Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

Resuspend 2×10^6 cells in electroporation solution, add 5 µg each of two CRISPR/Cas9 constructs and 0.5 µg GFP reporter plasmid, then electroporate at 250 volts for 5 milliseconds in a 2 mm cuvette. Immediately transfer to pre-warmed culture media and incubate at 30°C for 24–72 hours to allow plasmid expression.

Pass cultured cells through a sterile 50 micron filter and perform fluorescence-activated cell sorting (FACS) to isolate the top 3% of GFP-positive cells, which indicates successful plasmid uptake. Freeze half of the sorted bulk cells for future use and plate the other half for validation.

Plate sorted cells into a 96-well plate at a dilution of 30 cells per well in 100 µL culture media per well. Incubate clones at 37°C for 7–14 days depending on the cell line doubling time to allow single-cell derived colonies to expand.

Resuspend parental and bulk cell pellets in 50 µL DNA extraction solution and run in a thermocycler to extract genomic DNA. Measure the DNA concentration to quantify the extracted genomic material.

Assemble 20 µL PCR reactions containing 10 µL 2× PCR mix, 0.5 µL each of forward and reverse primers, and 50–100 ng genomic DNA, run for 35 cycles with 60°C annealing temperature. Resolve PCR products on a 2% agarose gel to identify deletion and non-deletion bands.

Transfer clone suspensions to a 96-well PCR plate, centrifuge at 400× g for 5 minutes, and resuspend cell pellets in 50 µL DNA extraction solution. Run PCR reactions to detect both non-deletion and deletion bands across all clones.

Resolve PCR products on a 2% agarose gel and identify clones with desired biallelic deletions. Confirm biallelic deletion by Sanger sequencing of PCR amplicons and perform RT-qPCR on isolated RNA to verify loss of target gene expression.