TITLE: DeRisi Lab Phage Immunoprecipitation Sequencing (PhIP-Seq)
AUTHORS: Hannah Kortbawi, Caleigh Mandell-Brehm, Brian O'Donovan, Saravazquez, Madhuraraghavan, Elze Rackaityte, Grace Wang, Aditi Saxena, David Yu, Joseph Derisi

[DESCRIPTION]
This protocol describes the process of Phage Immunoprecipitation Sequencing (PhIP-Seq) as it is performed in the DeRisi lab. It is the basis of many published papers from our group and is used for the discovery of protein targets of antibodies. The technique uses T7 bacteriophage that display a library of tiled peptide fragments that cover a human or microbial proteome. Human sera or monoclonal antibody controls are incubated with a phage display library and antibodies are immunoprecipitated using protein A/G beads. The purified phage library is then amplified in E. coli and re-incubated with sera, typically using three rounds of enrichment. Enriched phage DNA is then sequenced to identify antibody reactivity. In this way, a proteomic assay (immunoprecipitation) is converted into a genomic assay, leveraging the significant advantages of next-generation sequencing counting statistics.


[BEFORE_START]
Design and set up plates
Determine how many plates you will be doing based on number of samples, relevant healthy controls, GFAP antibody control (1:10 in SB), A/G bead-only control wells, and canary (no-bead, no-sample) control wells. Because individuals possess unique auto-reactive antibodies, we recommend using numerous (≥50) healthy control samples to account for non-disease-related enrichment - see Vazquez et al. 2022.
Design plate layout: for multiple-plate and multiple-batch runs, have a few duplicate healthy controls across plates and batches to help control for batch effect. Every plate should have A/G bead-only wells (no serum), and every batch of plates should have multiple canary wells. For patient cohorts with multiple disease statuses, disease status/outcome should be randomized within plates. For an example of a quality control validation plate layout, see attached spreadsheet.
Create working plates: aliquot serum into 96-well PCR plates (Bio-Rad Cat# HSP9601) and dilute 1:1 in storage buffer (SB). Serum should be arrayed in the plate layout planned for PhIP-Seq assay (put diluted GFAP in appropriate well, and leave A/G bead and canary wells empty). 5 uL of serum in 5 uL SB is a good amount in case assay needs to be repeated.
Store plates flat at 4°C until needed.

Make sure you have enough supplies and reagents
(See Materials tab)

[STEPS]
SECTION: Before Starting
1. Make buffers
SECTION: Day 1
4. Prepare overnight E. coli (BLT5403) stock
*This should be prepared from frozen for each day
SECTION: Day 1
4.1. Add 20-25 mL LB-carb into an autoclaved 125 mL Erlenmeyer flask
*Flame all flask and bottle openings upon opening and closing containers
SECTION: Day 1
4.2. Using a sterile loop to scrape some E. coli from a frozen BLT5403 E. coli stock
*It is critical to start all overnight cultures from the glycerol stock
SECTION: Day 1
4.3. Incubate in 37°C shaking incubator overnight.
It can be useful to start two overnight cultures in case one fails
SECTION: Day 1
5. Incubate peptidome with serum
SECTION: Day 1
5.2. Discard BSA from one of the blocked plates by upside-down flick over the sink.
Bring the plate back to the bench and blot the top dry using a clean paper towel. Do not wipe, only blot.
SECTION: Day 1
5.3. Pour appropriate volume of peptidome into a sterile reagent reservoir (50mL per plate).
Use multichannel P1000 to add 500 uL peptidome to each well of the plate using filter tips.
SECTION: Day 1
5.4. Change gloves
SECTION: Day 1
5.5. Add 1 uL of 1:1 serum in storage buffer (SB) to each well containing peptidome following the pre-defined plate map.
Use LTS tips and dispense serum directly into phage solution. Do not pipette up and down to mix.
To avoid contamination, when removing tips, bring pipette straight up from wells, move to the right so that the tips do not hover over any other rows, and eject tips into a bench height trash can.
DO NOT eject tips into benchtop waste container. This can aerosolize droplets from the tips or otherwise create splash that can contaminate other wells of the plate. Ejecting tips below the benchtop minimizes exposure.
SECTION: Day 1
5.6. Seal plates very well with rubber seals. A mechanical plate sealer can be used to improve ease. Visually inspect each plate to ensure complete sealing.

Rubber seals can be autoclaved on wet cycle after use for re-use.
SECTION: Day 1
5.7. Incubate plates on their side on the rocker at 4°C overnight. Use tape or other means to keep plates securely positioned on their sides.
 

SECTION: Days 2-4: IP and Washes (rounds 1-3)
6. Prepare A/G beads
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.2. Resuspend protein A and protein G beads by gentle swirling and end-over-end mixing. Avoid introducing bubbles.
*NEVER vortex because this can damage the beads.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.3. Add A and G in a 1:1 mix to a 15 mL or 50 mL conical tube.
12.5 uL of each bead type is needed per well. Make extra to account for pipetting error: 1 plate = 1.3 mL each bead
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.4. Place tube of A/G bead mixture on block magnet on ice. Any high magnetic field magnet will work - you can tape the tube to the magnet if you do not have a stand. 
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.1. Bleach work station and work over flame at all times.
The flame creates local updraft so no particles fall into wells. If a Bunsen burner is not available, use an alcohol burner.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.5. Using a new 2 mL serological pipette attached to vacuum manifold tubing, aspirate supernatant. Re-sheath the pipette in the plastic casing between washes to maintain sterility.
*A new tip should be used each day, but changing between washes isn't necessary.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.6. *Do not allow beads to dry out!
Immediately add cold TNP40 to beads at a volume enough to cover bead pellet and resuspend gently using serological pipette. Avoid adding bubbles.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.7. Repeat the previous two steps twice more for a total of 3 washes.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
6.8. Resuspend beads in cold TNP40 (volume proportional to IPs being done)
A good rule of thumb is 25 uL per well. Add extra TNP40 proportional to the amount of extra beads pipetted in 6.3: if 1.3 mL of each bead was used, resuspend washed beads in 2.6 mL TNP40.
SECTION: Before Starting
1.1. Make 3% BSA blocking solution in 1X TBS-Tween
1 mL of BSA is needed per well for blocking plates
Use spreadsheet for volume and recipe calculations
SECTION: Before Starting
1.2. Make TNP40
Very little TNP40 is needed per IP run - making 500 mL of TNP40 will last several runs of phage
Use spreadsheet for volume and recipe calculations
SECTION: Before Starting
1.3. Make RIPA buffer
2.5 mL of RIPA is used per well round of IP (5 washes x 500 uL RIPA per wash)
Use spreadsheet for volume and recipe calculations
Store RIPA at 4°C until ready to use
SECTION: Before Starting
1.4. Make 5M NaCl
78 uL of 5M NaCl is used per well per round of IP
Use spreadsheet for volume and recipe calculations
None. 
SECTION: Day 1
3. Re-spray work surface with 10% bleach and thoroughly wipe down before starting
For all following steps, work next to a lit Bunsen burner at all times
SECTION: Days 2-4: IP and Washes (rounds 1-3)
7. Add A/G Beads to peptidome+serum
SECTION: Days 2-4: IP and Washes (rounds 1-3)
7.1. Spin down plates at 1000 RCF for 1 minute at 4°C to pull down liquid from top of seals.
Visually inspect liquid levels in the plate from the side - if there are different heights in different wells, the plate was not correctly sealed and you should not proceed with the assay. 
SECTION: Days 2-4: IP and Washes (rounds 1-3)
7.2. Add 25 uL washed A/G beads to each well containing sample+library using a P200 multichannel pipette. Airdrop beads over the wells.
*Remember to leave A/G beads out of canary wells*
Mix beads in reservoir by gentle side-to-side rocking between each plate to avoid settling.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
7.3. Seal plates very well with rubber seals.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
7.4. Mix on rocker at 4°C for 1 hour.
*DO NOT EXCEED 1 hour of incubation - 50 minutes to 1 hour of bead binding is okay. Start washing first group of plates (if doing more than 2) at 50 minutes. Longer incubation may increase nonspecific binding.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
8. While the beads and samples are incubating, prepare new E. coli culture.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
8.1. Change gloves and bleach station
SECTION: Days 2-4: IP and Washes (rounds 1-3)
8.2. Aliquot enough LB-Carb for 550 uL per well into a flask and add 1 mL E. coli from overnight culture.
*Make sure to use a flask that can hold 5x the volume of culture used
*Flame all flask and bottle openings upon opening and closing
SECTION: Days 2-4: IP and Washes (rounds 1-3)
9. While the beads and samples are incubating, prepare wash station
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10. Wash phage-bound beads
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.1. Spin plate at 800-1000 RCF for 1 minute to pull down liquid from the top of the seal in each well
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.3. Using a multichannel set to 600 uL, remove all supernatant from each well.
For subsequent washes, set pipette to 500 uL.
Pipette from opposite side of well from the bead pellet to avoid touching beads
Discard supernatant into a waste bucket containing bleach in bench height garbage can. Eject tips into garbage can. DO NOT carry tips over other rows of the plate, as supernatant may drip into other wells.
 

SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.4. Open the lid of the RIPA reservoir and pick up 500 uL using a fresh set of tips. Carefully airdrop into each row, using the same tips for the entire plate (as long as tips do not touch any wells).
 

SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.2. Place plate on top of magnet on ice and allow the beads to pellet. Remove rubber seal very carefully, slowly peeling the seal back across the face of the plate, not upwards. This is to avoid creating a momentary vacuum in the wells, which could cause liquid to aerosolize. No liquid should be visible on the inside of the seal.
*If any liquid appears to fling up from the seal, IMMEDIATELY stop removing the seal, re-seal, and spin down again. The run may be contaminated at this point - consider re-starting. 
 
SECTION: Days 2-4: IP and Washes (rounds 1-3)
9.3. Set up wash station:
Place a 96-well plate magnet and an empty, clean P1000 tip box (tip holder insert removed) on ice in a rectangular ice bucket.
Place RIPA buffer on ice in a nearby ice bucket
Decant RIPA into empty P1000 tip box (used as reagent reservoir) and close the lid - do this right before bead incubation is over
 
SECTION: Days 2-4: IP and Washes (rounds 1-3)
9.2. Unwrap P1000 tip boxes - 7 boxes per plate
SECTION: Days 2-4: IP and Washes (rounds 1-3)
9.1. Change gloves and bleach station
SECTION: Days 2-4: IP and Washes (rounds 1-3)
9.4. Tape a large plastic bucket to the inside of a large, waist-height garbage can next to your bench.
Fill it with ~1 cm of 10% bleach.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.5. Carefully blot the top of the plate with a clean paper towel to remove any drips.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.6. Seal plate tightly with rubber seal and mark plate to keep track of wash number.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.7. Gently invert plate to resuspend beads, and then place plate on rocker for 5-10 minutes (10 min maximum) at 4°C.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.8. Repeat the RIPA wash for the next plate as the previous one is rocking.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
10.9. Repeat for a total of 5 washes per plate
Each addition of RIPA counts as a wash
Aim for no more than 2 hours for all 5 washes so the sample is not in RIPA for too long
When doing multiple plates, batch them in twos for easy centrifuging (e.g., six plates would be done in three pairs of two at a time)
SECTION: Days 2-4: IP and Washes (rounds 1-3)
11. Around the 4th incubation, check the OD600 of the E. coli culture - 0.3-0.6 is good.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12. Amplify enriched phage
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.1. After the 5th wash, remove all supernatant (set pipette to >500 uL), taking care to not touch the pellet, then remove plate from the magnet
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.2. Pour LB-carb into a sterile reagent reservoir (enough for 150 uL per well)
Immediately add 150 uL LB-carb to each well by airdrop (no need to change tips between rows as long as tips do not touch plate at all). Do not allow beads to dry out.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.3. Add 550 uL E. coli to each well of a square-well 2 mL deep-well plate.
There is no need to change tips between wells at this step.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.4. Use a multichannel P200 to mix each row of LB and beads, and then transfer to the plate with E. coli.
Be sure to get all beads transferred to the new plate - it is okay if the mixture gets frothy at this point, as the immunoprecipitated phage does not have to be bound to the beads and can just be in LB
Try to add the LB-carb/beads from each row to the E. coli around the same time so lysis occurs uniformly across the plate.
Start an off-plate canary, as well: aliquot 1 mL of E. coli culture into a glass test tube.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
11.1. *Change gloves and bleach bench or aliquot on a different bench entirely for this step!
When E. coli has grown to correct OD, decant it into a sterile 50 mL reagent reservoir or a clean, empty P200 tip box.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.5. Seal with a gas permeable seal and incubate in a 37°C shaker for 1.5-2 hours.
Place off-plate canary tube into shaker at the same time, too.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.6. Stop incubation when lysis occurs:
Try to catch the clarification as soon as possible
Place plate on 96-well magnet on the bench top (with flame on!) before checking for lysis to pellet beads to allow for easier observation of clarification vs. turbidity.
Compare lysis to on-plate canary well - the on-plate canary should remain turbid, indicating no lysis, while wells with patient samples should be cleared. This tells you that there has not been splashing between wells.
Compare lysis to off-plate canary - the off-plate canary should also remain turbid, which tells you that the original E. coli stock is not contaminated by phage.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.7. Add 78 uL of 5M NaCl to each well of the clarified culture (to a final molarity of 0.5M).
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.8. Spin plate at 4°C for 1 hour at 3220 RCF (or max speed) for 1 hour to pellet E. coli cell debris.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.9. While plate is spinning, empty out BSA from second round of plates as described previously.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.10. After spinning is complete, use a multichannel P200 to take out 50-100 uL lysate from the top of the wells, avoiding debris at the bottom and transfer this lysate to a 96-well PCR plate to be stored at 4°C. This is your Round 1 Lysate.
*We often only transfer and keep 50 uL of round 2 and round 3 lysate for sequencing. Keeping 50 uL of round 1 lysate is optional.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.11. Transfer 500 uL of lysate per well to new blocked deep-well plate. Again, slowly and carefully aspirate liquid from the top of the wells, avoiding any E. coli debris and beads.
If debris/beads are picked up, the plate can be spun again for 15 min to re-pellet and lysate transfer can be attempted again.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.12. Add 1 uL of 1:1 serum:SB from working plate (should be exact same plate configuration as added in round 1) to the lysate in the Round 2 deep-well plate.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.17. Repeat steps 6-12 for two more rounds of IP for a total of 3 rounds
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.13. Store remaining lysate and bacterial pellet in the square well plate, sealed with a foil seal, at 4°C until library prep, sequencing, and QC have been completed.
SECTION: Day 5: Library Prep
13. Prepare lysate
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.14. Incubate serum+phage on 4°C rocker overnight.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.15. Change gloves and bleach station.
SECTION: Days 2-4: IP and Washes (rounds 1-3)
12.16. Grow a new overnight stock of E. coli (from frozen) for the next day, as described in step 4.
SECTION: Day 1
5.1. Change gloves and re-clean your work surface with bleach.
SECTION: Day 5: Library Prep
13.2. Dilute phage lysate 1:4 in sterile, nuclease-free water (10 uL in 30 uL) in PCR plates.
SECTION: Day 5: Library Prep
13.3. Heat lysate at 70°C for 15 minutes.
While lysate is heating, prepare the PCR I master mix (see step 14).
Proceed immediately to PCR I after heating lysate. Do not let lysate sit for more than a few hours -  DNA will degrade and PCR will fail.
SECTION: Day 5: Library Prep
14. PCR I
SECTION: Day 5: Library Prep
14.1. Use the spreadsheet to scale the PCR mix for the number of samples being prepped. Make at least 5-10% more master mix than the number of samples you have.
PCR reaction volumes can be 12.5, 25, or 50 uL. All work and 12.5 uL is plenty for library prep.
SECTION: Day 5: Library Prep
14.2. Prepare the PCR I master mix.
Change gloves after pipetting phage lysate.
Add water, 5X Phusion buffer, dNTPs, and Strep and Flag primers first.
Add polymerase from -20°C when you are ready to aliquot the master mix.
Make 10uM STREP and FLAG primers from 100uM stock fresh for each run.
SECTION: Day 5: Library Prep
14.3. Aliquot master mix into new PCR plate
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13.1. Bleach station and work over flame until PCR II is complete
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14.4. Add volume of heated phage lysate proportional to volume of PCR reaction (see spreadsheet) to PCR I master mix in PCR plate, seal with foil seal, vortex, and thermal cycle with the following conditions:

 
While PCR I is running, prepare the PCR II mix (see step 15)
SECTION: Day 5: Library Prep
15. PCR II
SECTION: Day 5: Library Prep
15.1. Use the spreadsheet to scale the PCR mix for the number of samples being prepped. Make at least 5-10% more master mix than the number of samples you have.
PCR reaction volumes can be 12.5, 25, or 50 uL. All work and 12.5 uL is plenty for library prep.
SECTION: Day 5: Library Prep
15.2. Prepare the PCR II master mix.
Add water, 5X Phusion buffer, and dNTPs first.
Add polymerase from -20°C when you are ready to aliquot the master mix.
SECTION: Day 5: Library Prep
15.3. Aliquot master mix into new PCR plate.
SECTION: Day 5: Library Prep
15.5. Add volume of PCR I proportional to the volume of the PCR reaction (see spreadsheet) to PCR II master mix in PCR plate, seal with foil seal, vortex, and thermal cycle with the following conditions:

 

SECTION: Day 5: Library Prep
15.4. Aliquot TruSeq 12bp dual index primers into appropriate wells using the volume proportional to the volume of the PCR reaction (see spreadsheet).
*Carefully record which barcode IDs go into each plate. Do not re-use barcode plates across multiple plates destined for the same sequencing run.
*To avoid cross-contamination, seal barcode plates after use and do not reuse.
SECTION: Day 5: Library Prep
17. Bead clean
SECTION: Day 5: Library Prep
17.1. Allow beads to sit at room temperature for 30 minutes prior to use.
SECTION: Day 5: Library Prep
17.2. Calculate the total volume of your pooled samples (e.g., 100 uL).
Use AMPure XP beads at a 0.9x ration of beads to total volume of sample. E.g., if the sample is 100 uL, the appropriate bead volume is 90 uL.
SECTION: Day 5: Library Prep
16. Pool samples
SECTION: Day 5: Library Prep
16.1. Mix PCR II samples using a P20 multichannel and transfer 10-12.5 uL of PCR II (using the same tips) to a 50 mL reagent reservoir.
SECTION: Day 5: Library Prep
16.2. Mix well in reagent reservoir by gentle side-to-side agitation and then transfer pooled samples to clean DNA LoBind Eppendorf tubes. (One plate will fit into a 5 mL tube with enough room for bead cleaning volumes, but not into a 1.5 mL tube.)
SECTION: Day 5: Library Prep
17.4. Gently resuspend beads by swirling and end-over-end mixing, but avoid generating bubbles.
Add 0.9x of room temperature beads to DNA and mix well. Do not vortex.
SECTION: Day 5: Library Prep
17.3. Prepare fresh 80% EtOH (do not use stored diluted EtOH).
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17.5. Incubate sample at RT for 5 minutes.
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17.6. Place samples on magnetic rack and incubate for another 5 minutes (beads should be completely pelleted).
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17.7. Carefully remove supernatant without disturbing the bead pellet.
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17.8. While keeping the sample on the magnetic rack, add enough 80% EtOH to submerge the pellet of each sample.
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17.9. Incubate for 30 seconds, then remove supernatant.
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17.10. Repeat EtOH wash for a total of 2 washes.
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17.11. Place tube on magnetic rack and then sharply rap rack on the bench top to release remaining drop of EtOH from the pellet and then remove supernatant again.
SECTION: Day 5: Library Prep
17.12. Air-dry beads on magnetic rack until no EtOH is left (~ 5 min)
SECTION: Day 5: Library Prep
17.13. Remove tube from the magnetic rack. Elute DNA from beads in desired volume of 0.1x TE buffer, 10mM Tris-HCl, or nuclease-free water plus 3 uL of dead volume.
*A good rule of thumb is 50 uL of elution buffer per plate (plus 3 uL)
SECTION: Day 5: Library Prep
17.14. Vortex to mix beads and elution buffer.
SECTION: Day 5: Library Prep
17.15. Incubate for 2 minutes at room temperature off the magnetic rack.
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17.16. Place on magnetic rack and pellet ~5 minutes, until solution is clear.
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17.17. Remove desired volume of eluent and transfer to a clean nuclease-free DNA LoBind tube.
*Label the lid of this tube very clearly
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18. Qubit library
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18.1. Dilute samples 1:10
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18.2. Prepare Qubit working solution (dilute HS Reagent 1:200 in HS buffer, scaled to the number of samples + 2 extra reactions for standards)
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18.3. Aliquot 190 uL Qubit working solution into 2 tubes (one for each standard) and 199 uL working solution into tubes for each cleaned library.
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18.4. Add 10 uL standard to appropriate tubes and 1 uL 1:10 diluted library to appropriate tubes.
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18.5. Vortex samples thoroughly for 5 seconds each.
*Qubit will not correctly estimate concentration if samples are not vortexed.
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19. Bioanalyzer
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19.2. Allow HS DNA dye concentrate (blue) and HS DNA gel matrix (red) to equilibrate to RT for 30 min.
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19.3. Add 15 uL HS DNA dye concentrate (blue) to HS DNA gel matrix (red). Vortex and spin down.
SECTION: Day 5: Library Prep
19.1. Notes
Allow all reagents and samples to equilibrate to room temperature 30 minutes before use.
Protect dye and dye mixtures from light. Remove light covers only when pipetting.
Always insert the pipette tip to the bottom of the well when dispensing the liquid.
Use loaded chips within 5 minutes after preparation
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19.4. Transfer to a spin filter and centrifuge at 2240 g ± 20% for 15 min. Protect solution from light (wrap in foil). Store at 4°C. Use prepared gel-dye mix within 6 weeks of preparation.
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19.5. Allow the gel-dye mix to equilibrate to room temperature for 30 min before use.
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19.6. Put a new High Sensitivity DNA chip on the chip priming station.
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19.7. Pipette 9 μL of gel-dye mix in the well marked Dark G.
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19.8. Make sure that the plunger is positioned at 1 mL and then close the chip priming station.
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19.9. Press plunger until it is held by the clip.
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19.10. Wait for exactly 60 s then release clip.
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19.11. Wait for 5 s, then slowly pull back the plunger to the 1 mL position.
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19.12. Open the chip priming station and pipette 9μL of gel-dye mix in the wells marked Light G.
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19.13. Pipette 5 μL of marker (green) in all sample and ladder wells. Do not leave any wells empty.
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19.14. Pipette 1 μL of High Sensitivity DNA ladder (yellow) in the well marked Ladder.
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19.15. In each of the 11 sample wells pipette 1 μL of sample (used wells) or 1 μL of marker (unused wells).
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19.16. Put the chip horizontally in the adapter and vortex for 1 min at the indicated setting (2400 rpm).
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19.17. Run the chip in the Agilent 2100 Bioanalyzer instrument within 5 min.
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19.18. Results from Bioanalyzer will indicate the percentage between 300-700 basepairs and average basepair. Human peptidome library prep should show a peak around 349 bp.
1.5. Prepare work space.
Spray work surface with 1:10 dilution of bleach and thoroughly wipe down before starting
2. Block plates.
Plates are blocked with BSA to reduce non-specific binding of antibodies to phage.
Block enough plates for 3x the number of plates of samples you are running.
BSA should be made fresh for every PhIP-Seq experiment.
2.1. Decant 3% BSA solution into a sterile reagent reservoir
Add 1 mL BSA per well into a round-bottom 2mL 96-well deep well plate [see materials] using a multichannel
*Tips do not need to be changed between rows or plates at this step
2.2. Seal plates thoroughly with foil seals and incubate on rocker at 4°C overnight or for 4-6 hours at room temp.

SECTION: Days 2-4: IP and Washes (rounds 1-3)
8.3. Incubate E. coli culture in 37 degree incubator with rotational shaking.
SECTION: Day 5: Library Prep
20. Sequence using a sequencer with a maximum reads per run that will allow for 2 million reads per sample.