Protocol for Cloning Peptide Display Libraries in M13KE


  1. Design a library oligonucleotide following the convention. Bear in mind that the sequence VPFYSHS preceding the leader peptidase cleavage site is part of the pIII signal sequence and should not be altered. The first residue of the displayed peptide will immediately follow this sequence. For randomized positions, relative representations of each amino acid can be improved by limiting the third position of each codon to G or T (= A or C on the synthetic library oligonucleotide). We recommend including a short spacer sequence between the randomized segment and the first native pIII residue to improve target accessibility to the displayed peptide, e.g. the spacer Gly-Gly-Gly between the random peptide and the Ser-Ala-Glu (SAE). The oligonucleotide should be synthesized on a minimum of 0.2 µmol scale, gel-purified (1), and accurately quantitated by measuring the OD260 in a spectrophotometer (1 absorbance unit at 260 nm = 20 µg/ml of single stranded DNA).

  2. Anneal 5 µg of the library oligonucleotide with 3 molar equivalents of the universal extension primer 5´-CATGCCCGGGTACCTTTCTATTCTC-3´ (approximately 4 µg for a 90-nucleotide library oligonucleotide) in a total volume of 50 µl TE containing 100 mM NaCl. Heat to 95°C and cool slowly (15–30 minutes) to less than 37°C in a thermal cycler or water bath.

  3. Extend the annealed duplex as follows (mix in the given order)
    H2O: 119 µl
    10X NEBuffer 2: 20μl
    annealed duplex: 50μl
    10 mM dNTP's: 8μl
    Klenow fragment (NEB #M0210 ) (5 Units/µl ): 3μl
    Total: 200 µl
    Incubate at 37°C for 10 minutes, then 65°C for 15 minutes. Save 4 µl for later analysis (Step 5). 

  4. The extended duplex is then cut using EagI and either, KpnI or Acc65I. We favor digestion as follows:
    Extension reaction: 196 µl
    H2O: 158 μl 
    10X NEBuffer 2: 40 μl
    EagI-HF (NEB #R3505 ) (10 Units/µl ): 5 μl 
    KpnI (NEB #R0142 ) (10 Units/µl ): 6 μl
    Total: 400 µl 

    Extension reaction: 196 µl
    H2O: 154 μl
    10X NEBuffer 4: 40 μl
    EagI-HF (NEB #R3505 ) (10 Units/µl ): 5 μl 
    KpnI-HF (NEB #R3142 ) (10 Units/µl ): 5 μl
    Total: 400 µl
    Incubate at 37°C for 3–5 hours.* Purify the DNA by phenol/chloroform extraction, chloroform extraction and ethanol precipitation. 
    *EagI-HF is not recommended for > 1 hour digestions.

  5. Gel-purify the digested duplex on an 8% nondenaturing polyacrylamide gel (48), including as molecular weight markers, 4 μl of the undigested duplex as well as Low Molecular Weight DNA Ladder (NEB #N3233 ). Visualize by ethidium bromide staining, and excise the digested duplex from the gel, minimizing UV exposure time. Mince the excised band and elute the DNA by shaking overnight in several volumes of 100 mM sodium acetate, pH 4.5, 1 mM EDTA, 0.1% SDS at 37°C.

  6. Briefly microfuge to separate the gel fragments from the elution buffer, and transfer the supernatant to a clean tube. Repeat wash to improve yield, if desired. Purify the DNA duplex from the supernatant by phenol/ chloroform extraction, chloroform extraction and ethanol precipitation (48). Resuspend the pellet in 50 μl of TE and quantitate a small amount by PAGE or spectrophotometrically. One μg of purified insert is more than sufficient for a library of complexity 109.

  7. For a high-complexity library, digest 10–20 µg of M13KE Vector with the same enzymes used to prepare insert in step 4, for example, using 10 units each of EagI-HF and KpnI per µg DNA (total volume = 400–800 µl). Digest 3–5 hours at 37°C. Use KpnI in place of Acc65I if you used KpnI for insert digestion. Gel purify using standard methods (β-Agarase, QIAGEN®, etc). Quantitate a small amount of purified cut vector on an agarose gel or spectrophotometrically.

  8. Optimize the ligation conditions. Suggested starting parameters per 20 µl ligation: 40 and 100 ng of cut vector; 3:1, 5:1 and 10:1 molar excess of cut duplex; 2 µl of 10X ligase buffer; and 200 units (= 3 Weiss units) of T4 DNA Ligase (NEB #M0202 ). Incubate overnight at 16°C.

  9. Heat-kill the test ligations at 65°C for 15 minutes, then electroporate 1 µl of each into 100 µl of electrocompetent ER2738 or other F+ strain (e.g. NEB Turbo, NEB #C2986 ). For Bio-Rad Gene Pulser, use 25 μF, 200 Ω and 2.5 kV in a 2mm cuvette as electroporation parameters. Outgrowths are carried out in 1 ml of SOC medium for 30–45 minutes at 37°C with shaking. 

    SOC: Dissolve 20 g tryptone, 5 g yeast extract, 0.5 g NaCl in 950 ml deionized H2O. Add 10 ml of 250 mM KCl and adjust pH to 7.0 with NaOH. Bring the solution to 1 liter and sterilize by autoclaving. Just before use, add MgCl2 to 10 mM and glucose to 20 mM, from sterile stock solutions.

  10. Prepare 10, 100, and 1000-fold dilutions of the outgrowth in LB. Transfer 10 µl of each dilution to a test tube containing 3 ml of top agar + 200 µl of a mid-log culture of ER2738, equilibrated at 45°C. Vortex briefly and spread on LB/IPTG/Xgal plates. Incubate overnight at 37°C and count blue plaques the next day. 

  11. Scale up the protocol using the highest plaque/µg ratio to desired library complexity. For example, a library with a compexity of 1 x 109 clones would require a 5 µg ligation if the test ligations yield a ratio of 2 x 108 plaques/µg of vector. Use no more than 500 µl per individual ligation reaction; use multiple tubes if necessary.

  12. Purify the large-scale ligation by phenol/chloroform extraction, chloroform extraction and ethanol precipitation. Wash with 70% ethanol to desalt. Resuspend the DNA in low salt buffer and electroporate as described above. To reduce the likelihood of cells picking up more than one DNA sequence, the ligation should be divided and electroporated using as many cuvettes as convenient. For a 10–20 µg scale ligation we typically carry out 100 electroporations, using 3 µl of resuspended ligated DNA per 100 µl of electrocompetent cells. 

  13. Add 1 ml of SOC to each cuvette immediately after electroporation. For high-complexity libraries it may be convenient to pool the SOC outgrowths into groups of 5. Each outgrowth (or pool of 5) should be incubated for 30–45 minutes (no longer) before amplification. Titer several outgrowths or pools (as in Step 10) prior to amplification in order to obtain library complexity. 

  14. Amplify the electroporated cells by adding 20 ml of pooled SOC outgrowths to 1 liter of early-log cells (OD600 0.01–0.05) in LB medium. Incubate with vigorous aeration (250 rpm) at 37°C for 4.5 to 5 hours. Centrifuge at 5000 g for 20 minutes at 4°C. Transfer the supernatant to a clean bottle and discard the cells. 

  15. Recover the phage from the supernatant by adding 1/6 volume of 20% PEG/2.5 M NaCl and incubating overnight at 4°C. Pellet the phage by centrifugation at 5000 g for 20 minutes at 4°C. Discard the supernatant.

  16. Thoroughly resuspend the phage pellet in 100 ml of TBS by gently rocking over ~1–3 hour or overnight at 4°C. Remove residual cells by centrifugation at 5000 g for 10 minutes at 4°C.

  17. Transfer the supernatant to a new tube and discard the pellet. Reprecipitate the phage by adding 1/6 volume of 20% PEG/2.5 M NaCl and incubating for 1 hour at 4°C. Centrifuge at 5000 g for 20 minutes and discard the supernatant.

  18. Resuspend the final library in 10–40 ml of TBS by gentle rocking for 24–48 hours at 4°C. For long-term storage, add an equal volume of sterile glycerol, mix thoroughly and store at -20°C. The titer of the library should remain constant for several years at this temperature. Further amplification of the library is not recommended, as sequence biases may occur upon reamplification.