All-in-One CRISPR Genome Editing: A Method for Homology Directed Repair-Based Gene Knock-In in Cultured Cells Using CRISPR-Cas9 System

Begin by adding all-in-one CRISPR plasmid to immune cells present in a tube. This plasmid contains a Cas9 gene in combination with the sgRNA or single-guide RNA encoding sequence, complementary to the target sequence in the host genome.

Add a linearized donor plasmid containing a transgene sandwiched between homology arms or HAs - the DNA repeats necessary for homologous recombination. Perform electroporation, which uses electric current to inject the plasmid mix into the cell. Once inside, the processed sgRNA forms a complex with the expressed Cas9 nuclease.

The sgRNA binds the target site, allowing the Cas9 nuclease to identify the specific recognition sequence known as protospacer adjacent motif or PAM, and cleave the host DNA near it. This creates a blunt-ended double-strand break where the cellular machinery resects the DNA, generating 3' overhangs. The presence of HA flanking sites triggers the homology-directed repair mechanism where the cut DNA joins the HA sequence, forming a heteroduplex structure - the D-loop.

DNA synthesis continues, forming a strand complementary to the transgene sequence until it rejoins the original DNA strand. The top of the D-loop acts as a template to synthesize the missing part of the DNA, generating the double Holliday junctions - structures where four DNA strands interact. These junctions resolve and the gaps repair, facilitating target gene insertion.

Prepare a 24-well plate containing 500 microliters of complete growth medium without antibiotics in each well. Pre-warm the plate in humidified 37 degree 5% carbon dioxide incubator. Turn on the electroporation system. Input electroporation parameters for JURKAT or RAW cells.

Prepare cell-DNA mixture for electroporation. Collect JURKAT cells culture in 25-square-centimeter flask. Transfer cells into 50-mL centrifuge tube. Then, pellet the cells by centrifugation at 90 x g for eight minutes at room temperature.

As for the supernatants, for washing, resuspend the cells with 5-mL DPBS. Then, spin the cell suspension by centrifugation, using the same condition as the previous step. Remove the DPBS, and resuspend the cell pellet using 2 mL of DPBS. Use 10 microlitres of cell suspension, and mix with an equal volume of 0.2% trypan blue.

To estimate the cell count, load sample into this counting slide. Insert counting slide in the automated cell counter, and view the image of cells, then, get a cell count result. Calculate 2 million cells for 5 repetitions of electroporation, per knock-in experiment.

Transfer the required number of cells into 1.5-mL centrifuge tube. Pellet cells by centrifugation. To save time, electroporation mixture can be prepared during the centrifugation step. Add 2.5 microgram of each of CRISPR-Cas9 vector, 2.4 microgram of linearized targeting vector, and re-suspension Buffer R, to a total volume of 55 microliter, in a new 1.5-mL centrifuge tube.

Gently vortex, and briefly spin to mix well. Leave the electroporation mixture at room temperature before use. After spinning, aspirate the DPBS, then, centrifuge for an additional 30 seconds. Remove as much supernatant as possible. Re-suspend cells by adding the prepared electroporation mixture. Pipette up and down gently.

Electroporation: As for the cell electroporation mixture, use a 10-microliter nucleofection tip with pipette.

Important: Avoid air bubbles during pipetting, it will cause electroporation failure. Press "Start."

After electroporation, transfer the sample in one well of 24-well plates with prewarmed growth medium. Perform the same electroporation procedures as above, on the remaining 4 replicate samples. Gently rock the plate to assure even distribution of the cells. Incubate the cells in 37-degree incubator for 48 to 72 hours prior to FACS cell sorting.