Phage Display Library Prep Method

Sabrina A Mann, Sara Vazquez, Caleigh Mandel-Brehm, Lillian Khan, Joseph Derisi

Published: 2022-03-15 DOI: 10.17504/protocols.io.rm7vz3945gx1/v1

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Abstract

This protocol was designed to be used after the Scaled High Throughput Vacuum PhIP Protocol or the Scaled Moderate Throughput Multichannel PhIP Protocol. The immunoprecipitated phage targets are prepared for sequencing through a two PCR amplification rounds which first amplify the peptide target and second, add on UMIs.

Before start

Steps

Lysis

Dilute phage lysis product in water at a 1:4 ratio. Phage lysis should be used either immediately after IP or within 1 week of being stored at 4°C.

NOTE: we have done several tests and noticed that diluting the phage lysate 1:4 in water results in the most product at the end. Other dilutions ranging from 1:2 to 1:16 also worked. Using undiluted lysate also works, however there is the possibility that too much input can inhibit the PCR.

Heat lysate in preparation for PCR at 70°C for 0h 15m 0s (can be done on the PCR machine if aliquoted into PCR tubes or plates). Keep on ice afterwards.

PCR 1- Amplifying Insert

Add the following reagents. Reaction volumes of 50µL, 25µL or 12.5µL all work.
Multiply the number of samples by either the 50µL, 25µL or 12.5µL reaction coefficients to get the total volumes of each reagent to add to the master mix.
Make master mix and aliquot into a new PCR plate.
Add pre-heated phage lysis to PCR I master mix.

A	B	C	D
COMPONENT	50uL REACTION	25uL REACTION	12.5uL REACTION
Nuclease-free water	34.5	16.25	8.125
5X Phusion HF or GC Buffer	10	5	2.5
10 mM dNTPs	1	0.5	0.25
10 uM Forward Primer	1	0.5	0.25
10 uM Reverse Primer	1	0.5	0.25
Polymerase	0.5	0.25	0.125
Template DNA (lysis)	2	2	1
Total amount of Master Mix	48	23	11.5

Program PCR Machine for the following steps:

A	B	C
STEP	TEMP	TIME
Initial Denaturation	98°C	30 seconds
13-16 Cycles	98°C	5 seconds
	70°C	20 seconds
	72°C	15 seconds
Final Extension	72°C	2 minutes
Hold	10°C

PCR 2- Adding on UMI

Multiply number of samples by either the 50µL, 25µL or 12.5µL reaction number coefficients to get total volume of each reagent to add to master mix.
Combine the following reagents (except barcode primers and PCR I product) to make PCR II master mix, then aliquot master mix volume into new PCR plate
Next, add barcode primers and PCR I product, making sure to keep track of which barcode IDs corresponds to which sample well.

A	B	C	D
COMPONENT	50uL REACTION	25uL REACTION	12.5uL REACTION
Nuclease-free water	33.5	16.25	8.125
5X Phusion HF or GC Buffer	10	5	2.5
10 mM dNTPs	1	0.5	0.25
Polymerase	0.5	0.25	0.125
5 uM Forward and Reverse Primer Barcodes	4	2	1
Template DNA (product of PCR I)	1	1	0.5
Total amount of Master Mix	46	22	11

Program the PCR machine for the following steps:

A	B	C
STEP	TEMP	TIME
Initial Denaturation	98°C	30 seconds
5 Cycles	98°C	5 seconds
	70°C	20 seconds
	72°C	15 seconds
Final Extension	72°C	2 minutes
Hold	10°C

Pool and Bead Clean

Amplified samples are now uniquely labeled and are very similar in concentration and library size, so equal volume of each sample can be pooled together. If there is concern that samples are dissimilar, skip this step, bead clean, BioAnalyze and Qubit each sample individually. However, in our experience it is easier to pool first and then bead clean everything together prior to the QC check via BioAnalyzer and Qubit.

If pooling, it is recommend to pool anywhere between 5µL-10µL of each sample together by using a multichannel and combining all of the rows into a reservoir.

NOTE: It is also recommended to pool each plate separately from other plates. This allows us to protect individual plates from phage contamination.

10.

*Allow beads to sit in 70Room temperature for 0h 30m 0s prior

*Ratio of beads changes based on application and library size.

Determine the total volume of the pool. (i.e. 100uL)
Use Agencourt AmPure XP Beads 0.9x ratio of beads-to-total volume of sample. Prepare 80% EtOH.
Add 0.9x (i.e. 90uL) beads of room temperature AmPure Beads to pool. Mix well by pipetting up and down gently.
Pulse spin the tube but do not spin down beads. Incubate for 0h 5m 0s at 70Room temperature.
Place samples on magnetic rack and incubate for 0h 5m 0s on the rack.
Remove supernatant without disturbing the beads.
Add 2x original volume of 80% EtOH or enough to submerge bead pellet while on the magnetic rack. Incubate at 70Room temperature for 0h 0m 30s then remove the supernatant.
Repeat EtOH wash (step 7) for a total of 2 times.
Air dry the beads for 0h 5m 0s while on the magnetic rack.
Remove tube from magnetic rack. Elute DNA from beads in desired volume of 0.1x TE Buffer, 10mM Tris-HCl, or Nuclease free water plus 3µL of dead volume.
Vortex to mix. Pulse spin tubes and incubate for 0h 2m 0s at 70Room temperature off the magnetic rack.
Place on magnetic rack until solution is clear and bead pellet has formed ~ 0h 5m 0s.
Remove desired volume of eluant and transfer to a clean nuclease-free PCR tube. Do not disturbe the bead pellet.

Quantify and Qualify Library

11.

Qubit the samples for accurate concentration. BioAnalyzer also gives the concentration of sample but in practice this number not as accurate as the Qubit result.The Qubit is our gold standard for confirming DNA concentration for dilution prior to using the BioAnalyzer as to not overload it.
BioAnalyze or TapeStation library pools.
If the library size looks as expected, pool separate plate pools together at equal concentration or at desired ratio for read depth. Final libraries can be stored at 4°C for a few weeks. For longterm storage, samples should be kept at -20°C.

Expected result of library on a BioAnalyzer, it is normal to see these two peaks, the first peak should be smaller than the second. Make sure to confirm that the size of the main peak is the predicted size based on insert and primer scheme.