A Simplified Method for Generating Kidney Organoids from Human Pluripotent Stem Cells

All experiments using hPSCs were performed in compliance with institutional guidelines, and were carried out in a Class II biosafety hood with appropriate personal protective equipment. All reagents are cell culture-grade unless stated otherwise. All cultures are incubated at 37 °C, 5% CO₂ air atmosphere. At all stages of the assay, embryoid bodies or kidney organoids can be collected, and fixed or prepared for analysis. The hPSC lines used to generate this data have been fully characterized and published¹⁸.

1. Preparing culture plates

NOTE: Approximately 1 h prior to splitting hPSCs, coat 2 x 100 mm tissue culture plates with a stem cell qualified basement membrane matrix extract (BME). One may pre-coat the plates, seal them with a paraffin film and store at 4 °C according to manufacturers' instructions.

1. Prepare 2 x 100 mm tissue culture-treated plates (1 for kidney organoid assay, 1 to maintain the cell line) and a 15 mL conical tube in the Class II biosafety hood.
2. Aliquot 8 mL of cold, serum-free Dulbecco's Modified Eagle Medium (DMEM) into a 15 mL conical tube and ~4 mL into each of the 100 mm plates, enough to cover the bottom of each plate with medium.
3. Take a 100 µL aliquot of BME out of the freezer (-20 °C). Using a 2 mL serological pipette, take ~1 mL of cold DMEM from the 15 mL conical tube. Slowly thaw the BME aliquot by gently pipetting up and down with the cold DMEM, avoiding making bubbles.
   NOTE: Do not let BME aliquot sit at room temperature. Use immediately.
4. Transfer the thawed DMEM/BME back into the 15 mL conical tube with the remaining DMEM. With a 10 mL serological pipette, gently mix the diluted BME by pipetting up and down at least 8 times to evenly disperse the BME, avoiding making bubbles.
5. Transfer 4 mL of the diluted BME into each plate with DMEM and gently swirl the plate so that the BME is evenly distributed. Incubate the coated plate for 1 h at room temperature or 30 min at 37 °C.
   NOTE: Use 50 µL of BME per 100 mm plate. Use of other hPSC culture media and cell lines may require different concentrations of BME.

2. Passaging hPSCs

NOTE: For routine hPSC culture, passage cell lines at 70-80% confluency.

1. Aspirate the culture medium from the hPSC plate to be passaged. Add ~ 8 mL of Dulbecco's phosphate-buffered saline (DPBS) to the hPSC plate and gently swirl to wash the cells.
2. Aspirate DBPS and add 2 mL of gentle cell dissociation reagent (GCDR) to the 100 mm plate, drop by drop on top to cover the cells.
   NOTE: Other dissociation reagents may also be used. Adjust accordingly.
3. Incubate at room temperature for ~6-8 min until the colonies are breaking up and cells are refractive under phase contrast (Figure 2A).
   NOTE: The timing may vary between cell lines. Adjust accordingly.
4. While incubating, prepare a 50 mL conical tube. Add 16 mL of hPSC medium (8 mL per 100 mm plate) and add Rho-associated kinase inhibitor (ROCKi) to a final concentration of 5 µM.
5. Aspirate DMEM from the BME-coated plates, and add 8 mL of hPSC medium plus ROCKi to each plate.
6. When the cells are ready (as described in point 2.3, Figure 2A), aspirate the GCDR and tilt the plate ~45° towards the experimenter and scrape the cells with a cell lifter.
   NOTE: If cells are detatching, omit aspirating GCDR and proceed.
7. Turn the plate ~90° and scrape again to lift the remaining cells. Keep the plate ~45° and wash the cells down with 3 mL of hPSC medium using a 10 mL serological pipette.
8. Gently pipette up and down to break up large clumps (no more than 2-3 times) and seed the cells at the appropriate ratio for the cell line of interest onto the prepared plates. Place the plate with the cells in the incubator and move the plate gently in figure eight motions to distribute the cells evenly.
   NOTE: In this experiment hPSC lines were split at a ratio of 1:5, this may vary for other cell lines and conditions. Leave the plate undisturbed over night.
9. After ~ 24 h, examine the cells for attachment. Look for small individual colonies attached. Aspirate the spent medium and replenish with 8 mL of fresh hPSC medium (no ROCKi added).
10. Continue observing and feeding daily until the cells reach ~60% confluency to start the kidney organoid assay (usually reached 48 to 72 h post passaging). The colonies will ideally be discrete and not merging (Figure 2B).
    NOTE: It is very important to limit the cells to no more than 80% confluency in order to maintain their pluripotency state. Confluent cultures, rough handling or higher passages may lead to unwanted spontaneous differentiation or low efficiency of kidney organoid formation.

3. Day 0 – Setting up the kidney organoid assay

1. Before starting, prepare both the E5-ILP and Stage II media as per formulations (Table 1 and Table 2).
   NOTE: The media can be stored for up to 14 days at 4 °C.
2. For one kidney organoid assay (one 100 mm culture plate is needed for one 6-well plate), prepare complete E5-ILP medium in a 50 mL conical tube: 18 mL of E5-ILP supplemented with 8 µM CHIR99021 (14.4 µL), 3.3 µM ROCKi (6 µL), 0.1 mM beta-mercaptoethanol (32.7 µL).
3. Place 2 mL of complete E5-ILP medium into each well of a 6 well ultra-low attachment plate.
4. Wash hPSCs at ~60 % confluency (Figure 2B) twice with ~ 8 mL of DPBS. Aspirate DPBS then add 2 mL of dispase per 100 mm plate, drop by drop to cover the cells and incubate for 6 min at 37 °C.
   NOTE: After 6 min, the edges of the colonies will start to curl up (Figure 2C, red arrows) while the rest of the colony remains attached. If this is not obtained after 6 min, place the cells back into the incubator for additional 30 s. Other hPSC media and matrix may not be compatible with this timing. Laminin based BME coating is not compatible with dispase. If laminin based BME are the standard hPSC matrix, coat one of the plates in section 1 with the BME described in this method to be used for the kidney organoid assay.
5. Wash cells 3x with ~10 mL of DPBS. Aspirate DPBS then tilt the plate ~45° and scrape down with a cell lifter.
   NOTE: Dispase is not deactivated, hence it needs to be washed out thoroughly. Do not reduce the number of washes.
6. Wash the colonies down from the top with 6 mL of complete E5-ILP medium using a 10 mL serological pipette. Pipette up and down gently to break up any large colonies (2 or 3 times is usually enough).
7. Distribute the colony clusters evenly by adding 1 mL per well into the 6-well plate. Place the plate on an orbital shaker (settings: orbital = 30, reciprocal = 330°, vibration = 5° – 2 s) that is placed in the 37 °C incubator (Figure 2D).
   ​NOTE: The vibration feature is important for adequate distribution of organoids and to prevent clumping.

4. Day 2 – Feeding by half-medium change

NOTE: Within the 48 h, colony clusters will form embryoid bodies.

1. Prepare the complete medium: For one 6-well plate prepare 12 mL of E5-ILP medium + 8 µM CHIR99021 in a 15 mL conical tube.
   NOTE: Beta-mercaptoethanol and ROCKi are not required.
2. Let the embryoid bodies settle at the bottom of the plate, tilt the plate ~45° then aspirate the medium slowly from the top, leave ~1 mL per well.
   NOTE: Embryoid bodies at this stage clump rapidly. Do not leave them to settle for > 5 min.
3. Add 2 mL of prepared complete medium (section 4.1) per well. Return the plate back onto the shaker.

5. Day 3 – Transfer of embryoid bodies to Stage II medium

1. Prepare a 50 mL conical tube and DMEM (low glucose). Let the embryoid bodies settle at the bottom of the plate. Tilt the plate ~45° and aspirate the medium from the top slowly, leave ~1 mL per well.
2. Collect all the embryoid bodies carefully from each well using a 10 mL serological pipette and transfer them to the 50 mL conical tube.
3. Wash each well to collect any remaining embryoid bodies with ~ 1 mL of DMEM (low glucose) and add them to the same 50 mL conical tube.
4. Leave the embryoid bodies to settle to the bottom of the tube, ~5 min. While waiting, add 2 mL of Stage II medium to each well of the 6-well plate. Seive out large embryoid bodies (>300 µm) using a 200 µm cell strainer (Figure 2E).
   1. Use a new 50 mL conical tube and place the 200 µm cell strainer on top. Pipette all of the embryoid bodies using a 10 mL serological pipette carefully over the cell strainer.
   2. Rinse the cell strainer with an additional ~5 mL of DMEM (low glucose) to collect any embryoid bodies stuck in the cell strainer. Allow the embryoid bodies to settle to bottom of the conical tube.
5. When the embryoid bodies are settled, aspirate the supernatant and wash with ~10 mL of DMEM (low glucose).
6. Aspirate DMEM and re-suspend the embryoid bodies in 6 mL of Stage II medium.
7. Transfer the embryoid bodies back into the 6 well ultra-low attachment plate, distributing them evenly among the 6 wells.
8. Carry out half medium changes as described in steps 4.2 and 4.3 every other day.
   ​NOTE: From day 3 onwards, the embryoid bodies will have a 'golden' and smooth, spherical appearance (Figure 2F). From ~ day 6, tubule formation in individual embryoid bodies will become apparent, with increasing numbers over the following days reaching optimum numbers and growth by day 14 (Figure 2G,H). To eliminate occasional clumping forming, upon visually observing the kidney organoids, or very small embryoid bodies without tubules, sieve out the <200 and large >500 µm organoids with a 500 and 200 µm cell strainers as described in steps 5.4.1 and 5.4.2.

6. Transfer to spinner flask and feeding

NOTE: A spinner flask may be used anytime from day 3 onwards for experiments that require large numbers of organoids. Routine transfer of organoids happens in our lab between days 6-8. Please see the Discussion section for alternatives if equipment is not available.

1. Transfer embryoid bodies into a 125 mL spinner flask with 45 mL of Stage II medium. Set magnetic stirrer speed to 120 rpm and place into the incubator (Figure 2I).
2. To feed embryoid bodies or kidney organoids, let the kidney organoids settle briefly to the bottom of the spinner flask. Lift the lid from one side arm of the flask and place the aspirating pipette inside, with the tip touching the opposite inside wall.
3. Slowly angle the aspirating pipette down and aspirate approximately half of the medium. Replenish with 20 mL of fresh Stage II medium by pipetting it through the same opening.

7. Setting up 6-well magnetic stir plate (6MSP)

NOTE: The 6MSP format may be used in place of spinner flasks if multiple conditions need to be tested. Use the 6MSP for compound or nephrotoxin treatments. This saves the amount of medium used in the second stage while maintaining nutrient availability through diffusion.

1. Clean the oval magnetic stir bars in a 50 mL conical tube by washing in a tissue culture suitable detergent briefly (if never used) or soak for > 1 h if previously used.
2. Briefly wash 3x in sterile DPBS.
3. Wash 1x for 5 min in 70% ethanol, 1x in sterile DPBS.
4. Rinse with anti-adherence solution and wash 1x in sterile DPBS and aspirate.
5. Carefully, using long sterile forceps place one magnetic stir bar into each well of the 6-well plate with embryoid bodies or kidney organoids.
6. Place the plate onto the 6MSP and set the speed to 120 rpm (Figure 2J). Maintain kidney organoids with half medium change as per section 4.2 and 4.3.
   NOTE: In order for the magnetic stir bars to snap into position and start spinning, you may need to first put the power level to 100 briefly, then once they are all spinning, bring the power level down to 25.