the combinatorial His scanning library involves transforming E. coli SS320 cells, a highly efficient competent strain containing the F’ episome, with the heteroduplex dsDNA library. This enables infection by M13 bacteriophage resulting in phage production/packaging of the library. If needed, methods to amplify M13KO7 helper phage and prepare electrocompetent E. coli SS20 cells have been described by Sidhu and coworkers (21). Before electroporating E. coli SS320 cells with the library, it is recommended to determine the electroporation and transformation efficiency with a less valuable DNA sample, such as the original sdAb phagemid template. This dry run should be used to determine the maximum number of transformants that may be achieved by electroporation of E. coli SS320 cells.

  1. Incubate SOC medium in a water bath at 37°C. Place two 2×YT-Carb plates in a 37°C incubator. Thaw the heteroduplex degenerate phagemid dsDNA library on ice and split into two to three aliquots in microcentrifuge tubes, each containing 4 to 7 ug of DNA. Incubate a 2 mm gap electroporation cuvette on ice. Thaw two to three aliquots (350 μL each) of electrocompetent E. coli SS320 cells in an ice/water slurry.

  2. Transfer an aliquot of SS320 cells to one of the aliquots of degenerate phagemid dsDNA. Using a 1 mL pipettor, gently mix sample three times without introducing any air bubbles. Incubate the mixture for 2 to 3 minutes.

  3. Gently pipette the mixture to the center of a 2 mm gap electroporation cuvette without introducing bubbles and secure in the electroporation system (see Note 19).

  4. Set the field strength to 2.5 kV, the resistance to 129 Ω and the capacitance to 50 μF. Electroporate the sample.

  5. Immediately rescue the cells by adding 1 mL of 37°C SOC. Transfer to a 250 mL baffled flask. Repeat two to three times, rinsing the cuvette with SOC and transferring to the 250 mL flask. Finally, add SOC until a final volume of 25 mL is reached.

  6. Incubate the culture at 37°C for 20 minutes with shaking (200 rpm).

  7. Take 100 μL of the culture and set aside for serial dilutions (described below). The remaining ~25 mL of culture should be added to a 4 L baffled flask containing 500 mL of 2×YT-Carb broth and 1010 M13KO7 helper phage/mL. Incubate the culture overnight at 37°C with shaking (200 rpm) (see Note 20).

  8. With the 100 μL of culture set aside, perform serial dilutions to determine the library transformation efficiency. In a sterile 96 well plate, aliquot 90 μL of SOC medium across 12 wells. Add 10 μL of the culture stock to the 90 μL of SOC medium in the first row and mix. This represents a 10−1 dilution. Next, take 10 μL from the 10−1 diluted well and mix with the 90 μL of SOC medium in the second row to generate the 10−2 dilution. Repeat this until 12 ten-fold dilutions are made.

  9. Using a permanent marker, take the two 2×YT-Carb plates and mark up six evenly divided sections on the back of the plate. This is often most easily accomplished by drawing three straight lines to create six “pie slices.” Label each sector a dilution value, i.e., 10−1 through 10−12, across the two plates. Plate 20 μL from each dilution well into its corresponding section on the two plates. In addition, add 30 μL of the sterile SOC, used to dilute samples, to the center of the plate to serve as a control. Dry the plate in a sterile environment until no liquid remains. Incubate overnight at 37°C.

  10. The next morning, determine the total number of transformants and isolate phage particles.

  11. Examine the dilution series plates and calculate the number of bacteria transformed (i.e., antibiotic resistant colonies, due to uptake of the phagemid) in the 25 mL culture. For example, if 8 colonies are visible on the 10−6 dilution sector, the total number of transformants would be calculated as: 8 transformants × 50 mL−1 × 106 dilution factor × 25 mL = 1×1010 total transformants. If the number of cells is larger than the diversity of the library, then the entire library will be displayed (see Note 21).

  12. Centrifuge the 500 mL culture at 8,000 rpm using a JA-10 rotor (11,000 g) for 10 minutes at 4°C.

  13. Transfer the supernatant to fresh 500 mL centrifuge bottles and add 100 mL (representing 1/5 the culture supernatant volume) of PEG/NaCl. Mix and incubate on ice for 1 hour to precipitate the phage.

  14. Centrifuge the sample at 13,000 rpm using a JA-14 rotor (25,000 g) for 20 minutes (4°C). Discard the supernatant and centrifuge again at 2,000 g for 2 minutes to pool any remaining supernatant. Remove the rest of the supernatant with a serological pipet.

  15. Resuspend/resolubilize the phage pellet in 25 mL of PBS-A. Transfer to a new centrifuge tube and centrifuge at 15,000 rpm using a JA-20 rotor (27,000 g) for 20 minutes (4°C) to pellet any insoluble matter. Transfer the supernatant to a sterile 50 mL conical tube and store at 4°C for immediate use or −80°C for long-term storage.

  16. Phage concentration can be estimated using UV absorbance; OD (268 nm) = 1.0 for 5×1012 phage/mL (see Note 22).

3.7. Phage titering

Phage titering provides a quantitative assessment of the phage concentration, which is useful in evaluating the multi-step process of phage selection. As such, phage titers are typically performed on the original phage stock, as well as input and output phage samples during selection.

  1. Start a 2 mL culture of E. coli XL-1 Blue cells in 2×YT-Tet (14 mL Falcon culture tube) and grow at 37°C overnight with shaking.

  2. The next morning, inoculate 2 mL of fresh 2×YT-Tet with 4 μL of the overnight culture. Grow the new culture until mid-log phase is reached (target mid-afternoon). The target cell optical density should be between 0.5 and 0.8 for titering. If cell growth happens to go above this window, dilute cells (e.g., 10 to 20-fold) in fresh media and continue to grow until mid-log phase is reached.

  3. In a 96-well plate, make 12 10-fold serial dilutions of the phage solution (e.g., phage library). Add 90 μL of PBS-A to 12 wells. Then, add 10 μL of the phage stock to the first well and gently mix to generate the 10−1 dilution. Replace the filter tip before aspirating 10 μL of the diluted (10−1) phage solution and add it to the next well. Mix the dilution to generate the 10−2 dilution and replace the tip. Continue to repeat this process to generate the complete 12 10-fold dilution series.

  4. Pour the mid-log phase E. coli XL-1 blue cells into a sterile multi-channel pipet basin. Pipet 30 μL of E. coli XL-1 blue cells into 12 empty wells across the 96-well pate (parallel to the phage dilutions prepared above).

  5. Carefully add 10 μL of each of the 12 diluted phage samples into the 30 μL E. coli XL-1 blue cell samples. Incubate for 15-25 minutes at room temperature for phage infection.

  6. Using a permanent marker, take the two 2×YT-Carb agar plates and mark up six evenly divided sections on the back of each plate. This is often most easily accomplished by drawing three straight lines to create six “pie slices.” Label each sector a dilution value, i.e., 10−1 through 10−12, across the two plates. Carefully pipet 40 μL from each of the diluted infection wells into the confines of their specific dilution section. Pipet 30 μL of the uninfected E. coli XL-1 blue cells in the center of one plate, and 30 μL of the sterile PBS-A into the center of the second plate, which serve as controls. Dry in a sterile environment by an open flame. Incubate at 37°C overnight.

  7. The next morning, evaluate the most dilute sector that has clearly defined (countable) colonies and check that no colonies are observed in the controls. Calculate the phage titer of the original stock (in phage/mL). For example, if 10 colonies were observed in dilution sector 10−9, the phage titer would be 10 phage × 100 mL−1 ×109 = 1×1012 phage/mL.

3.8. Biotinylation of antigen

  1. Generate 1 mL of a 2 mg/mL solution of the antigen of interest in reaction buffer (100 mM sodium phosphate, 150 mM NaCl, pH 7.2).

  2. Ensure the antigen’s pre-existing buffer does not contain primary amines (e.g., Tris), which will interfere with the reaction. If present, dialyze the antigen solution against reaction buffer.

  3. Prepare a fresh 10 mM solution of EZ-Link Sulfo-NHS-SS-Biotin by adding 6 mg of the reagent to 1 mL of pure water.

  4. Add the dissolved EZ-Link Sulfo-NHS-SS-Biotin to the antigen solution and incubate on ice for 2 hours (see Note 23).

  5. Remove the unreacted biotin reagent by dialysis against 2 L of PBS-A (see Note 24).

  6. Store the biotinylated protein at 4°C for immediate use or flash freeze at −80°C for long-term storage.

3.9. Phage display panning with double selection

Prior to starting phage display panning, it is important to ensure that the transformation efficiency in library generation exceeded the theoretical library diversity. In addition, the library phage titer should be high enough to allow the addition of an amount of phage particles at least 100-fold higher than the theoretical library diversity, as only 1%-5% of the phage particles will display sdAb variants, thus ensuring full library coverage. It is also recommended to perform a “pull-down” test of the biotinylated antigen using the SAV beads. The ability to pull-down biotinylated antigen (versus unbiotinylated antigen) can be confirmed using SDS-PAGE or spectroscopy (monitoring the protein’s absorbance at 280 nm). This test can be extended to include pulling down biotinylated antigen/sdAb complex, which can ensure biotinylation events do not interfere with the antigen binding site.

The phage display selection, described below, is meant to serve as a guide to produce pH-sensitive sdAb variants. An overview of the two-step selection process is presented in Figure 5. The level of stringency of selection (e.g., target antigen concentration or pH) can be increased/decreased based on the enrichment readout values. In addition, to perform phage titers (described in section 3.7) the same day of selection, it is important to start overnight E. coli XL-1 Blue cultures the night before that day’s selection.

  1. Dispense 200 μL of resuspend SAV beads into a microcentrifuge tube. Perform a brief 10 second centrifuge spin at 5,000 g and place on the magnetic stand for 1 minute. Ensure supernatant is free of beads and gently remove supernatant avoiding beads. Remove the microcentrifuge tubes from the magnetic stand and resuspend/wash beads using 1 mL of TBS-T/BSA. Repeat this wash process three times. During the final wash, split the sample into two evenly split aliquots. These will be used for sample and control.

  2. For the first round of selection, add an amount of phage that represent at least 100-fold excess phage over the theoretical library diversity. For a combinatorial His scanning library with a max diversity of 1×1010, add 1 mL of a 1012/mL titer library to two different microcentrifuge tube (sample and control). Dilute a concentrated sample of biotinylated antigen to a final concentration of 2 μM (sample). For the control, add BSA to a final concentration of 2 μM. Incubate both at room temperature for 20 minutes with gentle rotation/rocking.

  3. Add 100 μL of the washed beads to each tube (sample and control) and incubate both at room temperature for 15 minutes with gentle rotation/rocking.

  4. Briefly centrifuge the sample and control for 10 seconds at 5,000 g in a tabletop centrifuge, before placing them on the magnetic stand for 1 minute. When the supernatant is clear, remove all but 200 μL of the supernatant and resuspend the beads.

  5. Transfer the sample and control samples to two separate row A wells in a Kingfisher magnetic bead handler. For each sample, add 200 μL of TBS-T/BSA to five wash wells. Finally, add 200 μL of sodium acetate, pH 4.0, which is used to select for pH sensitive clones, to a final well.

  6. Program the Kingfisher to transfer the SAV beads successively to each wash well, incubating with mixing for 20 seconds before transferring to next wash well. In the final transfer into the pH-sensitive selection well, allow beads to incubate with mixing for 10 minutes before removing beads (see Note 25).

  7. Take 10 μL of the final low pH well samples (both target and control) and determine the output titer of each. Use 90 μL of the low pH well sample for amplification (described below). Store the remaining low pH phage selection sample at 4°C.

  8. To amplify the output phage, add 90 μL of the low pH sample to 2 mL of mid-log phase E. coli XL-1 Blue cells in 2×YT-Tet and incubate at 37°C for 20 minutes with shaking (200 rpm).

  9. Add the 2 mL infected culture to 30 mL of 2×YT-Carb broth, containing 1×1010 M13KO7 helper phage/mL. Grow overnight at 37°C with shaking at 235 rpm.

  10. The next day, centrifuge the 30 mL overnight culture at 8,000 rpm for 10 minutes (4°C).

  11. Transfer the supernatant to a 50 mL conical tube and add 6 mL of PEG/NaCl and incubate on ice for 1 hour. Centrifuge for 20 minutes at 9,000 g and 4°C and discard the supernatant. Perform a second brief 30 second centrifugation at 2,000 g, if necessary, to pool any remaining PEG/NaCl solution and remove by aspiration (see Note 26).

  12. Resuspend the phage pellet in 1 mL of TBS. Centrifuge at 14,000 rpm (20800 g) for 10 minutes (4°C) to pellet insoluble material using a refrigerated tabletop centrifuge. Store the supernatant at 4°C.

  13. The morning after completing each round of phage selection, use the overnight titer plates to calculate the enrichment by dividing the phage count from the sample output by the control output. Since a combinatorial library is likely to have a sizeable sub-population capable of binding the higher first round antigen concentration, along with pH sensitivity, the enrichment ratio may be quite high (e.g., > 100) in the first round. The enrichment may even decrease with each round, as selection stringency also increases (e.g., enrichment values 10 to 90s). If the enrichment ratio is close to a value of 1.0 after the 2nd round or beyond, it is likely that the selection stringency may be too high and selection criteria can be made less stringent (e.g., smaller drop in antigen concentration or keeping low pH selection at pH 4.0).

  14. The remaining round of phage selection (second round and beyond) can be performed entirely automated using the Kingfisher. For round two, in two row A wells, add 100 μL of amplified phage to either 100 μL of 0.2 μM biotinylated antigen (sample) or 0.2 μM BSA (control). In row B wells add 100 μL of washed beads in a total of 200 μL TBS-T/BSA. In row C through G wells add 200 μL of TBS-T/BSA. In row H add 200 μL of sodium acetate, pH 4.5. Program the Kingfisher to incubate Row A wells with mixing for 15 minutes, before transferring SAV Beads. Capture SAV Beads with mixing for 15 minutes. Then, transfer beads successively to each wash well C through G, incubating with mixing for 20 seconds before transferring to next wash well. The final transfer to the pH-sensitive selection (wells H) should include incubation with mixing for 10 minutes before removing SAV Beads.

  15. Amplify the sample output for a third round of selection by following steps 8-12. Perform a phage titer for the amplified phage input, as well as both sample and control output wells using 10 μL of each sample.

  16. In the third round of selection, repeat steps 13-15. However, the biotinylated antigen and BSA control concentrations can be reduced 10-fold to 20 nM and the pH sensitive selection buffer can be raised to pH 5.0 (i.e., if the enrichment is not too close to 1.0).

  17. A fourth round of selection may be pursued further increasing the stringency of binding at pH 7.4 (by reducing antigen concentration) or pH sensitivity (by increasing pH or decreasing contact time).

Figure 5: Overview of phage display double selection method.

Figure 5:

Open in a new tab

Starting with a combinatorial His scanning phage library, which contains wild-type, non-functional, and pH dependent sdAb variants, two selection steps are performed. Selection 1 involves collecting phage displaying sdAbs that bind tightly to the antigen at the desired pH (e.g., pH 7.4). This is followed by selection 2, which collects phage displaying sdAb variants that possess binding sensitive to the pH change (e.g., pH 4.0). The identified phage particles are then amplified and subjected to additional rounds of screening (2-3 additional rounds).

3.10. Assessment of pH sensitive binding using phage ELISA

  1. Using up to 23 individual colonies from the third and/or fourth round output clones, inoculate individual 300 μL cultures of 2×YT-Carb media containing 1010 M13K07 helper phage/mL within a 96-well deep well plate. Repeat this process for a colony that was transformed/infected with the Wt sdAb phagemid to serve as a control. Mount the deep-well plate at a slight angle and incubate at 37°C overnight with shaking at 235 rpm (see Note 27).

  2. The same day use carbonate buffer to dilute concentrated stocks of antigen and BSA to final working concentrations of 2 μg/mL. Each individual clone will require four separate wells to test binding at two pH values (each pH includes a control well). For each clone including a Wt sdAb control (up to 24), add 100 μL of the antigen or BSA solution into two wells of a 96-well microtiter plate. Store plate overnight at 4°C.

  3. The next morning, centrifuge the culture plate for 10 minutes at 3,700 rpm (1,400 g) using a swinging bucket rotor and collect the phage-containing supernatant for each sample. Store at 4°C until ready to use.

  4. Continue microtiter plate preparation by removing the overnight coating solutions by inverting and then gently stamping on a paper towel to remove excess liquid.

  5. Next, block all wells by adding 200 μL of PBS-T/BSA to all sample and control wells. Incubate at room temperature with gentle shaking for 2 hours. Invert plate to remove liquid.

  6. Wash plate five times with PBS-T/BSA. Invert to remove liquid and gently stamp on paper towels between each wash to remove remaining liquid.

  7. To prepare phage samples, using a 96-well deep well plate, add 100 μL of each phage-containing supernatant collected in step 3 to 200 μL of PBS-T/BSA to dilute the phage into pH 7.4 buffer. Repeat this process with acetate-T/BSA to dilute the phage into pH 4.0 buffer.

  8. Add 100 μL of the diluted phage (whether pH 7.4 or pH 4.0) to the relevant sample and control wells. Incubate at room temperature for 1 hour with shaking.

  9. After incubation, wash the plate three times with PBS-T/BSA (for pH 7.4 wells) or acetate-T/BSA (for pH 4.0 wells). Then, wash all wells two times with PBS-T/BSA.

  10. Generate a dilution of an anti-M13 antibody HRP conjugate in the range of 1:2,500 to 1:4,000 using PBS-T/BSA. Add 100 μL of the diluted antibody to each well and incubate at room temperature with shaking for 30 minutes (see Note 28).

  11. Wash all wells four times with PBS-T/BSA and twice with PBS-B.

  12. Add 100 μL of TMB to each well and incubate at room temperature for 10 minutes. Quench reaction with 100 μL 1 M sulfuric acid and immediately measure the absorbance at 450 nm (see Note 29).

  13. Relative to background signals and the wild-type clone, evaluate the binding response for the two pH conditions to assess pH dependent sdAb clones.

Acknowledgment

This work is supported by NIH grant 1R15GM124607 to J.R.H.

4. Notes

1.

QIAprep 2.0 spin columns may be purchased separately from the kit.

2.

The total number of unique clones should be calculated by counting codons for the wild-type and His residues, as well as any additional codons that are introduced in the degenerate combination to include His.

3.

Most amino acid residues can be exchanged for His with only one or two nucleotide changes, resulting in 2 or 4 codons. Some residues, such as tryptophan, require additional degeneracy and may even introduce stop codons. Introduction of stop codons is allowed as long as total diversity is below the maximum number of phage produced. In addition, it is ideal to avoid using stop codons that are not frequently used by E. coli.

4.

Ultimately, the 1010 library limit stems from limitations in electroporation of E. coli SS320 cells with the degenerate phagemid vector (Section 3.6 step 4). It is important not to exceed the 1010 limit, otherwise one hundred percent coverage of the library is not achieved. It is recommended that prior to electroporation of the library, the user performs a test electroporation including serial dilutions to determine electroporation efficiency and verify that 1010 transformants can be reached. The library diversity can always be reduced to be below the experimental electroporation number. In situations where the designed library exceeds the upper limit of 1010, the user may choose to probe only one or two regions (e.g., CDR1 and CDR3). This approach has the advantage of selecting potential intraloop cooperative effects, which tend to stem from nearby residues.

5.

Since Kunkel mutagenesis is not 100% efficient, the use of a stop-codon template ensures only newly mutated, i.e., combinatorial mutant VHH variants, will produce full VHH polypeptides, thus eliminating Wt bias.

6.

By leaving large stretches of complementary base pairs between stop codons, the problem can arise of a mixture of a primer properly annealing and primer partially annealing to the template strand.

7.

When starting from a single colony, culture growth can be less predictable; therefore, it is often convenient to start more than one 2 mL culture.

8.

The phage pellet is a faint white color. It is advised to carefully note the expected location where the pellet should form to avoid inadvertent disruption. If a phage pellet is accidently disrupted or difficult to observe, collect the supernatant and repeat centrifugation.

9.

Make sure ethanol has been added to the PE buffer before use.

10.

On a molar scale, the oligonucleotide:template ratio should be 3:1. Values provided assume an oligonucleotide:template length ratio of approximately 1:100.

11.

The reaction can be stored in a −20°C freezer when complete.

12.

A QIAquick DNA purification kit (Qiagen) may be used instead.

13.

Work by the Kay lab suggested that growing at 25°C can enhance single strand DNA yield by 2- to 7-fold (22).

14.

Multiple tubes are used to purify the ssDNA as the ssDNA yield is often higher than the capacity of a single column (sometimes even more than two or three columns). The resulting uracil-containing stop codon ssDNA template may be used for additional libraries, if stop codon coverage is suitable. For a single combinatorial library generation, a yield of 20 μg of ssDNA is suggested.

15.

Two spin columns are used due to the high amount of DNA present. For some high yield phagemids, three or four spin columns may be used for maximum recovery.

16.

Kunkel mutagenesis is approximately 50%−80% efficient; therefore, it is expected to observe the original base at some frequency.

17.

Increasing the number of clones sent for sequencing will increase the reliability of the estimates of mutagenesis success efficiency. Modern high throughput sequencing may also be used to greatly enhance sampling/reliability. For troubleshooting, it may also be of interest to determine each primer’s success rate.

18.

The total library size of 1×1010 represents the “typical” upper limit for a routine phage display preparation and is dependent on the efficiency of the electroporation of E. coli SS320 cells with the dsDNA library and subsequent packaging into phage particles. It is important that the user establish they can reach this threshold (or what the “actual” threshold may be running through the protocol). The user can alter the size of the library, if needed. For those less familiar with phage display, one may want to choose to sample fewer residues and produce a library with a lower theoretical diversity. For example, if the final library theoretical diversity was 1×107, there would be plenty of headspace (103) to compensate for suboptimal electroporation.

19.

To avoid introducing bubbles into the system, avoid using the pipette’s second stop “blow out”.

20.

If using a phagemid with an isopropyl β-D-1-thiogalactopyranoside (IPTG) inducible promoter, IPTG should be included in the 500 mL culture.

21.

It is important to stress that the number of experimental transformants sets the limitation on library coverage. For example, if a library had a total theoretical diversity of 1×1010, yet only 1×108 transformants were achieved, only 1% of the designed library would be sampled moving forward. This fractional coverage would not yield suitable results for the combinatorial His scanning approach.

22.

Be mindful of potential phage contamination and proper decontamination.

23.

For antigen concentrations of ~ 1 mg/mL, a greater than 20-fold molar excess of EZ-Link Sulfo-NHS-SS-Biotin reagent to antigen is recommended. For a 15 kDa hypothetical antigen under these conditions, this ratio is 75:1. Add this volume to the antigen solution and incubate on ice for 2 hours.

24.

Alternatively, size exclusion chromatography or desalting columns may be used to remove unreacted biotin.

25.

If a Kingfisher is not available, selection washes and incubations may be carried out in a microcentrifuge tube. Brief 10 second centrifugation spins followed placing in a magnetic stand allows efficient condition changes.

26.

The use of disposable 50 mL conical tubes helps reduce the chance of phage contamination; however, it is important to verify the maximum g force to avoid potential tube collapse/shattering. Not all conical tubes can tolerate higher g force. Alternatively, Oakridge tubes and a JA-20 rotor can be used.

27.

While we have had success growing small scale cultures in deep well plates, sometimes larger phage yields may be necessary, which can be generated by increasing the overnight culture volume to 5 mL.

28.

While our early work used a GE Biosciences anti-M13-HRP antibody conjugate, it is no longer manufactured. In our hands, the anti-M13-HRP antibody conjugate from Sino Biological produces lower signal/noise. It may be necessary to explore other anti-M13-HRP antibodies or make use of alternate tags, such as anti-FLAG antibodies for detection.

29.

The absorbance can be measured at 652 nm before quenching.

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