CRISPR screened drop-out screens provide researchers with a simple, efficient, and inexpensive method to interrogate chain function on the genome-wide level. The advantage of CRISPR is its pliability to edit any gene by simply changing the guide sequence. CRISPR guide libraries enable researchers to interrogate the entire genome of any organism in one experiment in an unbiased, systematic way.
Currently, CRISPR screens are being used to identify essential genes across hundreds of human cancers and to map genetic interactions. Screens can also comprehensively profile drugs to reveal drug mechanisms of action. The CRISPR libraries described here target human cells.
However, guide libraries targeting other species, such as mouse, are available and can be screened similarly. Performing genome-wide screens in human cells can be daunting in practice, as it involves the handling of tens of millions in cells and requires analysis of large sets of data. Before starting a screen, ensure cell lines are carefully characterized.
This includes knowing the purity of your cell line, doubling time, lentivirus transduction efficiencies, and sensitivities to antibiotic selection agents. A visual demonstration will provide a picture of how to practically handle the millions of cells required in a screen and keep track of thousands of perturbations in a systematic manner. Demonstrating the procedure will be Andrea Habsid, Kamaldeep Aulakh, and Ryan Climie, all technicians from my laboratory.
Begin by transforming the ready-made CRISPR sgRNA plasmid into electrocompetent cells and growing them according to manuscript directions. When ready to harvest the colonies, add seven milliliters of LB with carbenicillin to each plate and scrape the colonies off with the cell spreader. Use a 10-milliliter pipette to transfer the scraped cells into a sterile one-liter conical flask, and rinse the plate with five milliliters of LB with carbenicillin.
Again, transfer the rinsing solution to the flask. Centrifuge the cells according to manuscript directions, and then, determine the weight of the wet pellet. Purify the plasmid DNA using a Maxi or Mega scale plasmid purification kit.
Prepare cells for transfection by seeding 293 T-cells and incubating them overnight. On the next day, prepare the three transfection plasmid mixture for 15-centimeter plates. Calculate the amount of plasmid needed for one transfection and make a mix of plasmids for the number of plates, plus one, to be transfected.
Next, prepare a lipid-based transfection reagent for each transfection and aliquot-reduced serum media into individual 1.5-milliliter microcentrifuge tubes for the number of plates to be transfected. Add the transfection reagent, mix gently, and incubate at room temperature for five minutes. After the incubation, add DNA to the transfection reagent at a three to one ratio of transfection reagent to micrograms of DNA complex.
Mix the solution gently and leave at room temperature for 30 minutes. Add the transfection mixture to packaging cells and incubate them according to manuscript directions. On the day of viral harvest, check the cells for abnormal and fused morphology as an indication of good virus production, and harvest the lentivirus by collecting the supernatant and transferring it to a sterile centrifuge tube.
Begin by selecting the CRISPR guide RNA library coverage to be maintained throughout the screen. Based on the library coverage, determine the number of cells required to maintain it per guide RNA and the number of cells required for infection at MOI 0.3. Next, determine the number of plates required to set up the infection and then, harvest and seed the cells to each plate.
Add hexadimethrine bromide to all plates and add the required volume of virus to screening and control two plates. Do not add virus to control one. Replace that volume with media.
Mix the plates thoroughly by tilting and place the plates into an incubator, making sure that they are level. Harvest infected cells according to manuscript directions and collect three replicates of cell pellets from the pooled cells for genomic DNA extraction. Centrifuge the cells at 500 times g for five minutes and wash them with PBS.
Label the tubes and freeze dry the cell pellets at minus 80 degrees Celsius. Split the pool of infected cells to three replicate groups, making sure to maintain library coverage within each replicate. Seed the cells at a density that would normally be used when expanding them.
Use the same number of cells for each replicate and the same total number of cells between replicates. Continue to passage the cells and harvest three replicates of cell pellets from each replicate of pooled infected cells for up to 15 to 20 cell doublings. At each passage, harvest the cells from all plates in each replicate with each other.
Label each pellet with the time and replicate designation. Set up PCR1 according to manuscript directions with a total of 100 micrograms of genomic DNA at 3.5 micrograms of genomic DNA per 50-microliter reaction and set up identical 50-microliter reactions to achieve desired coverage. Set up one PCR tube reaction according to manuscript directions.
Use five microliters of the pooled PCR1 product as a template and use unique index primer combinations for each individual sample to allow pooling of sequencing library samples. After completing the PCR, run the PCR tube product on a 2%agarose gel at low voltage for one to 1 1/2 hours. Visualize the product on a blue light transilluminator and excise the 200 base pair band.
Purify the DNA and measure its quantity and quality with both a spectrophotometer and a fluorometer. An ideal guide RNA library should have every single guide RNA represented at similar quantities. Next generation sequencing can be used to confirm that the library has a tight distribution of guide RNAs.
Precision-recall analysis can be used to evaluate screen performance. A high-performing screen should recover a large number of essential genes at a base factor larger than six and a false discovery rate of less than 5%The precision-recall curve should have a sharp elbow and a straight line to the terminal point. The Bayes factor represents a confidence measure that the gene knockout results in a fitness defect.
High scores indicate increased confidence, while low scores suggest that the gene knockout provides growth advantages. Genome-wide knockout pools can be cultured in the presence of excess drug agent to look for suppressor resistance genes. To perform a positive selection screen for suppressor of thymidine block, normalized read counts for all guide RNAs at T0 are plotted against mean-normalized read counts for thymidine-treated samples.
A key detail to successfully performing CRISPR screens is maintaining good distribution of each guide RNA, starting from transfection of the plasmid to transduction of cells. This will minimize the chance of random effects that can skew guide RNA representation and lead to false positive or negative results. Following a screen validation, experiments should be done to confirm hits because the primary screen only identifies potential hits.
Depending on the biology of the hits, various methods can be used. CRISPR editing, combined with next-generation sequencing, has enabled genome-scale loss of function screens in diverse human model systems. Due to the simplicity of CRISPR, these types of screens are now broadly accessible to all researchers, allowing more scientists to utilize functional genetic methods to study biological processes and disease.
