1. Preparation of Basement Membrane Matrix and Coating of Plasticware

1. Place the frozen basement membrane matrix stock from -20 °C onto ice and thaw overnight at 4 °C.
2. After thawing, pipette 2 mg aliquots of basement membrane matrix into pre-chilled eppendorf tubes. Store these at -20 °C until needed.
3. To prepare basement membrane matrix-coated plates, thaw one aliquot on ice until the last piece of ice in the eppendorf tube disappears (usually within ~2 hr).
4. After thawing, add basement membrane matrix to 12 ml of cold (4 °C) DMEM/F12 to make basement membrane matrix coating solution.
5. Add basement membrane matrix solution to the appropriate culture vessel. For a 6-well plate, dispense 1 ml per well. Swirl the plate to make sure the basement membrane matrix solution completely covers each well.
6. Parafilm seal the basement membrane matrix-coated plate/dish and incubate at room temperature for 1 hour before use. Alternatively, store basement membrane matrix-coated plates/dishes at 4 °C and use within 2 weeks of coating.
   NOTE: Add extra DMEM/F12 to the basement membrane matrix-coated plate/dish to prevent drying. Before using 4 °C stored basement membrane matrix-coated plate/dish, place it in a biological safety cabinet and allow it to come to room temperature for at least 30 min.
7. Aspirate basement membrane matrix completely before the addition of medium and cells.

2. Preparation of E8 medium

1. Prepare E8 culture medium by thawing E8 supplement overnight at 4 °C.
2. Remove 10 ml of E8 basal medium from the 500 ml stock and discard.
3. Pipette the entire 10 ml vial of E8 supplement directly into 490 ml of E8 basal medium. Do not warm complete E8 medium in a 37 °C water bath, as repeated temperature fluctuations can degrade the bFGF in the complete E8 medium.
4. Use complete E8 medium within 2 weeks of addition of the supplement.

3. Thawing of iPSCs

1. Remove a vial of frozen iPSCs from liquid nitrogen and place on dry ice.
2. Rapidly thaw the vial in a 37 °C water bath; swirl the vial in the water bath until only a tiny fragment of ice remains.
3. Spray the vial with 70% ethanol and transfer to a biological safety cabinet.
4. Add 1 ml of room temperature E8 medium dropwise directly to the vial.
5. Using a 2 ml pipette, transfer the cell suspension dropwise into 9 ml of E8 medium in a 15 ml conical tube. Swirl the tube frequently to ensure that the cells and medium mix well quickly.
6. Centrifuge the cells at 200 x g for 5 min.
7. Aspirate the supernatant and resuspend the cell pellet in an appropriate volume of E8 supplemented with 10 μM Y-27632.
8. Add the cells to an appropriate number of basement membrane matrix-coated wells, and place in a 37 °C, 5% CO₂ incubator overnight. It is recommended to plate at least 0.2 x 10⁶ iPSC per well of a 6-well plate to enable quick recovery after thawing.

4. Maintenance and Routine Passaging of iPSCs

1. Refresh E8 medium daily.
2. Monitor the morphology and confluency of cells with an inverted microscope. iPSCs of high-quality grow in flat colonies with distinct borders; individual colonies possess a “cobblestone-like” appearance.
3. Passage the iPSCs cells when they reach ~70% confluency.
4. Prepare an EDTA passaging solution by adding 0.9 g NaCl and 500 μl 0.5 M EDTA to 500 ml DPBS. Mix well to dissolve NaCl, and vacuum filter to sterilize. Warm an aliquot of the passaging solution in a 37 °C water bath prior to passaging.
5. To passage, aspirate spent culture medium and wash cells once with an equal volume of warm passaging solution. Aspirate, and pipette enough EDTA passaging solution to coat the cells (1 ml per well of a 6-well plate).
6. Place the cells under an inverted microscope and observe the iPSC colonies. The appearance of holes within colonies and raised borders should become apparent within 2 to 5 min.
7. Carefully aspirate the EDTA passaging solution.
8. Using a 10 ml pipette, dispense 4 ml of E8 medium (if using a 6-well plate) under high pressure directly into each well to be passaged.
9. Collect the iPSC clumps and split into an appropriate number of wells depending on the split ratio from 1:8 to 1:12. Do not over-pipette, as disaggregation of cell clumps will result in poor viability.
10. Place the plate in an incubator, and rock the plate back-and-forth and side-to-side several times to disperse the cells.

5. Preparation of MEFs and iPSCs for Transfection

1. 48 hr prior to transfection, passage iPSCs at ~1:6 into four or more wells of a basement membrane matrix-coated 6-well plate, so that they will be 70% confluent two days hence.
2. The next day, thaw DR4 MEFs into MEF medium consisting of DMEM (high glucose) supplemented with 10% FBS and 1x MEM-NEAA.
3. Plate DR4 MEFs into two 10-cm dishes at ~ 2 x 10⁴ cells/cm² and incubate overnight at 37 °C.
4. On the day of transfection, change MEF medium to E8 supplemented with 10 μM Y-27632 30 min before performing transfection on iPSCs.
5. Optional: 4 hr prior to transfection, supplement pre-transfection iPSC culture with Y-27632 at the final concentration of 10 μM.

6. Gentle-cell Dissociation Reagent Treatment and Transfection of iPSCs using an Electroporation System

1. Remove P3 primary cell transfection solution from 4 °C and allow it to come to room temperature for ~30 min. Add the entire 100 μl supplement to the transfection solution prior to use.
2. Warm gentle-cell dissociation reagent in a 37 °C water bath.
3. Obtain AAVS1 TALENs (pZT-AAVS1-L1 and pZT-AAVS1-R1) and AAVS1-CAG-EGFP donor from -20 °C.
4. Remove iPSCs from the incubator and wash once with DPBS.
5. Add 1 ml of gentle-cell dissociation reagent per well and incubate iPSCs at 37 °C for 5 min, or until greater than 50% of the cells have dissociated from the culture vessel.
6. Pipette the cells up and down a few times using a p1000 pipette to dissociate any remaining cells from the culture vessel, and to break up iPSC clumps.
7. Add 2 ml of E8 medium to each well and pipette up and down several times using a 10 ml pipette to further disaggregate cell clumps into single cells.
   NOTE: Transfection efficiency declines significantly if cell clumps are not sufficiently disaggregated.
8. Collect iPSCs in a 15 ml conical tube, and centrifuge at 100 x g for 3 min.
9. Aspirate supernatant and resuspend cells in a minimal amount of E8 medium.
10. Count the cells using a hemocytometer after application of a vital stain such as 0.4% Trypan blue. Ensure that cells are sufficiently dissociated while counting (1-3 cells per “clump”).
11. Dispense 3 x 10⁶ cells into each of two 15 ml conical tubes, and spin down again at 100 x g for 3 min.
    NOTE: Low speed centrifugation reduces the cell stress and allows easy re-suspension of iPSCs before electroporation.
12. Set the electroporation system to the cell-type specific program for the human embryonic stem cell line H9 (Program CB-150).
13. After centrifugation, aspirate the supernatant from the cell pellets. To one pellet, add 10 μg of the HR donor as control sample. To the other pellet add 10 μg of the HR donor, along with 5 μg of each TALEN (pZT-AAVS1-L1 and pZT-AAVS1-R1) as experimental sample.
14. Resuspend each cell pellet in 100 μl of P3 primary cell transfection solution, and transfer to a cuvette.
15. Perform the transfection and immediately add 500 μl of room temperature E8 medium to each cuvette.
16. Transfer the transfected iPSCs dropwise to one 10-cm dish containing DR4 MEFs prepared in step 5.4.