A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site—in essence, analyzing onsite mutagenesis.

Transcript

The overall goal of this experimental procedure is to outline a foundational approach in detailed methodology to measure genetic heterogeneity at the target site in a reliable and robust fashion. This method can help answer key questions in the gene editing field such as, how can onsite mutagenesis be analyzed and measured. The main advantage of this technique is that, it's a standardized methodology for homology directed repair or gene correction using single stranded oligonucleotides.

To begin the protocol, grow HTT11619 cells in 25 milliliters of previously prepared medium for the culture of HTT116 cells in a T-175 flask. Aspirate the medium and wash the cells with dulbecco's phosphate buffered saline or PBS, without calcium and magnesium, then aspirate the PBS. Next, add trypsin drop-wise to the T-175 flask, using a five milliliter pipette.

Allow the cells to detach in an incubator. Tap the flask to dislodge the cells and then quench the cells with the eight milliliters of complete medium over the entire surface of the flask. Then pipette the mixture up and down multiple times to break up cell clumps and transfer the cells to a 15 milliliter conical tube.

Before spinning the cells down, take 10 microliters from the 15 milliliter conical tube and combine it with 10 microliters of trypan blue to count the cells, then pellet the cells. Transfer 10 microliters of the cells mixed with trypan blue to the hemocytometer and count the four grids around the outside. For each 10 centimeter plate of cells to be synchronized, add five milliliters of complete medium and six micromolar of aphidicolin.

Next, transfer 100 microliters of the re-suspended cell pellet to each centimeter plate and swirl the plate gently to mix. Incubate the plates to synchronize the cells at the G1S border. Four hours prior to targeting, aspirate the medium, wash with PBS.

Aspirate the PBS and add five milliliters of complete medium. Place the plate back in the incubator for four hours. Enter the mutant EGFP gene sequence into the Zane Labs online generator and choose the CRISPR guide sequences that bind with close proximity to the target site.

Mix the RNA in equal molar concentrations to 45 micromolar. Add 6.75 microliters of a 200 micromolar stock of CR RNA and 6.75 microliters of a 200 micromolar stock of tracer RNA to a 1.5 milliliters centrifuge tube. Then, add 16.50 microliters of IDT buffer to make a final volume of 30 microliters.

Next, heat the mixture at 95 degree celsius for five minutes, in a PCR machine. Allow the samples to cool to room temperature. For each sample, dilute 2.22 microliters of CR RNA to tracer RNA complex and 2.78 microliters of IDT buffer to a final volume of five microliters.

Dilute 1.67 microliters of Cas9 protein from a 60 micromolar stock in 3.33 microliters of low sera medium, to a final volume of five microliters. Mix five microliters of Cas9 protein with five microliters of complex RNA. Aspirate the medium, wash with five milliliters of PBS.

Aspirate the PBS and add one milliliter of pre-warmed trypsin to each 10 centimeter plate. Put the plates in the incubator. Next, tap on the 10 centimeter plate to make sure all the cells are dislodged and then quench them with four milliliters of complete medium, by dispersing it over the entire surface of the plate.

Pipette up and down multiple times to break up cell clumps and transfer the cells to a 15 milliliter conical tube. Set aside 10 microliters from the 15 milliliter conical tube and combine it with 10 microliters of trypan blue to count the cells. Pellet the cells by spinning for five minutes at 125 x G at room temperature.

Aspirate the medium and wash with five milliliters of PBS. Pelt the cells by spinning out 125 x G for five minutes at room temperature. Transfer 100 microliters of the cell suspension to each electroporation four millimeter gap cuvette.

Add 10 microliters of the RNP complex to 100 microliters of cells, at a 5 x 10 to the fifth cell density. Add two micromolar ODN to each sample. Next, take the rack to an electroporation machine.

Lightly flick each of the samples and place them in the chamber. Place the rack back in the hood and transfer each sample to a well containing two milliliters of complete medium in a six well plate. Incubate the plate before checking for correction levels.

Aspirate the medium and wash the cells with two milliliters of PBS. Replace the PBS with 500 microliters of pre-warmed trypsin to each well of the six well plate. Place the plates in the incubator.

Next, tap the plate to make sure all cells are dislodged and then quench with one milliliter of complete medium by dispersing it over the entire surface of the well. Pass the cells into a 1.5 milliliter centrifuge tube and pellet. After pelleting the cells, aspirate the medium, then re-suspend the cell pellet in 500 microliters of FACS buffer.

Electroporate the synchronized and released HTT11619 cells at a concentration of 5 x 10 to the fifth cells per 100 microliters, with the RNP complex at 100 picomoles and 100 picomole of 72 NTODN at 2.0 micromolar. Transfer the cells to a six well plate and allow them to recover for 72 hours. Next, sort the cells individually into 96 well plates, using a fac sorter with a 488 nanometer laser for EGFP plus minus.

From the wells that have growth, isolate cellular gDNA using a commercially available DNA isolation kit. Amplify the regions surrounding the target base via PCR. Finally, perform DNA sequencing analysis on the samples.

72 hours after incubation flow cytometry analysis, confirm the functional repair of green fluorescent protein. A gradual dose response was observed as the coordinated levels of RNP and 72NT increase. A gene editing reaction in the absence of the RNP particle, corrected approximately 1%of the targeted cells when a 10 fold higher concentration of the 72NT oligonucleotide is used in the single agent gene editing reaction.

16 clones of the EGFP positive samples were selected and the genetic integrity surrounding the target site was analyzed by DNA sequencer. All 16 EGFP positive cells contained the predicted nucleotide exchange at the target site. 15 non-green clonal isolates were analyzed for heterogeneity at the target site.

No DNA base exchange was observed in approximately half the samples. The remainder of the clonal expansions examined, displayed a heterogeneous population of deletion mutations. The deletion size ranged from one base to 19 bases.

CRISPR, Cas9 and single stranded oligonucleotide donor DNA molecules working in tandem can lead to the precise repair of the point mutation in the EGFP gene as demonstrated in this new model for the repair of point mutations, a molecular pathway, in which the donor DNA access replication template for the repair of the mutant base. A process we have termed, ExACT. After its development, this technique paved the way for researchers in the field of gene editing to explore the degree of heterogeneity at the target site, as a result of gene editing activity in cell lines.

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Journal (Genetics)

Journal (Genetics)

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

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