After you have determined the cleavage efficiency of the pooled cell population, isolate single cell clones for further validation and banking. You can isolate single cell clones from the selected pool using limiting dilution cloning (LDC) in 96‑well plates or by single cell sorting using a flow cytometer.

Limiting dilution cloning (LDC)

  • Based on the editing efficiency and estimated cell viability, you can estimate the number of single clones needed to obtain a desired knock-out (KO) clonal cell line.

    For example, if you desire a homozygous KO with mutations in both copies of a gene and the resulting GeneArt cleavage detection efficiency was 50%, then the probability of having both alleles knocked out in any cell is 25% (0.5 × 0.5 = 0.25).

    If the probability of an indel leading to frame shift is 2/3, then the chance of having a homozygous KO is ~11% per cell [(0.5 × 0.5) × (0.66 × 0.66) = 0.11].
  • We recommend performing limiting dilution by targeting 0.8 cells/well, which requires you to resuspend the transfected cells (post-counting) at a density of 8 cells/mL in complete growth medium, then transferring 100 μL of this to each well of a 96‑well plate.

    If you plate at least ten 96-well plates in this manner and expect only 20% of cells to survive, then the probability of having homozygous KO clones in the 192 surviving cells will be 19–21 cells (192 × 11%).
  • Note that single cell clone survivability varies by cell type. Some cells that do not like to remain as single cells need to be plated at a low density to get well separated colonies, which will then have to be manually picked for further screening.
     

Example LDC procedure using 293FT cells

  1. Wash the transfected cells in each well of the 24-well plate with 500 μL of PBS. Carefully aspirate the PBS and discard.
  2. Add 500 μL of TrypLE cell dissociation reagent to the cells and incubate for 2–5 minutes at 37°C.
  3. Add 500 μL of complete growth medium to the cells to neutralize the dissociation reagent. Pipette the cells up and down several times to break up the cell aggregates. Make sure that the cells are well separated and are not clumped together.
  4. Centrifuge the cells at 300 × g for 5 minutes to pellet.
  5. Aspirate the supernatant, resuspend the cells in an appropriate volume of pre-warmed (37°C) growth medium, then perform a cell count.
  6. After counting, dilute the cells to a density of 8 cells/mL of complete growth medium. Prepare a total of 50 mL of cell suspension at this cell density and transfer to a sterile reservoir.
    Note: You can also perform a serial dilution to get a better estimate of cell density.
  7. Using a multichannel pipettor, transfer 100 μL of the cell suspension into each well of 96‑well tissue culture plates until the desired number of plates is seeded. Make sure to mix the cells in between seeding the plates to avoid the formation of cell aggregates.
    Note: In general, we seed ten 96-well plates to achieve a large number of clones. Number of plates to seed depends on the editing efficiency of pooled cell population and viability of cells post single cell isolation.
  8. Incubate the plates in a 37°C, 5% CO2 incubator.
  9. Scan the plates for single cell colonies as soon as small aggregates of cells are visible under a 4X microscope (usually after first week, depending on the growth rate of the cell line).
  10. Continue incubating the plates for an additional 2–3 weeks to expand the clonal populations for further analysis and characterization.
     

Example single cell sorting procedure in a 96-well plate using flow cytometer

You can sort single cells per well into a 96-well plate format using a flow cytometer with single cell sorting capability. After sorting and expanding the single cell clones, analyze and characterize the clonal populations using suitable assays. The following is an example single-cell sorting procedure with 293FT cells.

  1. Wash the transfected 293FT cells in each well of the 24-well plate with 500 μL of PBS. Carefully aspirate the PBS and discard.
  2. Add 500 μL of TrypLE cell dissociation reagent to the cells and incubate for 2–5 minutes at 37°C.
  3. Add 500 μL of complete growth medium to the cells to neutralize the dissociation reagent. Pipette the cells up and down several times to break up the cell aggregates. Make sure that the cells are well separated and are not clumped together.
  4. Centrifuge the cells at 300 × g for 5 minutes to pellet.
  5. Aspirate the supernatant, then wash the cell pellet once with 500 μL of PBS.
  6. Resuspend 1 × 106 cells in 1 mL of FACS buffer, then add propidium iodide (PI) to the cells at a final concentration of 1 μg/mL. Keep the resuspended cells on ice.
  7. Filter the cells using suitable filters before analyzing them on a flow cytometer with single cell sorting capability.
  8. Sort PI-negative cells into a 96-well plate containing 100 μL of complete growth medium. If desired, you can use 1X antibiotics with the complete growth medium.
  9. Incubate the plates in a 37°C, 5% CO2 incubator.
  10. Scan the plates for single cell colonies as soon as small aggregates of cells are visible under a 4X microscope. Colonies should be large enough to see as soon as 7–14 days (usually after first week, depending on the growth rate of the cell line). You can perform image analysis to ensure that the colonies are derived from single cells.
  11. After image analysis, continue incubating the plates for an additional 2–3 weeks to expand the clonal populations for further analysis and characterization.
     

Characterize edited clones

You can analyze the single cell clones for purity and the desired genotype (homozygous or heterozygous allele) by various molecular biology methods such as genotyping PCR, qPCR, next-generation sequencing, or western blotting.