Differentiation, Maintenance, and Analysis of Human Retinal Pigment Epithelium Cells: A Disease-in-a-dish Model for BEST1 Mutations

1. Differentiation of hPSC to RPE

1. Maintain and passage hPSCs as previously described¹⁸.
   NOTE: All cells (including hPSCs and hPSC-RPEs) are grown in 37 °C at 5% CO₂ throughout the duration of the growth and differentiation protocols.
2. Prior to differentiation, split confluent hPSCs to pre-coated 6-well plates.
   1. To coat plates, thaw basement membrane matrix on ice for about 1 h, suspend liquidized matrix in 4 °C DMEM medium at a 1:50 dilution, add 800 µL coating mixture to each well, and incubate for at least 1 h at 37 °C.
   2. When coating is complete, aspirate the mixture and immediately add 1.5–2 mL medium into the wells.
   3. To lift hPSCs from the old culture plates, incubate cells with PBS plus 0.5 mM EDTA for 5 min at room temperature, aspirate the solution, and resuspend cells in small clumps with culture medium.
   4. Seed cells at ~20% confluency in the new plates.
      NOTE: Store basement membrane matrix in 100 µL aliquots at -20 °C and use one aliquot to coat a 6-well plate. The time for coating can be extended up to 4 h at 37 °C, or overnight at room temperature. It is important to suspend and seed hPSCs as small clumps of 3–5 cells rather than single cells.
3. When cells are grown to full confluency, replace the culture medium with differentiation medium (4 mL/well) (Table 1, without activin-A). Culture for 14 days, changing the medium (4 mL/well) 3x/week.
   NOTE: The hPSC plate density doubles approximately every 24 h. Significant cell death is expected after the start of the differentiation procedure. Aggregates of dead cells and debris in the wells can be seen by the naked eye during medium changes.
4. From day 15 to day 28, supplement differentiation medium with 100 ng/mL human activin-A (Table 1, with human activin-A). Change the medium (4 mL/well) 3x/week.
5. Stop activin-A supplementation starting on day 29 (Table 1, without human activin-A). Continue cell culturing in differentiation medium for 8–10 weeks until pigmented hPSC-RPE clusters appear (Figure 1A-B).
   NOTE: Clusters of pigmented cells can be seen as small dark dots on a cell culture plate by the naked eye (Figure 1A-B, top). Individual pigmented cells with the signature cobblestone shape can be seen under a 20X microscope (Figure 1A-B, bottom).

2. Isolation and Culture of Differentiated RPE Cells

1. Remove the differentiation medium, wash once with PBS, add 1 mL of 0.05% trypsin plus 1 U/µL of collagenase into each well, and incubate at 37 °C for 20–30 min. Remove the trypsin/collagenase and gently add 2 mL of pre-warmed RPE medium into each well of the 6-well plate (Table 2).
   NOTE: Use an inverted microscope to monitor the morphology of the hPSC-RPE cells, which are polygonal before trypsin/collagenase treatment, and become round after sufficient digestion.
2. Use a Bunsen burner to pull glass Pasteur pipettes.
   1. With both hands, hold a long (9 inch) Pasteur pipette at both ends so it is horizontal, and above the Bunsen burner about 2/3 down the length of the thin part of the pipette.
   2. Once the glass becomes soft from the flame, quickly pull the two ends apart, such that a micro scraper-like tip is formed at the new end of the pipette (Figure 2).
   3. Repeat several times to create multiple "cell scrapers". Spray the pulled pipettes with 70% ethanol for sterilization and air-dry them in the cell culture hood.
3. Under a microscope, use a micro "cell scraper" to gently dissociate pigmented hPSC-RPE clusters from the plate.
   1. Gently tap (do not scrape) directly on the pigmented cells, which will float up into the culture medium once they dissociate from the bottom of the well.
   2. Immediately use a 20 µL micropipette to gently suction the dissociated hPSC-RPE cells and transfer them to a pre-coated 6-well plate with RPE medium²⁴.
   3. Collect ~5 x 10³ cells (~10% confluency as observed under a 20X microscope) in each pre-coated well of a 12-well plate by repeating steps 2.3.1–2.3.2 multiple times, as needed. Bring the final volume of medium to 2 mL.
      NOTE: To prevent dissociated hPSC-RPE cells from floating away, avoid movement of the plates during the dissociation process such that the dissociated cells are floating in the medium but still remain locally concentrated.
4. Culture the newly isolated hPSC-RPE cells (P₀) on 12-well plates in RPE medium for another 6-8 weeks to allow them to form a pigmented monolayer (Figure 1C).
5. Change the medium 3x/week. When changing the medium, leave approximately 1/3 volume of the old medium in each well, and add 2/3 volume of fresh, pre-warmed RPE medium. At this stage, use 2 mL of RPE medium for each well in a 12-well plate (thus, exchange 1.3 mL of medium each time).
   NOTE: The monolayer of P₀ hPSC-RPE cells may form domes, suggesting that the cells are actively transporting fluids, and are anchored by tight junctions²⁵. Therefore, dome formation is an indicator of mature RPE status.

3. RPE Fate Validation by Immunoblotting

1. Make the hPSC-RPE cell lysate using mammalian protein extraction buffer supplemented with proteinase inhibitor cocktail. Incubate the cell lysate on ice for 30 min before centrifugation at 13,000 x g for 15 min at 4 °C to discard undissolved cell debris. Measure the total protein concentration of lysate.
2. Denature 40 µg of total protein in Laemmli SDS sample buffer at 75 °C for 10 min. Separate the protein samples on a 10% Tris-glycine SDS-PAGE gel at a constant voltage of 90 V for 15 min and then 150 V until the dye front reaches the bottom of the gel.
3. Transfer the proteins onto a nitrocellulose membrane in pre-cooled Tris-glycine buffer supplemented with 10% methanol at 100 V for 1 h.
4. Block the membrane in Tris buffered saline with 0.1% non-ionic detergent (TBST) and 5% non-fat dry milk for 1 h at room temperature.
5. Incubate the membrane with primary antibodies diluted in the blocking buffer at 4 °C overnight. Dilute the antibodies targeting RPE specific marker proteins as follows: RPE65, 1:1,000; CRALBP, 1:1,000; BEST1, 1:1,000. Dilute the antibody against β-actin at 1:10,000 to detect β-actin as a loading control.
6. Wash the membrane with TBST 3 times, with 5 min for each wash.
7. Incubate the membrane with fluorophore conjugated goat anti-mouse IgG secondary antibody diluted 1:10,000 in blocking buffer for 1 h at room temperature.
8. Wash the membrane with TBST 4 times, with 10 min for each wash.
9. Detect the proteins using an infrared imaging system (Figure 3).

4. Checking BEST1 Subcellular Localization by Immunostaining

1. Wash hPSC-RPE cells with 2 mL of PBS once and fix in 2 mL of 4% paraformaldehyde for 45 min at room temperature.
2. Wash with 2 mL of PBS twice, and incubate the cells in 2 mL of PBS with 0.1% non-ionic surfactant and 2% donkey serum for 45 min to block the non-specific binding sites.
3. Incubate the cells with BEST1 antibody (1:200 dilution) for 2 h at room temperature.
4. Wash the cells with 2 mL of PBS 3 times. Incubate with fluorophore conjugated secondary IgG (1:1,000) for 1 h at room temperature.
5. Wash with 2 mL of PBS 3 times to remove unbound secondary antibody.
6. Observe stained cells by confocal microscopy (Figure 4).

5. Recording Ca²⁺-dependent Cl^– Current in hPSC-RPE by Whole-cell Patch Clamp

1. 24–72 h before the patch clamp, split a fully confluent well (on a 12-well plate) of mature P₀ hPSC-RPE cells (cobblestone-shaped and pigmented). Aspirate the medium, wash once with PBS, and add 1 mL of 0.05% trypsin plus 1 U/µL collagenase into the well.
   NOTE: Pre-warm the trypsin/collagenase to 37 °C right before use.
2. Incubate at 37 °C for 8 min.
   NOTE: After this step, the hPSC-RPE cells are still adherent and connected.
3. Use a 1 mL micropipette to gently wash the cells off the plate, and transfer cells in the digestion mixture to a 15 mL conical tube. Wash the well with 1 mL of pre-warmed trypsin/collagenase to collect the residual cells and combine them into the 15 mL conical tube.
   NOTE: The hPSC-RPE cells are still in large clumps at this point. Do not try to break the cell clumps by vigorous pipetting. Some clumps may stick to the inner wall of the 1 mL tip.
4. Incubate the conical tube in a 37 °C dry bath for 8 min.
5. Use a 1 mL micropipette to gently pipette up-and-down 10–15 times the digestion mix.
   NOTE: Cell clumps become much smaller but still visible. Avoid vigorous pipetting.
6. Repeat the previous two steps.
   NOTE: Most of the cells should be in single-cell suspension after pipetting. There may still be some tiny cell clumps, which are acceptable. Optional: Incubate the conical tube at 37 °C for an additional 5 min followed by gentle pipetting. The total time in the trypsin/collagenase at 37 °C should not exceed 30 min (8 + 8 + 8 + 5 = 29).
7. Add 5 mL of pre-warmed RPE medium to the 15 mL conical tube containing the digested cells. Spin at 200 x g for 5 min to collect the cells. Re-suspend and seed cells on pre-coated 35 mm dishes at 10–20% confluency (e.g., a 100% confluent well on a 12-well plate to five 35 mm dishes for 10% confluency).
   NOTE: At all times, avoid vigorous pipetting, which may cause significant cell death as represented by apparent dark cell debris in the supernatant after centrifugation. Well-separated single-cell seeding is essential for patch clamp recording.
8. Perform the whole-cell patch clamp as described previously^{18,26,27,28,29,30,31}. Acquire current traces from a family of step potentials (-100 to +100 mV from a holding potential of 0 mV) (Figure 5).
   NOTE: The recipes of internal and external solutions are described in Table 3 and Table 4, respectively.