I. Making the library peptide-fragment:
Always save at least 5uL from each step for future analysis and verification.
1uL negative control template (p102M linked to a neg. ctrl. peptide or no peptide, use a 1:10 dilution of a midi-prep)
1uL dNTP mix
1uL Library oligo (HPLC purified)
1uL Taq (Invitrogen)
5uL Taq Buffer (Invitrogen)
38.5uL MilliQ water
95oC 30 sec, 60oC 1 min, 70oC 1 min, 25 cycles
- Run the whole reaction out on a 1% gel, purified by GeneClean or Qiagen kit. Make sure everything used to run this gel is clean (washed with soap and water) and only run one library fragment per gel to avoid cross-contamination. Change razors in between slices and gels.
- Sometimes I see a second band that is about twice as big as the real fragment, I’m careful to avoid this band during the purification.
2uL dNTP mix
2uL Taq (Invitrogen)
10uL Taq Buffer
77uL MilliQ water
- Run multiple reactions (usually 3) to ensure you have enough fragment for an entire ligation reaction. Combine during the purification.
- Purify this PCR using a purification column, or you can run the fragment on a 1% gel and isolate by GeneClean or Qiagen.
- Sometimes I still see a second band in this PCR, avoid it during the purification.
When using GeneClean and Qiagen, I resuspend in 100uL final volume.
- Submit about 5uL of PCR#2 for sequencing. Use primers 781 (for just the peptide section) or 3173 and 3175 for the whole fragment. This is to check your oligo and to make sure your fragment has the BstXI sites on both sides. Randomized positions should show all four colors (A, T, G, or C) and each randomized nucleotide position.
30uL fragment (1/3 of one PCR reaction)
10uL NEB Buffer 3
100uL reaction. Digest more than one reaction tube if more insert is needed.
- Incubate at 55oC for 1 hour then at 37oc for 1-2 hours.
- Purify the digested fragment by running on a 1% gel, isolate by Gene Clean.
- I had problems getting one of my inserts to digest properly, for some unknown reason. To fix this, I increased the digestion time to 8 hours at 55oC.
Notes about GeneClean and gel isolation:
- I started isolating my fragment by gel isolation because I was getting some other non-specific bands during my PCR. Especially after the digest, these were hard to purify away from the real fragment.
- Always do the GeneClean isolation as quickly as possible, don’t leave the fragments in the gel piece, in the NaI buffer, or incubating with the glassmilk too long.
- Dissolve gel in 1-2 mL of NaI buffer, incubate about 20 min at 40-50oC.
- Add 25uL of glass milk. Incubate 10 minutes, rocking at RT.
- I split this into two microcentrifuge tubes to wash. Spin 1 min at 14k. Decant.
- Add 1mL of Wash Buffer, spin, decant. Dry the pellet completely.
- Resuspend in a total of 50uL water (combine tubes here), spin, remove into a new microcentrifuge tube and resuspend glassmilk in a second 50uL of water, for a total final volume of 100uL.
To quantify and verify your fragment:
- Run 5uL of PCR#1, PCR#2, and digested PCR#2 on a 3% gel (all fragments should be purified before running, changing the buffers can affect how the fragments run on a gel).
- Use a ladder with a 400 bp marker. PCR#1 should be slightly blurry and should be smaller than the 400 bp marker. PCR#2 should not be blurry and should be bigger than the 400 bp band. Digested PCR#2 should be smaller than the 400 bp band and should be the only band in the well (if you have more than one band, you should try the gel purification again).
- Use this gel to quantify how much fragment you have (ng/uL). This varies a lot but I usually get between 5ng/uL–20ng/uL. Using a ladder made for quantification helps.
Store at 4oC until the GFP DNA is ready for use. Don’t freeze. Use ASAP.
II. Making Baculovirus GFP (or ICAM) DNA
The preparation of this DNA makes or breaks the efficiency of your ligation.
- Make a stock of GFP or ICAM virus (interrupting the B2M side of the vector, your MHC-I on the other side) from 3-5 flasks of 3x107 cells per flask, infected with 150uL virus. Allow all cells to die (7-12 days) before harvesting. Filter the supes.
- Spin down 100mL of viral supernatants in the ultracentrifuge (100k x g) in the Ti50.2, I usually split into four 25mL Oakridge tubes. Balance carefully using a digital scale. Spin 2 hours at 4oC.
- Pour off the supernatant, wash carefully (don’t disrupt the pellet on the side of the tube) with 5mL TlowE buffer (10mM Tris pH 8, 0.1mM EDTA). Decant, carefully tap dry. Add 100ul of TlowE to each Oakridge tube. Turn the tube on its side (close the lid) so that the buffer covers the virus pellet. Resuspend O/N at 4oC without disrupting.
- Remove the buffer into a 1.5mL microcentrifuge tube (combine 2 oakridge tubes together).
- Rinse the Oakridge tubes with 200-300uL of TlowE. Sometimes I can still see a white precipitate stuck on the side of the oakridge tube. I try to remove this by gently pipetting up and down.
- Add SDS (25uL of 10% SDS) and pK (5 uL of 10 mg/mL), incubate 1 hour at 37oC. This incubation is lysing viral particles and releasing the viral genome. From this point on, you must be EXTREMELY gentle with the DNA.
Phenol Chloroform extraction:
Pipette the DNA as few times as possible. Use gloves to avoid DNAses.
- Add 500uL phenol to the DNA tube, gently rock for 1 minute to mix. Spin at 13k for 1 min.
- Using a large-bore pipet tip, carefully and slowly remove the upper aqueous layer into a new microcentrifuge tube (add to the new tube drop-wise).
- Gently add 500uL phenol/chloroform/isoamyl alcohol. Gently rock for 1 min to mix. Spin 1 min at 13k.
- Again using a large-bore pipet tip, carefully and slowly remove the upper aqueous layer, add dropwise to a new microcentrifuge tube. Add 500uL choloroform, gently rock to mix. Spin 1 min at 13k.
- For the final time, remove the aqueous layer into a new microcentrifuge tube (sterile). (DNA has been pipetted 3 times at this point).
- Add 1/10th volume of NaOAc (50uL), gently rock until you can’t see any viscous layer (~1-2minutes).
- Add 2x volume of 100% EtOH, 1mL, (make sure this is a new bottle that hasn’t been pipetted into, trying to avoid any DNAse). Gently rock again until you can’t see any viscous layer (1-2 minutes).
- Place on a slow/gentle rocker at room temp and incubate 5-10 minutes. Before spinning, make sure the tube has been mixed well, you should be able to see a white cloud of DNA and should not be able to see any viscous layer. Spin at 13k for 5 min.
- Decant and add 1mL 70% EtOH (fresh), spin 1 min at 13k. Decant and add 1mL 100% EtOH (fresh). Spin 1 min at 13k.
- Decant and tap gently on a kimwipe, allow to air dry upside down (avoid using the hood air-flow to dry, it can suck your pellet right out of the tube).
- Add 100uL sterile-filtered TE (10mM Tris pH 8, 1mM EDTA). Incubate for 1-2 hours at 37oC. If you can spare the time, its best to let this resuspend by neglect O/N at 4oC after the 37oC incubation.
- Perform a control transfection to ensure your DNA prep is good before continuing with the digest and ligation. Transfect 2x106 SF9s in a 6-well dish with 500uL Transfection buffer A, 500uL transfection Buffer B and a 1ug-50ng titration of BV DNA. At day 3, harvest cells and run flow (FL1 for GFP, or stain with an anti-ICAM antibody). A good DNA prep will still have some ICAM/GFP positive cells at 50ng, and should have well over 50% positive with 1ug of DNA.
Baculovirus DNA digestion:
- Heat DNA to 55oC for 10 minutes (to ease pipetting). Take out 4uL for an OD, continue incubating at 55oC while you OD the sample. In general my DNA preps are between 750ug/mL-2mg/mL. Everything should be kept “sterile”.
- Using a large-bore tip, pipet 12.5ug of BV DNA into a 1.5mL microtube (prepare 6 tubes).
- To each tube add:
15uL CeuI (5k U/mL)
15uL SceI (5k U/mL)
water (up to 300uL)
- Use 1/10th of everything in a 0.5mL microtube (prepare 3 tubes with 1.5ug BV DNA).
- 3.0 NEB4
- 3uL diluted BSA
- Add 1.5uL enzymes to 2/3 tubes (one control tube should not have any enzymes added).
- The controls are uncut BV DNA (no enzyme added), cut BV DNA (no ligase added later), and cut re-ligated (add ligase to this tube later).
Gently rock to mix.
Incubate for 3 hours at 37oC, 20 minutes at 65oC to inactivate.
Precipitate to buffer exhange: (this is where my DNA was lost most of the time)
Add 30uL 3M NaOAc, pH 5.2, mix gently
600uL 100% EtOH, mix gently
- Gently rock until there is no visible viscous layer.
- Incubate at least 5 minutes rocking gently at room temperature.
- Spin 5min at 13k. Decant, add 1mL 70% EtOH, spin, decant, add 1mL 100% EtOH. Decant, tap on kimwipe and allow it to air-dry.
- Add the necessary volume of TE buffer (usually between 30-70uL, but it depends on the concentration of your peptide fragment. You will be adding 100ng of fragment.)
- Incubate to resuspend for 2 hours at 37oC and O/N at 4oC… extremely important!
III. Ligation and Transfection:
Heat to 55oC for 10 minutes.
Add 100ng of fragment.
20uL 5x ligation buffer (invitrogen)
5uL T4 DNA Ligase (2 million U/mL)
- Incubate 1-2 hours at room temp
- Add 10uL of 10mM ATP
- Incubate O/N at 19oC.
- Take transfection buffers out of the fridge to bring them up to RT.
- Add 3x107 SF9s to 3 large culture flasks.
- While the cells are settling onto the plastic, incubate the ligation reaction at 65oC for 10 minutes to inactivate the ligase. Heat at 55oC until ready to pipette.
- Once settled, carefully wash the cells 1x with Grace Plain (15mL). Remove Grace Plain and add 10mL of Transfection buffer A (5% serum for our current lot of serum).
- Add 1mL of transfection buffer B to the ligation reaction, gently rock to mix.
- Pipet 8mL of transfection buffer B into 3 15mL conical tubes.
- Using a large-bore pipet tip, add 2 tubes of the diluted ligation reaction to 1 15mL conical tube (bringing the total volume of each conical tube to 10mL). Gently rock to mix.
- Carefully and slowly add the transfection buffer B/ligation reaction to the cells, dropwise while rocking. Use a large-bore pipet tip to remove any extra buffer from the conical tube.
- Gently rock to mix and incubate for 4 hours at 27oC.
- Remove transfection buffer, wash 1x with Grace Plain, and add 30mL Graces Supplemented (10% serum). Incubate for 3 days.
IV. Sorting and Screening
Estimation of the Library size
- On Day 3, remove a small amount of cells for FACs analysis (I usually take 1mL from each flask).
- Stain the cells with an MHC antibody (FL2) and ICAM (if necessary). Secondary antibody controls are important for proper gating and calculations.
- I run at least 100,000 events (for a nice picture), although the actual number of events needed depends on how many MHC-expressing cells there are.
- Calculate the estimated size of your library by multiplying the percent of MHC+ cells by the number of cells infected. Add up the three flasks for your total estimated library size. This calculation assumes that one virus has infected one cell, and that each peptide is only represented once. For complete saturation of the library, 10-fold excess is recommended.
Sorting the Library
- Titer your virus prior to beginning. Infected Hi5’s (2x104 per mL) in 96 well plates, 100uL per well, with a titration of virus (a 103 dilution through 109). Most virus stocks are in the range of 108U/mL.
- Sign up for sort time with Shirley. She needs to know which colors you will be using and approximately how long you think the sort will take. For one flask of cells (3x107) it usually takes 1.5 hours.
- Infect 3x107 SF9s in a large flask with 1U/cell of virus (Day 0). For large libraries I infect 2 flasks of SF9s, just to make sure I’m representing all of the peptides in the library.
- Make sure everything is sterile and be extremely careful when handling the cells, contamination is a common problem.
- On Day 3 post-infection, spin down the cells, wash 2x with FACs buffer (no azide). Resuspend in 500ul of sterile filtered TCR-AD0304-AF647 stain (1:10 dilution of stock, dilute in FACs no azide). For two flasks of cells, I resuspend in 1mL of stain. Incubate at room temperature, in the dark (or wrapped in foil), for 1 hour.
- Add 500uL of MHC-I antibody (sterile filtered, diluted in FACs no azide, for 28.14.8s the dilution is 1:800). Again, for two flasks of cells I add 1mL of MHC-I stain. Incubate at 4oC for 1 hour.
- Wash 1x with FACs (no azide), and resuspend in 500ul of MHC-I secondary antibody stain (sterile filtered, for IgG2a-PE the dilution is 1:800). Resuspend in 1mL of stain for two flasks of cells. Incubate at 4oC for 30 minutes.
- Wash 2x with FACs (no azide). Resuspend in 2-3mL FACs, no azide. Transfer into the appropriate polypropylene tube (blue lid). Dilute 50uL of these cells into 500uL FACs no azide for a set-up tube (also in a polypropylene tube). Put 2mL of complete media into a third polypropylene tube (for collecting cells).
- After the sort, put all cells recovered into a flask with 3x107 SF9s and Grace’s Supplemented Media.
- The next round of sorting can be done as soon as Day 3. Repeat the staining process as above. Alternatively a viral stock can be grown up at each stage of the sorting