Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

The goal of this protocol is to generate human induced pluripotent stem cells that express endogenous fusion proteins fused to end frame N or C terminal fluorescent tags. To achieve the desired knock in, we use the CRISPR-Cas9 system and a ribonucleoprotein based approach using wild type S.pyogenes Cas9 protein and a synthetic two part CRSPR RNA and tracrRNA. This RNP is co-delivered with the desired fluorescent tag sequence as a donor plasmid via electroporation.

Putatively edited cells expressing the fluorescent tagged protein, are enriched by FACS. Clonal lines are then generated for analysis of precise editing outcomes. This gene editing protocol has many advantages over traditional over-expression constructs.

Because a fluorescent tag is introduced into the genome, the subcellular localization and dynamics of the resulting fusion protein can be studied under endogenous regulatory control. Here we use induced pluripotent stem cells as a model system because it allows us to study the tagged proteins in diploid non-transformed cells. Additionally, since the cells can be differentiated into multiple cell types, like the cardiomyocyte cells shown here, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

However, with careful optimization, this protocol could be adapted for editing the genomes of other mammalian cell types. Hi, I'm Amanda Haupt, I'm part of the stem cells and gene editing team here at the Allen Institute for Cell Science. Today, we have used this approach to generate over 35 different cell lines expressing endogenous fusion proteins.

We use these to study a wide variety of unique cellular structures and processes. This approach has been successful for up to 95%of the loci attempted to date with our group. This method will be particularly useful to cell biologists who are interested in better understanding the dynamics of a protein in live cells.

However once the edited clonal lines are generated the possibilities for study are endless. With this method we're able to consistently identify clones that have monoallelic fluorescent protein tags without any other disruptions in the genomic sequence. We are also occasionally able to obtain biallelically edited clones.

In this video we will show some of the key steps in transfecting, FACS enriching, and isolating clonal cell lines that express endogenously regulated fluorescent fusion proteins. Two to three hours before transfection, treat the hiPSCs with ten micromolar ROCK inhibitor supplemented into growth media. This will promote survival after electroporation.

Cells will have spiky appearance after this treatment. Cells should be removed from the tissue closure plate using a single cell dissociation region and the cell suspension collected in a conical tube prior to transfection. Prepare an aliquot of 1.84 times 10 to the six cells for each experimental condition to be transfected.

Prepare ribonucleoprotein complex tubes for each transfection by adding 2.88 microliters of 10 micromolar guide RNA and 2.88 microliters of 10 micromolar Cas9 to a 1.5 mil tube. Incubate at room temperature for a minimum of 10 minutes, a maximum of one hour. Pellet the cell aliquot prepared in the previous step at 211 times G for three minutes at room temperature.

Then aspirate the supernatant and race to spin the cell pellet in 220 microliters of electroporation buffer. Add 220 microliters of re suspended cells into the 1.5 mil RNP complex tube previously prepared. Add 4.6 microliters of one microgram per microliter donor plasmid.

Use the nucleofection tip and pipette to mix the two contents two to three times then transfer a hundred microliters of suspension to the prepared electroporation device. Avoid introducing any bubbles in the tip. Apply thirteen hundred volts for one pulse of thirty milliseconds.

Gently transfer the suspension into a prepare six well plate with a swirling motion. Disperse the cells by gently moving the plate side to side and front to back. After repeating for all of the experimental conditions and controls, incubate transfected cells at 37 degrees celsius and 5%CO2 for 24 hours changing the media to regular growth media without ROCK inhibitor at 24 hours.

Then continue to feed the hiPSCs every 24 hours for 72 to 96 hours monitoring their confluency. When hiPSCs reach 68 to 80%confluency proceed to the FACS enrichment. Two to three hours before transfection treat hiPSCs with 10 micromolar ROCK inhibitor supplemented into the growth media to promote survival after FACS enrichment.

To sort cells, remove them from the plate using the single cell dissociation reagent then filter the hiPSC suspension through 35 micron mesh filter into polystyrene round bottomed tubes. Sort cells using forward scatter and side scatter including:height versus width, triplets debris, and doublets. Use live buffer only control cells to set the FP positive gate such as that less than 1.1%of buffer only cells fall within the gate.

When sorting stem cells adapt instrument settings to promote cell survival as suggested in the discussion section. Typical knock in efficiencies as determined by the percent FP positive cells by FACS can range anywhere from 0.1%to 5%Sort cells into a 1.5 to 15 mil polypropylene tube containing 0.5 to 2 mils of room temperature growth media supplemented with 10 micromolar ROCK inhibitor. Centrifuge the collected cells at 211 times G for three minutes at room temperature.

Carefully aspirate the supernatant and re suspend the cell pellet in 200 microliters of growth media supplemented with 10 micromolar ROCK inhibitor. Transfer up to three thousand assorted cells to a single well of a fresh matrigel coated 96 well plate. Incubate sorted cells at 37 degrees celsius in 5%CO2 for 24 hours changing the media to growth media supplemented with 5 micromolar ROCK inhibitor at 24 hours.

Then at 48 hours, continue feeding cells regular growth media, no ROCK inhibitor every 24 hours for 72 to 96 hours monitoring the confluency. Once the cells are mature enough to be passaged out of the 96 well plate we recommend passaging them gradually into larger plates. So for example from one well of a 96 well plate, move them up to one to two wells of a 24 well plate.

And then from there into a 6 well plate, et cetera, until you have enough cells for downstream purposes. Usually three to four million cells minimum. To generate clonal lines from the FACS enriched population of punitively edited cells, passage into single cell suspension and plate ten thousand cells onto a 10 centimeter plate and allow the colonies to grow for five to seven days.

When the colonies are visible macroscopically they're large enough to be isolated. On a dissecting microscope use a P200 pipette to gently scrape and aspirate individual colonies from the plate surface. Transfer the volume containing the colony to a single well of the 96 well plate.

After all colonies have been transferred, incubate the plate in a tissue culture incubator at 37 degrees celsius and 5%CO2 for 24 hours. Change the media to regular growth media without ROCK inhibitor at 24 hours. Then continue feeding the cells every 24 hours for 72 to 96 hours until colonies have approximately tripled in size.

The cells will need to be grown and passaged in 96 well plates in order to have enough cells to cryopreserve antalise for genomic DNA extraction. See the written protocol for more details. To passage in 96 well plates use an eight channel aspirator to remove and discard media from the first column.

Using a P200 multichannel pipette add about 200 microliters of DPBS to the first column of the plate to wash the cells. Using an eight channel aspirator then remove and discard the DPBS wash from the first column. Then add 40 microliters of dissociation reagent to the first column.

Repeat these steps working one column at a time for up to a total of six columns of the 96 well plate. Change tips to be sure to not cross contaminate wells. Place the plate in the incubator for three to five minutes from the time the dissociation reagent was added to the first column.

When the cells in the first column of the plate have begun to lift off the bottom, use a P200 multichannel pipette to add 160 microliters of DPBS to the first column and gently triturate the cells in the 12 o'clock, 3 o'clock, 6 o'clock, and 9 o'clock positions of each well. Transfer the entire volume with cell suspension to a V-bottom 96 well plate. Repeat these steps for the remaining columns that have dissociation reagent in them changing tips as to not cross contaminate the wells.

Spin the V-bottom plate in the centrifuge at 385 times G for three minutes at room temperature. The sub shown up until this point for 96 well passaging are performed in the same way any time the cells are passaged in this plate format. Next we will show you how to proceed if you are ready to cryopreserve the cells in 96 well format.

See the written protocol for more information. After the centrifugation step shown in the previous section, aspirate the supernatant using a P200 multichannel pipette and then re suspend in 60 microliters of growth media supplemented with 10 micromolar ROCK inhibitor. Repeat for all wells, changing tips as to not cross contaminate.

Transfer 30 microliters of cell suspension to a non-matrigel coated 96 well tissue culture plate. Transfer the remaining 30 microliters of suspension into a sister plate then quickly add 170 microliters of freezing buffer to each well without mixing. Note that this process is done in duplicate sister plates so that a back up population of cells exists after one of the individual plates is thawed.

Putting cryopreserved cells into only every other column of the 96 well plate allows for faster thawing. Wrap the plate with parafilm and place it in a room temperature styrofoam box with a lid. Place the whole box in the minus 80 degree celsius freezer.

After 24 hours plates can be transferred out of the styrofoam box and sort at minus 86 degrees celsius for up to four weeks. Cryopreservation of human induced pluripotent stem cells in 96 well format allows for a temporal break in cell culture. The researcher can perform genomic validation assays on the clonal mines to identify the genetically desirable clones to thaw for future studies.

Four days after transfecting the WTC hiPSC line with a target specific guide RNA Cas9 protein and a carefully designed donor template plasmid, 0.95%of the cells were observed to be expressing GFP. FACS was used to sort out all of the GFP positive cells as an enriched population. Preliminary microscopy studies revealed the expected localization of the mEGFP nuclear lamin B1 fusion protein to the nuclear envelope.

The fluorescently labeled protein was observed in about 95%of the cells with varying intensities. After clonal line generation, clones with uniform mEGFP intensity were obtained and further analyzed for precise editing. To conclude, we have shown a transfection method for gene editing human induced pluripotent stem cells for the expression of endogenously regulated fluorescent fusion proteins.

Because typical knock inefficiencies are low, some enrichment strategy like the FACS method shown here results in a population of putatively edited cells expressing the fusion protein. The enriched population can be used for biological studies or clonal lines can be generated as described in this method making it possible to identify cell lines that are precisely gene edited. It's important to remember that Cas9 induced double strand breaks in the genome are often repaired imprecisely despite incorporation of the tag at the intended locus.

Additionally in monoallelically edited clones, the untagged allele frequently contains indels most likely caused by non-homologous end joining repair. These finding underscore the importance of genetic validation after a CRISPR Cas9 knock in only revealed through analysis of the genome of clonal cell lines. Expression of the expected fusion protein alone does not guarantee precise editing.

When using this protocol for editing human induced pluripotent stem cells take care to follow the recommendations detailed in the written protocol to ensure gentle handling of the cells for optimal cell survival and minimal spontaneous differentiation. We hope that this video and the associated protocol and materials list help you perform similar experiments in your own lab on your favorite protein. Thanks for watching.