Genome-wide CRISPR Screens in T Helper Cells Reveal Pervasive Crosstalk between Activation and Differentiation

Summary T helper type 2 (Th2) cells are important regulators of mammalian adaptive immunity and have relevance for infection, autoimmunity, and tumor immunology. Using a newly developed, genome-wide retroviral CRISPR knockout (KO) library, combined with RNA-seq, ATAC-seq, and ChIP-seq, we have dissected the regulatory circuitry governing activation and differentiation of these cells. Our experiments distinguish cell activation versus differentiation in a quantitative framework. We demonstrate that these two processes are tightly coupled and are jointly controlled by many transcription factors, metabolic genes, and cytokine/receptor pairs. There are only a small number of genes regulating differentiation without any role in activation. By combining biochemical and genetic data, we provide an atlas for Th2 differentiation, validating known regulators and identifying factors, such as Pparg and Bhlhe40, as part of the core regulatory network governing Th2 helper cell fates.


Manual for CRISPR KO screening in primary mouse T helper cells
These instructions are similar but not exactly those we use in the paper "Genome-wide CRISPR screens in T helper cells reveal pervasive cross-talk between activation and differentiation". Several steps can be improved further; in particular, higher virus titres can be achieved using caffeine (https://www.ncbi.nlm.nih.gov/pubmed/20626321). There are mixed reports of the ability to freeze mouse-specific retrovirus, which would simplify the protocol execution greatly. Concentration by ultracentrifugation has not worked well for us, but overnight spins at lower g might work better. This protocol does not have a freezing step and need careful logistical planning. The final sequencing of the CRISPR library is the same as for Kosuke Yusas mouse v.2 KO library, from which our library is derived (https://www.addgene.org/pooled-library/yusa-crispr-knockout-mouse-v2/).

Maintenance of 293T cells for virus production
We use 293T for virus production normally, but it is possible to also use a producer cell line. Even in this case it is worth supplementing the with pCL-ECO, the envelope plasmid. Do not under any circumstance let the cells go overconfluent as this reduces the proliferation rate. Subsequent virus production/transfection is severely impaired by this. Avoid running too many passages (<20-30?).

Splitting: Expect 2x growth per day
If you want to put 293T cells onto gelatin coated plates then first take 10cm dishes and add 4ml 0.1% gelatin. Incubate them at 37°C for at least 30 min. Then just remove the gelatin-containing media before adding the T cells.
To split: 1. Remove media from plate.
4. Put plate into incubator for 2 minutes (or 2.5). Do not stack the plates -if you do then reheating will take longer, and possibly be uneven, affecting the trypsinization.
5. Quick method: Flush off the cells with any amount of media (e.g. 5ml). Pipette up and down to resuspend single cells. Slow method: Flush of the cells with any media, centrifuge 450g 5min. Remove media, add 1ml, resuspend by pipetting.
6. Dilute to the amount of media needed. Add geneticin unless you are making virus the day after (50mg/µl is 100x but can be used as 500x or 1000x). Geneticin is optional but supposed to maintain and select for the cassette with the SV40 large T antigen. This may boost virus production. We frequently omit this step.

Thawing
Heat media first. Then heat cells in water bath (fast heating is good . It is key that the plates have the right amount of dryness, or the media will either be sucked in, or the streaks will mix. Do up to 3 plates/8 channels to ensure you have enough colonies (2 plates is good). Notation: Write the letter of the column (before streaking) on the side you will streak from. Streak from left to right, with (1) on the top. Move to the incubator with the streaks still turned up.

Validation of sgRNA plasmid insert
 The final validation is best done by Sanger sequencing, using primer mathU6seq: CGATACAAGGCTGTTAGAG (thanks to Mathias Friedrich).


A quick first check can be done by digestion and running the product on a gel. Digest by NEB EcoRI in buffer 2. Two fragments (8400+630) will appear if the original insert is left, otherwise just linear band (~8400). Do not do overnight digest. I have noticed that the entire plasmid can get digested (at least with this restriction enzyme). This may be due to remaining exonucleases if you miniprepped the plasmid. This is a master mix for 500ng/µl plasmid, 20µl/reaction and 20 reactions:

400µl Total
Mix 19µl + 1µl plasmid. Digest for 1h and run the product on a gel.

Virus production with lipofectamine LTX
The production also works with LT2000, but then instead use Kosuke Yusas protocol that only uses optiMEM during transfection. The most crucial parameter is confluence. If the cells ever hit the limit and becomes retarded then discard them and restart. There is no saving them from there. Lipofectamine can vary 35 -45µl without major change in efficiency, and plasmid 12-15µg with ease.
Grow 293T in Advanced dmem/f12 with FBS and P/S. Maintain on 1-10µg/ml geneticin (recommendation from Sebastian Łukasiak who uses 10µg; we prefer 1µg). Be sure to remove geneticin the day before transfection or the titre will be low. Aim for 80% confluence on the day of transfection.
On the day of transfection:

Virus production with PEI
This much cheaper methods is suitable for targeted KO but not for library making as the titre is 50% of LTX. Concentration of MSCV does not seem possible or at least not easy, but has not been tested enough.
This method, like calcium-phosphate transfection, seems based to creating cell-piercing crystals with DNA. Getting the crystal size right is crucial. Calcium-phosphate appears to give comparable results to LTX in the right hands, but PEI is much easier and the buffer lasts forever once made.
2. For 10cm plates: Change media to 9ml media on day of transfection.
3. In a sterile tube dilute total plasmid DNA (µg) in 1m, optiMEM (volume of media is 10% of final volume in culture vessel). Total plasmid (pCL-eco + MSCV, 50/50%) does not seem to be a major effector. 8-12µg total is a good number. 3. Drop the pH to at least 4.0 (Important! Not written in any other protocol; thanks to Laura Wood for pointing this out). Neutralize to pH 7.0, filter sterilize (0.22µm), aliquot and store at -20°C; a working stock can be kept at 4°C.

Estimation of required number of cells
The number of required cells is easiest to estimate by computer simulation. While we have not validated the results, this may still provide a back-of-an-envelope estimate of the required amount. There are several bottlenecks through-out the protocol and it makes sense to try and balance these: • Many cells die during activation and will thus restrict the effective number of available cells • Any skew in the complexity of the library will propagate. We assumed the library to be uniform

• The number of infected cells is a key parameter
• The sequencing depth is another key parameter • The expected fold-change of the KO that you wish to detect • The number of sgRNAs expected to work (unknown) and the number of sgRNAs per gene For discussion, we talk about the rate at which viruses enters cells, MOI (λ), and the probability of a cell to be seen as infected (c). The relationship is, assuming a poissonian distribution, λ =−log( 1−c) . It is generally easier to talk about MOI than infected cells. A higher MOI (λ) means more cells to measure, but also more cells with more than one sgRNA inserted. The ideal tradeoff depends on your scored phenotype. Commonly MOI=0.3 is used. The following R code shows the probability of an observed transduced cell being a single transduction: lambdas <-(0:150)/100 prob_0 <-exp(-lambdas) prob_1 <-lambdas*exp(-lambdas) prob_2plus <-1 -prob_0 -prob_1 #Not 0 and not 1 plot(lambdas,prob_1) plot(lambdas,prob_1/(prob_1+prob_2plus),xlab="MOI", ylab="P[Observed cell is single-transduced]",type="l") To get an idea of the number of required cells, we chose to look at the expected number of false positives. It is difficult to reason about how sgRNAs are merged together as a per-gene score (indeed, we came up with a whole Bayesian framework in our paper to calculate this). Thus we only look at the scores for one sgRNA. How many sgRNAs will by chance be reported to have a >2x fold change? The following code calculates this using the EAT model  runif(1e5,min=0,max=88000)),stage=1) numdiv <-round(5*24/14) ## Number of divisions in 5 days numdiv <-8

General notes on methods for T cell extraction
We have compared Stem Cell Technologies, Miltenyi and a home-brew negative T cell selection protocol. Generally the home-brew protocol worked but found that screening is already sufficiently complicated to not involve additional steps. SCT has given more varying purity results than Miltenyi, possibly due to the extraction being at room temperature. You also obtain fewer T cells for the same purity level with SCT double-negative selection as opposed to Miltenyi negative-positive selection. However, the Miltenyi protocol does not scale well to the number of cells required for screening, and takes more time to execute even for small numbers of mice. Nowadays we use Miltenyi only for individual KO, if ever. Both Miltenyi and SCT recommend against using red blood cell lysis because the debris may clog cells in subsequent steps. However, we find that with proper care, especially the filter step afterwards to remove lipids, this need not be a problem. With the much reduced number of input cells, a smaller amount of antibodies must be used, or the negative selection will be too harsh.

CRISPR screening & KO on mouse T cells
This assumes that you get T cells from mice expressing Cas9. Avoid mice over 12w as the number of T-cells are reduced quite a bit, but these will also work. We use mainly 8-10w. Expect ~7M cells per spleen with the following protocol.
Plan to use 20-25 mice per screening, if 100% of the cells express the gene of interest. If fewer, scale up accordingly or perform a rescreening. Assuming an MOI of 20-30%, for 30M cells output cells, this means 100M-150M cells are needed on a bare minimum.

Day 0 or before -making of plates
Use 6w plates. Coat with 1µl/ml CD3 with 1.2ml per well. Put in 37°C for at least 2 hours or overnight 4°C (and 3µl/ml CD28 if using the old protocol with both of these plate bound, see later remark)

Day 0 -extraction of T-cells
Idea: Use cold PBS in the beginning of the protocol to keep cells happy. Then use room-temp PBS when at SCT stage. Simple solution is take it out of fridge and just keep it on the bench, and let it warm up as you perform the protocol.
1. Take a 10cm plate and add up to 4-6 sets of 70µm sieves. In each sieve add 3 spleens. More spleens risk loss of T cells due to clogging. Add 10ml cold PBS in total over the spleens. Use the back of a syringe to macerate the spleens. I believe more spleens/sieve will reduce the viability of the cells. Unconfirmed rumors also has it that the back of a syringe is bad and that the front is better, but this is much slower in my hands. I use the hard side of the syringe to first cut the spleen in pieces, then the rubber side to macerate.
2. Add as much PBS as needed to wash off the cells and pass through a 70µm into a 50ml falcon. Reusing the 10cm plate might help in retaining cells.
3. Spin down 400g/5min and resuspend 3 spleens per 50ml falcon. Up to 4 spleens can be processed in only falcon, but cell loss may be higher.
4. Perform RBC lysis: vortex cells to resuspend. add 5ml 1x RBC lysis buffer (eBioscience, use and predilute the 10x as it is much cheaper than the 1x). Wait 4 min, then fill with cold PBS to 50ml. Spin down and remove supernatant. b. CRUCIAL: do not put more than 6 spleens into one large magnet. That is, you need to dilute it more than the protocol says. For 20-25 spleens, use all 4 magnets. If you lose the cells in the magnet then resuspend the cells on the walls of the magnet in PBS and use the magnet again. Loss is lower with 15ml polypropylene tubes than with falcon tubes.

Day 0 -Plating T cells
Resuspend cells into IMDM with 1ul/ml BME, 1ul/ml CD28 and any IL* of choice. If using only IL-2 then 5ul/ml. It is possible that adding IL-2 might boost differentiation here but we don't do it anymore. Keeping the CD28 in the media might help budding cells which later will not have direct contact with plate surface, so it makes it more even.
For a 6w plate, aim for 40M cells per plate (ready next day). Do not by any means go below 20M. 30M can be a compromise. This is a step we think deserves further optimization as other groups have reported higher infection efficiency with lower plating density.

Day 1 -Transduction
If cells are floating, remove media, spin them down and add them in new media without stirring the cells already attached to the plate too much. I use 6ml media/plate. Note that this is pushing the limit for the amount of media/well for 6w, and you will see some cells detach. The problem seems particularly bad the more plates you stack. I resuspend the cells lightly after centrifugation to make up for this Put plates into suitable centrifuge preheated to 32°C. Having it at 32°C keeps the cells happy while the virus is degraded slower (this has been published before, and is called "cold infection", but we have not verified it to be better). Several 6/96 well plates can normally be stack in a centrifuge. However, if too many plates are stacked, we noted that the angle is off. This can be a problem if you centrifuge with a small amount of virus media.
Spin for 2h at 1000-1100g. Put the plates in zip-lock bags or too much liquid will evaporate. It might be beneficial to work with 12 well-plates here to also reduce use of virus but we have in practice gotten stuck on using 6 well-plates out of habit. Put the plates for up to 16h in 32°C incubation. One paper says that "extended centrifugation does not improve efficiency" but we have not tested the influence of this. It is possible to get infected cells even if you skip the incubation after the spinning, but this is not recommended.
Resuspend cells in IMDM with differentiating cytokine of choice, BME 1µg/ml and 1µg/ml CD28. Keep cells in at least 2ml/well, possibly 3ml.

Day 2 -Optionally transduct again
Redo infection if you wish. Approximately doubles the efficiency according to others. We have not verified this.

Day 2 -Puromycin selection
This is to be done on the day after transduction. 24h after spinning, change the media of the cells (IL*, BME, CD28) but now in addition 2µg/ml puromycin. For each well, use at least 3ml media as the cells are depleting it very fast.
We have estimated kill rates and think anywhere 2-3µg/ml is suitable. We once did a quick experiment and got the following kill rates: With 2.5µg/ml puromycin, 38% of the cells were transduced on day 5. With 5µg/ml, 53% of the cells were transduced.

Day 3 -Media change
It is likely that another media change is needed here too, same composition as the day before. But so many cells will already have died that the media change need will be much reduced from here on