The goal of this procedure is to isolate and identify clonal knockout cell lines generated by two types of CRIPSR/Cas9-mediated genome editing. One method detects protein knockout caused by nonhomologous end joining while the other detects genomic perturbations generated by homology-directed repair. This method is particularly useful in the field of molecular biology, where generation of cell lines with genetic perturbations is an important tool in dissecting gene function.
The main advantage of this technique is that it is a quick and relatively high-throughput method that can be easily executed using basic molecular biology techniques. Nonhomologous end joining introduces small deletions or insertions, and are assessed by a dot immunoblot, while homology-directed repair allows for larger and more precise perturbations, and are assessed by colony PCR. This experiment uses T-REx-293 cells cultured in DMEM media, supplemented with 10%FBS at 37 degrees Celsius, and 5%carbon dioxide.
Prior to transvection, plate the cells onto six well plates and include a well for an untransvected control. Grow the cells to approximately 70%confluency. Transvect the cells with 2.5 micrograms of total plasmid using a commercial transvection protocol.
For transvection of cells without selection, use 2.5 micrograms of Cas9-sgRNA plasmid. For transvection of cells with selection, use 0.75 micrograms of Cas9-sgRNA1, 0.75 micrograms of Cas9-sgRNA2, and one microgram of homologous recombination template. Incubate the transvected cells and the untransvected control at 37 degrees Celsius and 5%carbon dioxide for 48 hours.
For cells transvected with selection, begin the drug selection by treating the cells with the appropriate drug, which is Neomycin in this case. Return the six-well plate to the incubator. Observe the treated cells once a day to check for rates of cell death, until all cells in the untransvected control die.
To isolate clonal populations, first grow the cells to 100%confluency in the original well after selection. Next make serial dilutions of the cells and count the cells. Since the proper generation of monoclonal colonies is key to obtaining a full knockout, care must be taken to determine the correct dilution for seating cells at the desired density.
Once the correct dilution has been determined, seat the cells into 96-well plates at a density of 0.33 cells per well. Seating three 96=well plates is a good starting point to ensure isolation of more than one correct clone. Over a two-to-four week period, observe the colonies until colonies are visible to the eye.
These representative images show colonies at different stages of growth. A single cell at seating, cells at day five, and a well-established colony at day 10. Using a sterile pipette tip, pick visible colonies with care and precision, and reseat in new wells to encourage monolayer growth.
Continue growing the cells at 37 degrees Celsius and 5%carbon dioxide. Knockout candidates without selection are screened by dot immunoblot to assess the protein product in a high-throughput manner. After growing individual clones to 50 to 100%confluency, dislodge the monolayer of cells by pipetting within the well.
Aliquot 90 microliters of the 100 microliter total volume from each well to a clean microcentrifuge tube. For the purposes of this video, only two samples will be processed. Spin down at 6, 000 RPM for five minutes, remove the media, and lyse the cell pallet in 10 microliters of 1X SDS loading buffer.
Add 90 microliters of new media to the remainder of the cells in the well, and return the plate to the incubator to continue propagating the cells. Pipette one microliter of cell lysate onto a dry nitrocellulose membrane to form a dot. Blot each sample twice on two separate membranes, creating two identical patterns of samples.
Block the membranes in 5%milk in TBST at room temperature, with rocking, for one hour. After one hour, blot the membranes with primary antibodies at the recommended primary antibody dilution, and TBST plus 5%milk. On one membrane use the primary antibody against the target protein, ELAVL1 in this experiment.
On the other membrane, use the primary antibody for a control protein that is not expected to change, PUM2 is used here. Incubate at room temperature for one hour. Remove the primary antibody solution from each blot, add TBST, and incubate for five minutes.
In this manner, wash the blots three times with TBST. Next, add to each membrane the appropriate HRP conjugated secondary antibody, and blot at room temperature for one hour. After one hour, wash the membranes three times with TBST for five minutes each time.
Apply a chemiluminescent substrate solution to the blots, following the manufacturer's instructions, and image the blotted membranes on a digital chemiluminescence imager. Lastly, quantify dot intensities for the target and control proteins using the appropriate software, and analyze the results as described in the text protocol. Candidates with the selectable resistance marker are screened by colony PCR.
Begin this procedure by duplicating individual colonies in a new 96-well plate, and growing them to 100%confluency for preparation of cell lysates. Remove the media from one set of the clones, resuspend the cells from each well in 30 microliters of extraction buffer from a commercial DNA preparation kit, and transfer to a clean 1.5 milliliter microcentrifuge tube. Heat the solution to 96 degrees Celsius for 15 minutes, and then let cool to room temperature.
Add 30 microliters of stabilization buffer to each microcentrifuge tube and mix well. To identify wild-type and monoallelic or biallelic mutant lines by colony PCR, design two separate sets of PCR primers to amplify regions around the homology arms based on either successful or unsuccessful integration of the resistance cassette. On each side, use a common primer that anneals outside of the homology arm.
To test for the wild-type allele, use the common primer with the corresponding paired primer that is complementary to the endogenous sequence spanning the homology region. To test for the desired mutation, use the common primer with another paired primer, complementary to the inserted resistance cassette. For each sample to be tested, prepare a reaction mixture containing the following, 0.5 microliters of cell lysate, 1.25 microliters of 10X KOD buffer, 0.75 microliters of 25 millimolar magnesium sulfate, 1.25 microliters of two millimolar dNTPs, 0.375 microliters of forward primer, 0.375 microliters of reverse primer, 0.25 microliters of KOD polymerase, and 7.75 microliters of water.
Perform PCR using the following conditions, 95 degrees Celsius for two minutes, followed by 25 cycles of 95 degree Celsius for 20 seconds, primer melting temperature for 10 seconds, and 70 degrees Celsius for 20 seconds per KB.Subsequently, visualize the PCR products by agarose gel electrophoresis. Genome editing using non-homologous end joining was performed to generate ELAVL1 knockout lines followed by dot immunoblot, probing for ELAVL1 and PUM2. Clones that displayed little control signal were marked with an X, and excluded from further analysis.
Clones with low ELAVL1 signals were further tested by western blot. Confirmed candidates are indicated in green, and invalidated candidates are indicated in gray. This graphic shows the ratios of ELAVL1 to control protein signal in increasing order.
In most cases, clonal populations with the lowest relative ELAVL1 signal were correctly identified as knockouts. Candidates generated by the homology directed repair method were screened by colony PCR, and the PCR products visualized by agarose gel electrophoresis. Shown are example results using primers spanning the upstream integration junction using endogenous inside primers and knockout inside primers.
Primers were first validated in both parental cells and bulk selected cells. Results from 24 individual candidate clones are shown together with the expected band patters for wild-type, knockout, and monoallelic deletion clones. The black arrows indicate the correct amplification products, stars indicate biallelic knockout clones.
The biallelic mutant lines were further validated by western blot. Nonhomologous end joining uses a single Cas9 cut and selection-independent screening, thus requiring little up-front preparation, and screening for knockout clones by dot blots can quickly determine if a knockout generation was successful. PCR-based screening accurately detects correct candidates with large genomic perturbations generated by homology-directed repair.
These techniques enable researchers in molecular biology to easily and efficiently identify knockouts, allowing them to explore normal or dysfunctional cellular processes using cell lines. After watching this video, you should have a good understanding of how to isolate and identify clonal knockout cell lines generated by CRISPR/Cas9 mediated genome editing.
