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Ph.D.™-12 Phage Display Peptide Library Kit > FAQ |  | Ph.D.™-12 Phage Display Peptide Library Kit FAQ
See the Protein Tools FAQ also.
Q1: Where can I find many more detailed FAQs for Phage Display Peptide Libraries?
Q2: Can a different bacterial strain be used with the Ph.D.™ Phage Display?
Q3: No plaques are visible when titering using the Ph.D.™ Phage Display kit.
Q4: I am using Ph.D.™ Phage display and the amplified phage titer is low.
Q5: I am using Ph.D.™ Phage Display and the phage DNA templates do not yield readable sequence.
Q6: I am using Ph.D.™ Phage Display and the sequencing templates do not run where they should on a gel.
Q7: I am using Ph.D.™ Phage Display and after 4 or more rounds of panning all clones are wild-type phage (white plaques).
Q8: When performing an experiment using Ph.D.™ Phage Display, the ELISA indicates that background binding to the plate is as high as binding to the target.
Q9: When using the Ph.D.™ Phage Display, panning yielded a consensus sequence, but no ELISA signal.
Q10: I am using Ph.D.™ Phage Display and the streptavidin control experiment did not yield the HPQ consensus sequence.
Q1: Where can I find many more detailed FAQs for Phage Display Peptide Libraries?
A1: You may find the complete set of FAQs for Phage Display Peptide Libraries under "Technical Reference > Protein Tools" on our Web site.
Q2: Can a different bacterial strain be used with the Ph.D.™ Phage Display?
A2: In theory other F+ strains containing the supE suppressor mutation (such as XL1-Blue and DH5αF´) should work with our phage display system. However, we have not tested these strains with our libraries and do not know whether here will be any subtle effects on the expression or transport of certain peptides out of the cell. Since the Ph.D. libraries were made in ER2537, the immediate predecessor to ER2738, we know that all the peptides in the libraries can be successfully expressed in this strain. Therefore, we recommend this strain over any other one.
Q3: No plaques are visible when titering using the Ph.D.™ Phage Display kit.
A3: Unlike lambda, M13 is a non-lytic phage and does not produce clear plaques. M13 plaques are areas of diminished cell growth, not lysis, and consequently can be difficult to see. Try holding the plate up to a light. Also, since the vector used to prepare the library carries the lacZα gene, plaques will be blue, and easier to see, when using an alphacomplementing strain such as the supplied strain ER2738 and plating on Xgal/IPTG plates.
Also, be sure the dilution range is appropriate for the phage you are titering. For amplified phage, plate 10 µL of 1:109 - 1:1011 dilutions; for unamplified panning eluates, try 1:10 - 1:104 dilutions for early rounds, 1:104-1:107 for later rounds. If the phage is not sufficiently dilute, the plaques will be confluent on the plate and it will look like there are no plaques at all (or a bluish tinge when using Xgal plates). Occasionally after PEG precipitation, the phage will clump and not dilute properly. As a result, you might have a plate containing too many plaques merged together. Make sure to give the phage ample time to resuspend after precipitation (> 1 hour) and vortex each dilution tube very well (~10 seconds).
Q4: I am using Ph.D.™ Phage display and the amplified phage titer is low.
A4: In order for M13 phage to be efficiently amplified, it is critical that cultures be well aerated, and that cultures be infected early in their growth phase. We recommend amplification in 20 mL cultures in 250 mL Erlenmeyer flasks, in a shaker set to 250 rpm. Amplification in smaller vessels, such as 50 mL conical tubes, will result in much lower yields of amplified phage. M13 phage should either be added to an early-log culture, A600 < 0.01, or to a 1:100 dilution of an overnight culture. Yield of amplified phage is maximal after 4.5-5 hours at 37°C; longer incubation may result in deletions and is not recommended. If carrying out nonspecific elution with pH 2.2 glycine buffer, the eluted phage must be neutralized as described in the Manual prior to amplification.
Q5: I am using Ph.D.™ Phage Display and the phage DNA templates do not yield readable sequence.
A5: The sequencing template purification protocol in the manual should provide single-stranded template of sufficient purity for dideoxy sequencing with Sequenase™ (Amersham), or automated cycle sequencing with dyelabeled terminators (ABI). The procedure should be followed exactly as described in the manual: prolonged ethanol precipitation, precipitation at 20° C or centrifugation longer than 10 minutes will result in co-precipitation of salt and phage proteins, which will inhibit sequencing. Additionally, it is crucial that the phage pellet is thoroughly suspended in the iodide buffer prior to adding ethanol. If problems persist, or if another sequencing method is used, a phenol:chloroform extraction step can be added: Following suspension in Iodide Buffer, add 2 volumes of TE, extract once with phenol:chloroform (1:1) and once with chloroform, and ethanol precipitate. 5 µL of suspended template (approximately 0.5 µg) should be sufficient for sequencing; quantitation should be confirmed by agarose gel electrophoresis using 0.5 µg single stranded M13 DNA (NEB #N4040S) as a standard.
Q6: I am using Ph.D.™ Phage Display and the sequencing templates do not run where they should on a gel.
A6: The sequencing templates prepared by the method in the manual are single-stranded (approx. 7250 nucleotides), and as a result will not line up with double-stranded markers of the same length. The apparent size will vary depending on the applied voltage, ethidium and agarose concentration in the gel, and whether TBE or TAE is used as running buffer. We strongly recommend using single-stranded M13 DNA (e.g. single-stranded M13mp18, NEB #N4040) as a marker.
Q7: I am using Ph.D.™ Phage Display and after 4 or more rounds of panning all clones are wild-type phage (white plaques).
A7: In a typical round of biopanning, ~2 x 1011 input phage are reacted with the target, and between 103 and 107 total phage are eluted off following washing. This corresponds to an enrichment of 104 to 108-fold per round. Since the library contains ~2 x 109 different clones, the eluted pool of phage should in theory be fully enriched in favor of binding sequences after only 2 or 3 rounds. Once this point is reached, further rounds of amplification and panning will result only in selection of phage that have a growth advantage over the library phage. For example, vanishingly small levels of contaminating environmental wild-type phage (less than one part per billion) will completely overtake the pool if too many rounds of amplification are carried out, regardless of the strength of the in vitro selection.
Q8: When performing an experiment using Ph.D.™ Phage Display, the ELISA indicates that background binding to the plate is as high as binding to the target.
A8: If panning against a polystyrene plate coated with the target (direct coating method), it is possible to inadvertantly select peptides that specifically bind the polystyrene surface (see Adey, N. B. et al. (1995) Gene 156, 27-31). These peptides will yield identical ELISA signals in the presence and absence of target, since the ELISA plate is also made of polystyrene. Such "plastic binders" are typically rich in aromatic residues (Phe, Tyr, Trp, His), which often alternate (the sequence FHWTWYW is a plastic binder discovered and characterized at NEB). Selection of plastic binders often occurs in the absence of a strong target preference for peptide sequences present in the library: other libraries may yield the desired targetspecific sequences. Selection of polystyrene-specific peptides can be avoided by using the bead capture protocol described in the Manual. The phage is reacted with the target in solution, and the phage-target complexes are then captured onto beads that specifically bind the target (protein A-agarose for antibody targets, glutathione-agarose for GST fusions, etc.). Unbound phage is removed by extensively washing the beads in a microfuge tube. Unlike polystyrene, neither the beads (typically crosslinked agarose) nor the microfuge tube (polypropylene) are likely to select specific peptide sequences from the library, although the species conjugated to the beads (protein A, glutathione, etc.) might. To avoid selection of bead-specific ligands, we suggest either alternating rounds between different beads specific for the target (e.g. protein A beads for rounds 1 and 3, protein G beads for round 2 for antibody targets), or adding a subtractive panning step, beginning with round 2, in which the phage pool is first reacted with the beads alone (no target), the beads discarded, and the supernatant from this step reacted with the target.
Q9: When using the Ph.D.™ Phage Display, panning yielded a consensus sequence, but no ELISA signal.
A9: When characterizing phage clones by the ELISA protocol in the manual, it is difficult to add more than 1012 virions per 100 µL well. This corresponds to a phage concentration of only 16 nM. At this concentration, an unambiguously positive ELISA signal can only be observed if the binding affinity is in the micromolar range or better. The iterative nature of phage selection permits identification of ligands with a broad range of affinities, from subnanomolar to 1 millimolar, so lower affinity ligands will not show a positive ELISA signal. In this case it is necessary to increase the concentration of the selected ligand, either by synthesing a peptide corresponding to the selected sequence (be sure to include the spacer sequence GGGS at the C-terminus, and amidate the C-terminal carboxylate if possible), or by expressing the selected sequence as an N-terminal fusion to a smaller protein. Alternatively, a sandwich ELISA can be carried out in which the selected phage is immobilized and an excess of target applied in the liquid phase. This procedure requires an antibody against the target protein, or some other means of detecting bound target protein. Coat the wells overnight with anti-M13 antibody (no HRP), wash, and add serial dilutions of each phage clone (one clone per row). After 1 hour, wash away unbound phage and add an excess of target protein (0.1 - 1 µM) in TBST. Incubate 1-2 hours at RT, wash away unbound target, and detect bound target with an enzyme-linked antibody.
Q10: I am using Ph.D.™ Phage Display and the streptavidin control experiment did not yield the HPQ consensus sequence.
A10: If you used low pH glycine rather than biotin to elute your phage, you will likely not get an HPQ consensus sequence. Due to the relatively low affinity of the peptide-streptavidin interaction, nonspecific elution is incapable of selectively enriching for HPQ-containing peptides. HPQ-containing peptides can be competitively eluted using the natural ligand biotin. If you used biotin to elute and still did not get a consensus sequence, the most likely explanation is that you did not carry out sufficiently rigorous washes. When you wash, pour the wash buffer in the plate from a bottle (don't gently pipet it in) and swirl it for about 10 seconds each time. The number of phage that you elute after the first round of biopanning should be in the range of 103 - 107 (closer to 103 for an ELISA well and closer to 107 for larger wells). If you are eluting more phage, you are not washing well enough and as a result, not getting sufficient enrichment. It also may help to add 0.1 µg/ml streptavidin to the blocking buffer to complex any contaminating biotin in your BSA, which could otherwise complex the streptavidin on the plate during the blocking step.
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