Establishment of Genome-edited Human Pluripotent Stem Cell Lines: From Targeting to Isolation

The procedures described in this manuscript were reviewed and approved by the UC Berkeley Stem Cell Research Oversight Committee.

1. Prepare Stem Cells for Editing

1. Grow and culture human pluripotent stem cells (hPSCs) on a 6-well plate containing 2.4 × 10⁶ cells/plate of mitomycin C-inactivated mouse embryonic fibroblast (MEF) feeders grown on gelatin ³². Maintain hPSCs in 3 ml of human embryonic stem cell media per well (hESC media) and grow in a 37 °C incubator with 3% O₂/5% CO₂.
   Note: For the success of this protocol, it is not necessary to maintain hPSCs in a low-oxygen incubator; however it is important to note that hPSCs proliferate faster in a high O₂ environment, so times and cell numbers should be adjusted accordingly. It should also be noted that hPSCs maintained in low oxygen have lower rates of spontaneous differentiation ³³.
   1. To make 500 ml hESC media combine 380 ml DMEM/F12, 75 ml Fetal Bovine Serum (FBS), 25 ml KnockOut Serum Replacement (KSR). Add Glutamine (1 mM final concentration), 5 ml 100x Non-essential Amino Acids, 100 units/ml Penicillin-Streptomycin (P/S), basic Fibroblast Growth Factor (bFGF) (4 ng/ml final concentration), and 2-mercaptoethanol (5.5 µM final concentration).
2. Change media by removing entire volume of media (3 ml) using a glass pipette and vacuum. Replace with 3 ml warm hESC media using a serological pipette. Repeat media change every day until hPSCs are approximately 50% confluent (Day -1).
3. One day before targeting (Day -1), change hESC media, removing the old media and adding warm hESC media supplemented with 10 µM Y-27632.
4. Also on Day -1, prepare one to two 6-well or 10 cm plates of drug resistant MEF feeder cells from DR4 mice (2.4 × 10⁶ cells/plate) ³⁴.
   Note: In general, 6 well plates are advantageous over 10 cm plates, as they accommodate more media. 6 well plates also ensure that different clones from different wells are independent. However, a 10 cm plate will allow easier picking, depending on the microscope available for this process.

2. Editing Pluripotent Stem Cells

1. Prepare transfection solutions by pipetting 5 µg each of ZFN 1 and 2, TALEN 1 and 2, or 15 µg of the CRISPR/Cas9 px330 encoding plasmid (Figure 1) into a 1.5 ml tube. Pipette 30 µg of the repair plasmid into this tube as well. Finally, pipette 1x Phosphate Buffered Saline (PBS) into the tube to bring the volume up to 300 µl.
   Note: Prepare plasmids as a midiprep (any kit is suitable) with a minimum concentration of 300 ng/µl in order to keep the total volume of the transfection solution under 300 µl. It is not necessary to perform phenol/chloroform extraction, to linearize the plasmid, or to use an endotoxin free plasmid prep kit.
2. Remove media from hPSCs using a glass pipette and vacuum. Pipette 2 ml warm 1x PBS into each well to wash the cells.
3. Remove the PBS immediately using a glass pipette and vacuum. Add 0.5 ml 0.25% Trypsin-EDTA solution directly onto cells in each well. Place in the incubator (37 °C/5% CO₂/3% O₂) for approximately 10 min. or until feeder layer starts to lift off the plate.
4. Add 2 ml warm esWash media (470 ml DMEM/F12, 25 ml FBS, 100 units/ml P/S) to each well to stop the trypsin reaction.
5. Ensure that feeder cells come off as a sheet. Pipette the contents of each well into a single 50 ml conical tube, combining all wells. Triturate cells using a 10 ml serological pipette. It is not necessary to break up the feeder layer; hPSCs will come off of the feeder layer by gentle trituration. There should be about 15 ml of the cell suspension.
6. Add 25 ml of esWash media to the tube to bring the volume up to 40 ml total. Allow large feeder chunks to settle at the bottom of tube for 1-2 min. Remove the supernatant (~38 ml) from the tube using a serological pipette and deposit into a new 50 ml conical tube.
7. Spin down for 5 min at 190 x g. Remove the supernatant from the tube using a glass pipette and vacuum. Be sure to not disturb the cell pellet. Resuspend the cells in 500 µl 1x PBS. Combine with plasmid transfection solution prepared earlier. Count the cells at this step. Use 5-10 million cells per electroporation.
8. Pipette the entire 800 µl suspension into a 4 mm electroporation cuvette, place on ice for 3-5 min. Electroporate cells using the exponential program on the electroporation system with the following parameters: 250 V, 500 µF, ∞ resistance, and 4 mm cuvette size. After electroporation, place cuvette back on ice for 3 min.
   Note: Observe the time constant of the electroporation on the electroporation system. The time constant varies with cell number and DNA purity. Successful transfections usually have a time constant between 10-14 msec when using the listed conditions and a gene pulser II. Lower electroporation efficiencies can occur when time constants varies from these values.
9. Resuspend electroporated cells in 18 ml warm hESC Media supplemented with 10 µM Y-27632. Plate 3 ml of this single cell suspension into each well of a 6 well plate with DR4 feeder cells. Return to incubator (37 °C/5% CO₂/3% O₂).

3. Selection of Positive Colonies

1. Day 2, remove all media using a glass pipette and vacuum. Replace with 3 ml of warm hESC media supplemented with 10 µM Y-27632. Day 3, remove all media using a glass pipette and vacuum. Replace with 3 ml of warm hESC media without including Y-27632. Day 4, remove all media using a glass pipette and vacuum. Replace with 3 ml warm hESC media supplemented with antibiotics for selection. Return to incubator (37 °C/5% CO₂/3% O₂).
   Note: The type of antibiotic used will depend on the resistance cassette included in the repair template. When working with WIBR#3 hPSCs (NIH registry 0079), 0.5 µg/ml puromycin, 70 µg/ml G418 (geneticin), and 35 µg/ml hygromycin have been used successfully. Concentrations for selection should be determined empirically by establishing the minimal concentration of antibiotic needed to kill wild-type cells within approximately one week.
2. Days 5-12, change media daily, replacing old media each time with warm hESC media supplemented with antibiotics.
   Note: Expect a large amount of cell death. Individual colonies will become apparent around day 8-10. Regular hESC media without antibiotics can be used after 12-14 days of continuous selection. If cell density is high or cell death is slow, acidification of the media should be avoided and it may be necessary to increase the media volume (up to 4-5 ml) during the first few days of selection.

4. Picking Selected Colonies (Day 12-14)

1. Observe colonies on day 12-14. Observe the colonies that are ready to pick on a dissection microscope and ensure that they do not contain cells that are starting to differentiate. The approximate size should be 800-1,200 µM. If colonies reach this size before day 12, it is recommended that they should be picked then.
   Note: For each targeting experiment, pick as many colonies as necessary to ensure the desired genotype is isolated. As an example, AAVS1 editing with the AAV-CAGGS-EGFP repair template has shown consistently robust integration, and requires only 12-24 colonies to be picked to obtain approximately 5-10 heterozygously targeted clones. Other targeting experiments may require more colonies to be picked (Table 1). The frequency of correct targeting events depends on factors such as the efficiency of the SSN to introduce a DSB, the size of insert, and the selection strategy. In the case of the gene trap approach for the AAVS1 locus presented here, the selection marker is only expressed when correctly integrated at the target site, reducing the number of colonies required to obtain a properly targeted clone.
2. One day before picking, prepare 12-well plates of MEFs (2.4 × 10⁶ cells/plate, one well will be used for each colony picked).
3. On the day of picking, remove all media from the 12 well MEF plates using a glass pipette and vacuum and replace with 1 ml hESC media. Also on day of picking, change hESC media on the 6 well plates that are going to be picked.
4. Pull glass pipettes for mechanical dissociation of individual colonies from the feeder layer. Lower the pipettes over an in-hood Bunsen burner and softened at the expansion point until glass is malleable. Quickly remove from flame and pull the pipette apart creating an angular point. Break the point approximately 2 cm from the bent axis, leaving a narrow channel. Polish the end of the channel by exposing to flame for 1-2 sec.
   Note: Optionally pick colonies a p20 pipet tip, however, this may reduce the clonality of the culture, as the dull tip may dislodge greater amounts of cells from each colony into the media, potentially contaminating subsequent pickings.
5. Assemble picking device by taking a p1000 filter pipette tip and attaching a suction bulb to the narrow end. Insert the pulled glass pipette in the wide end.
6. Place a 6-well plate of hPSCs to be picked on the stage of a dissection microscope mounted in a tissue culture hood. Compress the bulb of the picking device and gently excise and cut an individual colony into 10-20 equally sized pieces, taking care not to release pieces into the media. Take excised hPSC pieces of the colony into the pipette by releasing the bulb. Try to take as little media as possible while transferring.
7. Transfer the individual colony to a single well of a 12-well MEF plate, compressing the bulb again directly into the well, releasing the now broken colony. Label each well to allow unique identification of single-cell derived clones. Repeat for as many colonies as necessary, changing the glass pipette each time.
   Note: Picking colonies may take some time to learn. It is recommended that the experimenter practice on some control cells before trying to isolate targeted colonies.
8. Return 12 well plates to the incubator (37 °C/3% O₂/5% CO₂), gently rocking the plate first to avoid accumulation of cells in the middle of each well.
9. The next day and each subsequent day, remove full volume of media using a glass pipette and vacuum and replace with 1.5 ml of warm hESC media until cells are about 50% confluent (this usually takes 10-12 days).
10. After 10-12 days, pick one to two colonies from each well and transfer to new 12-well MEF plates repeating steps 4.1-4.8 to generate a replica plate.
11. Extract DNA (see below) from the remaining colonies in each well of the original MEF plates.
    1. Remove media from all wells using a glass pipette and vacuum. Pipette 1 ml 1x PBS onto each well to wash cells. Remove PBS using glass pipette and vacuum. Pipette 250 µl of cell lysis buffer (final concentrations in H₂O: 10 mM Tris HCl, 5 mM EDTA, 0.2% SDS, 200 mM NaCl, 0.08 mg/ml Proteinase-K) onto each well. Place in incubator (37 °C/3% O₂/5% CO₂) O/N.
    2. The next day, pipette contents of each well into individual 1.5 ml tubes. Pipette 250 µl of isopropyl alcohol into each tube to precipitate the DNA. Shake tube vigorously. A white precipitate should be visible.
    3. Spin each tube down for 3 min at 13,000 rpm in a table top centrifuge. After spin down, discard supernatant by decanting into liquid waste. A small DNA pellet should remain stuck to the bottom of the tube. Pipette 250 µl of 70% ethanol into each tube to wash the DNA pellet. Shake vigorously. Spin each tube down for 3 min at 13,000 rpm in a table top centrifuge.
    4. After spinning down, discard supernatant by decanting into liquid waste. A small DNA pellet should remain stuck to the bottom of the tube. Pipette out the remainder of the supernatant so there is no liquid left in the tube. Leave tube open to dry on benchtop for 5-10 min. After drying, resuspend DNA in 250 µl TE buffer. Place at 37 °C for 6 hr to allow DNA to dissolve.
12. Genotype each sample using a pre-optimized PCR or southern blot strategy.
    Note: For AAVS1 targeting using the AAV-CAGGS-EGFP repair template, the southern blot strategy can be found in Hockemeyer et al., 2009 ⁴. A comprehensive southern blotting protocol can be found in Southern, 2006 ³⁵. For multiplex PCR genotyping of the same experiment, primers and conditions can be found in Table 2 ³⁶.
13. Discard wells that are not properly targeted and continue changing hESC media on properly targeted cells every day. Freeze down the cells when they are approximately 50% confluent, see below for freezing protocol.
    1. One day before freezing, change hESC media, removing the old media and adding warm hESC media supplemented with 10 µM Y-27632.
    2. On the day of freezing prepare 0.5 ml each of solution A and solution B per well of a 12-well plate that is being frozen down in two 15 ml conical tubes. Place solutions on ice. Solution A: 50% hESC media and 50% FBS; Solution B: 80% FBS and 20% Dimethyl Sulfoxide (DMSO).
    3. Dissociate hPSC colonies into single cells before freezing. Follow steps 2.2-2.8 to do this, scaling volumes accordingly. Fully resuspend cell pellet in 0.5 ml of solution A.
    4. Add 0.5 ml solution B to cell suspension, pipette up and down to make cell suspension homogenous. Take the total volume of the cell suspension (~1 ml) and deposit into a 2 ml cryotube. Screw cap on tightly.
    5. Place cryotube in a -80 °C freezer O/N. The day after freezing, remove frozen cells from -80 °C freezer and immediately place into liquid nitrogen tank for long-term storage.