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Large-Scale Production of Kidney Organoids from Human Pluripotent Stem Cells (Version 1.0) | ATLAS-D2K Center

PLEASE NOTE: ATLAS-D2K closed July 31, 2025 and this website is for reference purposes only.

Large-Scale Production of Kidney Organoids from Human Pluripotent Stem Cells (Version 1.0)

Version

1.0

Notice

This page is the corresponding protocol tomestone page generated as part of the ATLAS-D2K shutdown in July 2025. Many links on this page may be broken.

Authors

Aneta Przepiorski; Veronika Sander; Neil Hukriede; Alan Davidson

Keywords

[‘3D culture’, ‘hiPSC’, ‘induced pluripotent stem cells (iPSC)’, ‘kidney’, ‘kidney development’, ‘organoid’, ‘nephron’, ‘Podocytes’]

Subjects

[‘Cell culture’]

Release Date

2021-08-09

Abstract

Kidney organoids represent a physiologically advanced model for studying the mechanisms of kidney development and disease. Here, we describe a simple two-step protocol for the differentiation of human pluripotent stem cells into kidney organoids. Our approach involves suspension culture that allows for rapid and cost-effective bulk production of organoids, which is well-suited for large-scale assays such as drug-screening. The organoids correspond to fetal human kidney tissue and may be of limited use for modeling adult kidney function. For complete details on the use and execution of this protocol, please refer to Przepiorski et al. (2018).

Introduction

The principle of in vitro kidney organoid differentiation is based on mimicking fetal kidney development, for which Wnt signaling is one of the key factors. Our protocol is a two-stage approach, whereby the small molecule Wnt-agonist CHIR99021 is used in the first stage medium to promote mesoderm formation and initiate nephrogenesis in form of embryoid bodies. In the second stage, the supplementation of our ‘Stage II’ medium with KnockOut Serum Replacement is sufficient to further differentiate the embryoid bodies into kidney tubule (nephron)-containing organoids. The formation of kidney organoids with our protocol is achieved without the need for additional growth factors, such as FGF9 and Activin A that are commonly used in other kidney organoid protocols. Thus, our protocol differs from other methods in that it is inexpensive, allowing kidney organoids to be grown in suspension culture in large scale. Despite the simpler approach, our organoids resemble those generated with other methods (Taguchi et al., 2014; Takasato et al., 2015; Morizane et al., 2015; Taguchi and Nishinakamura, 2017; Kumar et al., 2019) in that they develop the major renal cell types, i.e. podocytes, proximal and distal tubule cells, presumptive collecting duct cells, endothelial cells and interstitial cells. For a more detailed comparison of protocols, please refer to Przepiorski et al. (2018).

Reagents

  • 1-Thioglycerol; Sigma Cat#M6145
  • 2-Mercaptoethanol; Thermo Fisher Cat#21985023
  • Accutase; STEMCELL Technologies Cat#7920
  • AlbumiNZ™ protease reduced, immunoassay (EIA) grade, ≥97% (BSA used for immunohistochemistry); MP Biomedicals Cat#219989880
  • Chemically Defined Lipid Concentrate; Thermo Fisher Cat#11905031
  • CHIR99021; STEMCELL Technologies Cat#72054
  • Dispase; STEMCELL Technologies Cat#7923
  • DMEM, low glucose, pyruvate; Thermo Fisher Cat#11885084
  • DPBS 1x, no calcium, no magnesium; Thermo Fisher Cat#14190250
  • Geltrex™ LDEV-Free Reduced Growth Factor Basement Membrane Matrix; Thermo Fisher Cat#A1413202
  • Gentle Cell Dissociation Reagent; STEMCELL Technologies Cat#7174
  • GlutaMAX Supplement; Thermo Fisher Cat#35050061
  • Ham’s F12 Nutrient Mix; Thermo Fisher Cat#11765054
  • HEPES; Thermo Fisher Cat#15630080
  • IMDM; Thermo Fisher Cat#12440053
  • Insulin-Transferrin-Selenium-Ethanolamine (ITS -X); Thermo Fisher Cat#51500056
  • K-252a (BDNF inhibitor); Sigma Cat#K1639
  • KnockOut™ Serum Replacement - Multi-Species; Thermo Fisher Cat#A3181502
  • L-Ascorbic acid 2-phosphate sesquimagnesium salt hydrate; Sigma Cat#A8960
  • Matrigel® Growth Factor Reduced Basement Membrane Matrix, LDEV-free BD Biosciences; Cat#354230
  • MEM Non-Essential Amino Acids Solution; Thermo Fisher Cat#11140050
  • mTeSR1 STEMCELL Technologies; Cat#5850
  • Penicillin-Streptomycin; Thermo Fisher Cat#15140122
  • PFHM-II Protein-Free Hybridoma Medium; Thermo Fisher Cat#12040077
  • Plasmocin; InvivoGen Cat#ant-mpt
  • Poly(vinyl alcohol); Sigma Cat#P8136
  • Probumin® Bovine Serum Albumin; Sigma Cat#821006
  • ROCK inhibitor Y-27632; STEMCELL Technologies Cat#72304
  • StemFlex™ Medium; Thermo Fisher Cat#A3349401

Reagent preparation

hPSC culture and organoid generation The aliquot sizes suggested here correspond to the volumes required for preparing 250 mL of mBPEL medium or 500 mL of Stage II medium.
CRITICAL: Do not use a 37°C water bath for pre-warming media or other reagents. This should be done at room temperature (15–25°C). Thawing of reagents should also be done at room temperature, or overnight (16-20 hrs) at 4°C.

mTeSR1 hPSC culture medium Thaw 5x mTeSR1 Supplement then combine with 400 mL mTeSR1 Basal Medium. Add 50 µL Plasmocin (dilute 1:10,000 for a final concentration of 2.5 µg/mL) and 5 mL Penicillin-Streptomycin 100x (for final concentration of 1x). Store at 4°C for up to two weeks. Alternatives: Other hPSC culture media such as StemFlex or TeSR-E8 media can be used.

Probumin Bovine Serum Albumin (BSA) Prepare 10% w/v in IMDM medium, aliquot into 6.25 mL and store at -20°C for up to one year. The lyophilized stock powder is stable for 2 years at 4°C. Alternatives: Other low endotoxin and low IgG BSA preparations that are labeled as suitable for cell culture can be used.

L-Ascorbic acid 2-phosphate sesquimagnesium salt hydrate (AA2P) Prepare 5 mg/mL in ddH2O, aliquot into 2.5 mL and store at -20°C for up to one year.

Penicillin-Streptomycin (P/S) Prepare 2.5 or 5 mL aliquots of the 100x stock and store at -20°C for up to one year.

Poly(vinyl alcohol) (PVA) Prepare 10% w/v in 1x DPBS. Leave to dissolve in 95°C water bath (~8 hrs). Store at 4˚C for up to 6 months.

KnockOut Serum Replacement (KOSR) Prepare 50 mL aliquots of the 100x stock and store at -20°C for up to 18 months. Avoid additional freeze-thaw cycles.

Dispase Prepare 2 or 4 mL aliquots and store at -20°C until expiry date indicated on the label. Thawed aliquots are stable for up to 2 weeks at 4°C. Do not re-freeze.

Accutase Prepare 2 or 4 mL aliquots and store at -20°C until expiry date indicated on the label. Thawed aliquots are stable for up to 2 months at 4°C.

CHIR99021 Prepare 10 mM stock in DMSO, aliquot (e.g. in 50 µL) and store at -20°C for up to 6 months.

ROCK inhibitor Y-27632 Prepare 10 mM stock in ddH2O, aliquot (e.g. in 50 µL) and store at -20°C for up to one year.

Geltrex Thaw Geltrex at 4°C for 16-20 hrs. Prepare ice-cold (-20°C) pipette tips and cryotubes. Place thawed Geltrex on ice, mix carefully and aliquot (e.g. into 100 or 200 µL aliquots), then freeze immediately. Store at -20°C for up to 18 months. Note: We recommend preparing Geltrex aliquots in cryotubes with internal thread and wide-diameter opening (e.g. Nunc CryoTubes™) to facilitate sterile pipetting.

Modified Bovine Serum Albumin (BSA) Polyvinylalcohol Essential Lipids (mBPEL) medium BPEL medium was originally developed for embryoid body-based differentiation of hPSCs (Ng et al., 2008). Our modified (m)BPEL medium is a variation of the original in that it contains 0.1x ITS-X (adapted from Orlova et al., 2014).

see TABLE 1

CRITICAL: Prepare this medium by pipetting all ingredients except Lipid Concentrate and Plasmocin directly into the upper chamber of a 0.22 µm SteriCup filtration unit. Add Lipid concentrate and Plasmocin after filtration. Store at 4°C for up to two weeks.

Alternatives: Albumin Polyvinylalcohol Essential Lipids (APEL) medium, the animal-product free formulation (Ng et al., 2008) of this medium, can be used instead of BPEL.

Complete mBPEL medium Freshly prepare complete mBPEL medium on Day 0 and Day 2 of organoid generation (starting from steps 2 and 13, respectively). For one assay, add 8 µM CHIR99021 (14.4 µL), 3.3 µM ROCK inhibitor Y-27632 (6 µL) and 100 µM 2-Mercaptoethanol (32.7 µL) to 18 mL of mBPEL to make complete mBPEL medium for Day 0. Add 8 µM CHIR99021 (9.6 µL) to 12 mL of mBPEL to make complete mBPEL medium for Day 2.

Stage II medium Note: We changed from high-glucose DMEM (as used in the original protocol) to low-glucose DMEM. This modification was made as the low-glucose DMEM formulation (containing 5.5 mM glucose) more closely resembles physiological fasting glucose levels of 5.5 mM, whereas high-glucose DMEM (containing 25 mM glucose) exposes the organoids to hyperglycemia.

see TABLE 2

CRITICAL: Prepare this medium by pipetting all ingredients except Plasmocin directly into the upper chamber of a 0.22 µm SteriCup filtration unit. Add Plasmocin after filtration. Store at 4°C for up to two weeks.

Equipment

  • Cell lifter; Corning Cat#3008
  • Corning tissue-culture treated culture dishes, D × H 100 mm × 20 mm; Merck Millipore Cat#430167
  • Corning disposable spinner flasks; Sigma Cat#CLS3152
  • Fisherbrand™ Multi Function 3D rotator; Fisher Scientific Cat#15514080
  • pluriStrainer® 500 µm; Pluri Select Cat#43-50500-03
  • Stericup-GP Sterile Vacuum Filtration System (500 mL); Merck Millipore Cat#S2GPU05RE
  • Stericup-GP Sterile Vacuum Filtration System (200 mL); Merck Millipore Cat#SCGPU02RE
  • Ultra-Low Attachment 24-well plates; Merck Millipore Cat#CLS3473
  • Ultra-Low Attachment 6-well plates Merck Millipore; Cat#CLS3471

Procedure

Passaging hPSCs (Day -2)

Timing: 20 min

This split is performed to achieve 40-50% confluency at Day 0 of the organoid generation protocol. In the original publication of this protocol, we determined the average cell number in a 40-50% confluent 100 mm culture of MANZ-2-2 iPSCs to be 6.6E+06 ± 2.3E+05 (Przepiorski et al., 2018).

  1. Passage hPSCs into one or more 100 mm Geltrex-coated culture dishes as described above. Splitting the cultures at 1:4–1:6 usually leads to the desired confluency after two (maximum three) days.

Starting the organoid assay (Day 0)

Timing: 30 min

This step describes how hPSC colonies are prepared for differentiation. Our protocol starts with a 3-day treatment with 8 µM CHIR99021 to induce mesoderm formation in form of spherical embryoid bodies in suspension culture (Figure 2A). The volumes given are for one assay that is initiated from one 100 mm culture dish.

  1. Check for 40-50% confluency of the culture (Figure 2B). The colonies should be mostly discrete.
    Note: If the optimal confluency is missed, we recommend splitting the culture and starting organoid differentiation with the subsequent passage.
  2. Pre-warm mBPEL medium to room temperature (15–25°C) and thaw an aliquot of Dispase.
  3. Prepare 18 mL of complete mBPEL medium in a 50 mL centrifuge tube (add 14.4 µL CHIR99021, 6 µL ROCK inhibitor Y-27632 and 32.7 µL 2-Mercaptoethanol to 18 mL mBPEL). Mix well by inverting the tube.
  4. Aliquot 2 mL of complete mBPEL medium into each well of an ultra-low attachment 6-well plate.
  5. Wash hPSCs twice with 10 mL 1x DPBS.
  6. Aspirate DPBS and add 2 mL of Dispase dropwise (ensuring that all colonies are covered) then incubate at 37°C for 6 min. Upon this treatment, the edges of the hPSC colonies should start to curl up but not detach from the dish (Figure 2C). Note: Depending the type of culture medium used, this timing may need to be adjusted. CAUTION: Over-digestion will lead to detachment of the colonies, decreased cell survival and low yield of embryoid body formation (Figure 2D).
  7. Aspirate Dispase, then wash 3x with 10 mL 1x DPBS each. CRITICAL: The 3 washes are crucial to completely remove Dispase. Insufficient washing may lead to cell death or clumping of embryoid bodies.
  8. After the last wash, aspirate DPBS and scrape the colonies off the culture dish using a cell lifter.
  9. Use a 10 mL serological pipette to wash down the colonies with the remaining 6 mL of complete mBPEL. Pipette up and down (3-5x) to break up large colonies into evenly sized fragments of ~ 100 µm, then distribute 1 mL to each well of the 6-well plate (Figure 2E). Troubleshooting 2 (Figure 4B)
  10. Place the 6-well plate into a 37°C incubator, move side-to-side for even distribution of the colonies.
  11. Leave the plate undisturbed for 48 hours.

Half-medium change (Day 2)

Timing: 10 min

On Day 2, a half-medium change is performed to supply fresh mBPEL and CHIR99021.

  1. Pre-warm mBPEL to room temperature (15–25°C), then prepare 12 mL of complete mBPEL (with 9.6 µL CHIR99021 added).
  2. Check the 6-well plate for embryoid body formation. At this stage, the embryoid bodies should be obvious, appearing as spheres of 50 – 150 µm in diameter among floating single cells (Figure 2F and F’). We use an EVOS™ XL Core inverted Imaging System to monitor embryoid body (and organoid) appearance.
  3. Place 6-well plate into the tissue culture hood and tilt to achieve a ~45° angle (e.g. place on the edge of a tube rack, Figure 2G). Let embryoid bodies sediment on the bottom (3-5 min). CAUTION: Do not leave the embryoid bodies in this position for extended time as clumping will occur. Troubleshooting 3 (Figure 4F)
  4. Carefully aspirate approximately half of the medium from each 6-well.
  5. Return 6-well plate into flat position and add 2 mL of complete mBPEL to each well.
  6. Place 6-well plate on an orbital shaker in the incubator (Figure 2H).
    CRITICAL: We recommend using an orbital shaker that can be set to orbital and reciprocal agitation, as well as vibration. We use the following settings on a Fisherbrand Multi Function 3D Rotator: Orbital rotation at 30 rpm for 10 sec; Reciprocal rotation for 10 sec including 2 changes of direction; Vibration for 5 sec.

Transfer to Stage II medium (Day 3)

Timing: 20 min

On Day 3, mBPEL is replaced by Stage II medium. Stage II medium contains 15% KnockOut Serum Replacement in DMEM. This is sufficient to drive renal tubule formation, which becomes apparent by Day 7 - Day 9 of the protocol.

  1. Pre-warm DMEM and Stage II media to room temperature (15–25°C).
  2. Check appearance of Day 3 embryoid bodies. The embryoid bodies should be spherical, light-golden in color with smooth edges (Figure 2I and I’). Troubleshooting 4
  3. Aspirate embryoid bodies from all 6 wells using a 10 or 25 mL serological pipette and combine in a 50 mL centrifuge tube.
  4. Rinse the 6 wells with 1 mL DMEM per well to collect remaining embryoid bodies then add these to the 50 mL tube.
  5. Leave the embryoid bodies to sediment at the bottom of the tube (3-5 min) at room temperature.
  6. Carefully aspirate the supernatant then wash with 15 mL of DMEM. Repeat step 23. CAUTION: Do not leave the embryoid bodies accumulate at the bottom of the tube for extended time as clumping will occur. Troubleshooting 3 (Figure 4F)
  7. Aliquot 2 mL of Stage II medium into each well of the 6-well plate.
  8. Carefully aspirate the supernatant from the sedimented embryoid bodies.
  9. Take up the embryoid bodies in 6 mL of Stage II medium and redistribute into the 6-well plate, 1 mL per well. CAUTION: Distribute embryoid bodies evenly as high density of embryoid bodies in a 6-well can affect organoid formation. Troubleshooting 5
  10. Return plate to the orbital shaker.

Alternatives: Instead of continuing to culture the embryoid bodies in the 6-well plate format, they can be transferred into a spinning bioreactor, e.g. the Corning spinner flask (Figure 2J) from Day 3 onwards.

27a. Take up embryoid bodies in Stage II medium and transfer into a spinner flask, then fill up to 45 mL with Stage II medium. 28a. Place on magnetic stirrer (at 90 rpm) in the incubator (Figure 2J).

Optional: Size-filtration of embryoid bodies / organoids

Timing: 10 min

Occasional fusion may lead to abnormally large embryoid bodies and organoids. (Troubleshooting 3). We have observed that organoids exceeding a diameter of 700 µm display signs of core apoptosis and necrosis, perhaps due to insufficient levels of oxygen and/or nutrients (Przepiorski et al., 2018). To eliminate large specimens and achieve a uniformly-sized organoid culture, size-filtration can be performed from Day 3 onwards.

  1. Place a 500 µm cell strainer onto a 50 mL centrifugation tube (Figure 4E).
  2. Take up embryoid bodies / organoids with a 10 or 25 mL serological pipette and slowly drop onto the cell strainer then rinse with 5 mL Stage II medium.
  3. Leave embryoid bodies / organoids in the flow-through to sediment, then re-distribute into 6-well plate or transfer into spinner flask.

Maintaining embryoid body cultures (Day 3 – Day 8)

Timing: 10 min

From Day 3 onwards, Stage II medium needs to be changed every 2-3 days.

  1. Pre-warm Stage II medium to room temperature (15–25°C).
  2. Remove spinner flask or 6-well plate from incubator, leave embryoid bodies to sediment (in tilted position for 6-well plates) at room temperature.
  3. Carefully aspirate most of the medium from each 6-well and roughly half of the medium from a spinner flask.
  4. Replace with 2 - 3 mL fresh Stage II medium per 6-well, or adjust to 45 mL per spinner flask.

Note: For spinner flasks, it is sufficient to perform medium changes every 3 days. For 6-well plates, the amount of Stage II medium per well can be adjusted to the frequency of medium changes, i.e. add 3 mL when changing every 3 days (e.g. over the weekend), add 2 mL when changing every other day.

Maintaining kidney organoid cultures (Day 8 – Day 20+)

Timing: 10 min

Between Day 7 and Day 9, the outlines of the kidney tubules should become visible on the surface of the embryoid bodies (here forth referred to as kidney organoids; Figure 2K) when observed by bright-field microscopy. (Troubleshooting 6).

  1. Keep changing Stage II medium every 2-3 days as described in steps 32-35.

The organoids will reach their optimal state of differentiation by Day 12 (Figure 2L), when gene expression for specific markers of the renal cell types (i.e. podocytes, proximal tubule cells, distal tubule and collecting duct cells) can be detected by quantitative (q)PCR, and marker proteins are detectable by immunohistochemistry (Figure 3; Przepiorski et al., 2018). Validated oligonucleotide primers and antibodies recommended for determining optimal organoid maturity are listed in the Key Resources Table. The organoids maintain expression of these gene and protein markers through to approximately Day 20. We recommend performing assays, such as drug treatments, during this period (Digby et al., 2020).

PAUSE POINT: For gene expression analysis, wash organoids once in 1x DPBS and then transfer into TRIzol reagent. Store at -80°C until RNA isolation, cDNA synthesis and qPCR analysis. For detailed protocols and expected results, please refer to our previous publications (Przepiorski et al., 2018; Digby et al., 2020). For immunohistochemistry, fix whole organoids in 1 mL 4% Paraformaldehyde for 20 min at room temperature (15–25°C), then wash once with 1x DPBS and store at 4°C until further processing (for up to two weeks).

Timing

Timeline of the protocol

Critical_Steps

CRITICAL: Freshly thawed hPSC lines should be passaged at least once and cultured for a minimum of one week before starting organoid generation. This allows the hPSCs to adapt to culture procedures and establish a regular growth pattern. Depending on the hPSC line, we observed that the efficiency of organoid formation may improve after ~2 weeks of culture and >2 passages. (Troubleshooting 1)

CRITICAL: Do not use a 37°C water bath for pre-warming media or other reagents. This should be done at room temperature (15–25°C). Thawing of reagents should also be done at room temperature, or overnight (16-20 hrs) at 4°C.

CRITICAL: On day 0 of organoid generation, over-digestion with Dispase will lead to detachment of the colonies, decreased cell survival and low yield of embryoid body formation (Figure 2D).

CRITICAL: On day 0, the 3 washes are crucial to completely remove Dispase. Insufficient washing may lead to cell death or clumping of embryoid bodies.

CRITICAL: On day 2 for half medium change, do not leave the embryoid bodies in the tilted position for extended time as clumping will occur. Troubleshooting 3 (Figure 4F). This also applies to the transfer to stage II medium on day 3 and every medium change thereafter.

CRITICAL: From day 2 on, we recommend using an orbital shaker that can be set to orbital and reciprocal agitation, as well as vibration. We use the following settings on a Fisherbrand Multi Function 3D Rotator: Orbital rotation at 30 rpm for 10 sec; Reciprocal rotation for 10 sec including 2 changes of direction; Vibration for 5 sec.

CRITICAL: On day 3, distribute embryoid bodies evenly as high density of embryoid bodies in a 6-well can affect organoid formation. Troubleshooting 5

Trouble_Shooting

Troubleshooting

Problem 1:

Low-efficiency organoid formation (Figure 4A).

Potential Solution:

Using bright-field microscopy, specimens without tubules are clearly distinguishable from tubule-containing organoids by Day 8 or Day 9 of the protocol. Low efficiency (<50%) of tubule formation is most likely due to sub-optimally maintained hPSC cultures. It is most critical that cultures are passaged regularly and before reaching 80% confluency, as high-density cultures can lead to spontaneous differentiation and low-yield differentiation assays. Do not start organoid differentiation if an hPSC culture has become over-confluent. Instead, passage the cells twice, then resume with organoid differentiation. Furthermore, ensure that ROCK inhibitor is added when passaging, the split ratio is not too high (we do not recommend higher than 1:8) and the cultures are negative for mycoplasma.

Problem 2:

Unevenly-sized embryoid bodies / organoids (Figure 4B).

Potential Solution:

Avoid excessive trituration of Dispase-treated colonies (step 10), as cell aggregates smaller than ~50 µm are less likely to form tubular organoids. On the other hand, aggregates >200 µm will likely develop into large embryoid bodies / organoids that may form apoptotic or necrotic cores (Przepiorski et al., 2018).

Problem 3:

Clumping of embryoid bodies (Figure 4C-F).

Potential Solution:

Occasional merging of two or more embryoid bodies can occur during the first 48 hrs of the protocol (Figure 4C). This can result in large specimens (Figure 4D) that can be removed by size-filtration (steps 29-31; Figure 4E). To prevent severe clumping of embryoid bodies, ensure that the cultures are not left to sediment for more than 5 min, e.g. during half-medium change at Day 2 (step 15), transfer into Stage II medium at Day 3 (step 23) or medium changes at later stages of the protocol. Severe clumping will also occur if the orbital shaker or spinner flask is accidentally left switched off (Figure 4F).

Problem 4:

Sub-ideal appearance of embryoid bodies.

Potential Solution:

Based on our experience, embryoid bodies that appear bean-shaped, dark-colored or exhibit rough edges by Day 5 – Day 7 of the protocol are unlikely to develop into tubular kidney organoids. We recommend discontinuing such assays and instead starting over, using healthy, non-differentiating hPSC cultures. Non-spherical shaped embryoid bodies can be a result of differentiating hPSCs. See also Problem 1.

Problem 5:

High density of embryoid bodies / organoids.

Potential Solution:

The optimal number of embryoid bodies or organoids per 6-well is ≤150. If too dense, distribute into several 6-well plates or use a spinner flask, which can hold up to 6,250 organoids (Przepiorski et al., 2018). For short-term culture up to 72 hrs, e.g. drug treatments after Day 12, ~100 organoids can be cultured per 24-well (as shown in Figure 3A, B).

Problem 6:

Delayed tubule formation, or inconsistent maturation of organoids generated from different (isogenic) lines.

Potential Solution:

This could be due to intrinsic differences between hPSC lines. If observed consistently, refer to Phipson et al., 2019 and check organoid maturation state by qPCR or RNA-sequencing.

Problem 7:

Outgrowth of non-renal cells (Figure 4G).

Potential Solution:

We observed that some hPSC lines trend towards forming large areas of non-renal cells, including cells of neuronal and myofibroblast appearance, when cultured >20 days. For example, the CRL1502 (clone C32) iPSC line shows a tendency towards neuronal differentiation, which is also noticeable by single cell RNA sequencing (Wu et al., 2018). These organoids are usually large and can be removed by size-filtration (steps 29-31). Alternatively, organoid cultures could be treated with K-252a, an inhibitor of neuronal differentiation, as described in Wu et al. (2018). We also observed that more stringent hPSC culture conditions (e.g. using TeSR-E8 medium instead of mTeSR1) helped with reducing non-renal outgrowths on organoids derived from some hPSC lines. emphasized text

Anticipated_Results

This protocol produces kidney organoids within less than two weeks. An assay starting from a 100 mm culture dish of hPSCs typically yields 500-1,000 organoids (Figure 3A, B, showing a highly efficient assay of >1,000 Day 12 organoids distributed into 10 wells of a 24-well plate for compound testing). Organoid efficiency, i.e. the number of organoids that contain nephron-like structures within one assay, is typically >90% of the total number of organoids (Figure 3C; Troubleshooting 1). Our protocol has successfully been applied to >30 human induced pluripotent stem cell and embryonic stem cell lines, including isogenic lines. Commercially available and published hPSC lines are listed in the Key Resources Table. Organoid production with this protocol is very robust, i.e. no adjustments in CHIR99021 concentration, media composition or treatment periods are required to achieve the above-mentioned high efficiency of organoid formation.

Immunostainings on paraffin sections confirm the presence of the main kidney tissues in the organoids, i.e. nephrons and their sub-segmentation into podocyte clusters (resembling primitive glomeruli), proximal tubules, distal tubules and connecting/collecting duct epithelia. Figure 3D shows representative images of organoid sections at Day 12 with WT1-labelled podocytes, EPCAM and HNF1B-labelled tubules, CUBN- and LRP2-labeled proximal tubules and CDH1-labeled distal tubules and connective/collecting duct epithelia. For more examples of immunohistologically labeled organoids, please refer to our previous publications (Przepiorski et al., 2018; Digby et al., 2020).

Limitations

Comparative analyses with fetal human kidney tissue revealed that the maturity of our kidney organoids corresponds to the ‘late capillary loop’ stage of first trimester human kidneys (Przepiorski et al. 2018). As such, markers of nephron segments that form at later stages of fetal kidney development are not detectable. The fetal-like state of the organoid kidney tissues may limit studies that aim to address structural aspects and functions of fully differentiated human kidneys, as well as adult-onset disease modeling. Nonetheless, our kidney organoids (and organoids with comparable maturity generated with other protocols) have been shown to recapitulate congenital kidney diseases (Forbes et al., 2018; Tanigawa et al., 2018; Combes et al., 2019; Przepiorski et al., 2018; Freedman et al., 2015) and acute kidney injury (Morizane et al., 2015; Hale et al., 2019; Digby et al., 2020). As an alternative to the fetal-like kidney tissues derived from hPSCs, organoids can be generated from adult human kidney biopsies or urine-derived epithelial cells (Schutgens et al., 2019; Jun et al., 2018). These so-called tubuloids provide a rapidly achievable and stable system that could be more suitable for personalized disease-modeling of e.g. kidney malignancies. However, in contrast to hPSC-derived organoids that are comprised of multiple renal cell types and reflect the structural complexity of the kidney, tubuloids only contain epithelial cells that grow in a tubulocystic configuration, thus have their own limitations.

Extended culture (>20 days) of kidney organoids generated with our protocol may lead to expansion of non-renal cell types, including neuronal cells and myofibroblasts (Figure 4G; Troubleshooting 7), leaving the organoids less representative of healthy human kidney tissue. This limits the optimal time frame for experiments on the organoids to ~Day 10 – Day 20.

References

please find the full list of references in the original publication https://star-protocols.cell.com/protocols/229

Associated_Publications

  • Przepiorski, A., Sander, V., Tran, T., Hollywood, J.A., Sorrenson, B., Shih, J.-H., Wolvetang, E.J., McMahon, A.P., Holm, T.M., and Davidson, A.J. (2018). A Simple Bioreactor-Based Method to Generate Kidney Organoids from Pluripotent Stem Cells. Stem Cell Reports 11, 470-484.
  • Digby, J.L.M., Vanichapol, T., Przepiorski, A., Davidson, A.J., and Sander, V. (2020). Evaluation of cisplatin-induced injury in human kidney organoids. Am J Physiol Renal Physiol 318, F971-F978.
  • Sander V, Przepiorski A, Crunk AE, Hukriede NA, Holm TM, Davidson AJ. Protocol for Large-Scale Production of Kidney Organoids from Human Pluripotent Stem Cells. STAR Protoc. 2020 Oct 29;1(3):100150. doi: 10.1016/j.xpro.2020.100150.
  • Przepiorski A, Crunk AE, Holm TM, Sander V, Davidson AJ, Hukriede NA. A Simplified Method for Generating Kidney Organoids from Human Pluripotent Stem Cells. J Vis Exp. 2021 Apr 13;(170). doi: 10.3791/62452.

Acknowledgement

We thank G. Chang and T. Perreau for critical reading of the manuscript and L. Conrad for helpful input on the troubleshooting. This work was supported by the Health Research Council of New Zealand (17/425), Auckland Medical Research Foundation (1116018), Cystinosis Research Foundation USA, Cystinosis Research Ireland Foundation (MRCG2014-8), the National Institute of Diabetes and Digestive and Kidney Diseases (R01-DK-069403), the United States Army Medical Research and Development Command (W81XWH-17-1-0610), and Valrae Collins Philanthropic support for AP.

Consortium

(Re)Building a Kidney (RBK) Consortium