Abstract

Adult stem cell-derived organoids are three-dimensional epithelial structures that recapitulate fundamental aspects of their organ of origin. We describe conditions for the long-term growth of primary kidney tubular epithelial organoids, or ‘tubuloids’. The cultures are established from human and mouse kidney tissue and can be expanded for at least 20 passages (>6 months) while retaining a normal number of chromosomes. In addition, cultures can be established from human urine. Human tubuloids represent proximal as well as distal nephron segments, as evidenced by gene expression, immunofluorescence and tubular functional analyses. We apply tubuloids to model infectious, malignant and hereditary kidney diseases in a personalized fashion. BK virus infection of tubuloids recapitulates in vivo phenomena. Tubuloids are established from Wilms tumors. Kidney tubuloids derived from the urine of a subject with cystic fibrosis allow ex vivo assessment of treatment efficacy. Finally, tubuloids cultured on microfluidic organ-on-a-chip plates adopt a tubular conformation and display active (trans-)epithelial transport function.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Data availability

The data that support the findings of this study are available from the corresponding author upon reasonable request. WGS data have been deposited at the EGA, which is hosted by the EBI, under accession code EGAS00001002729. Single-cell and bulk sequencing data can be found in the Supplementary Information and have been deposited at GEO under accession code GSE107795.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  1. 1.

    Clevers, H. Modeling development and disease with organoids. Cell 165, 1586–1597 (2016).

  2. 2.

    Drost, J. & Clevers, H. Organoids in cancer research. Nat. Rev. Cancer 18, 407–418 (2018).

  3. 3.

    Takasato, M. et al. Kidney organoids from human iPS cells contain multiple lineages and model human nephrogenesis. Nature 526, 564–568 (2015).

  4. 4.

    Sato, T. et al. Single Lgr5 stem cells build crypt-villus structures in vitro without a mesenchymal niche. Nature 459, 262–265 (2009).

  5. 5.

    Dekkers, J. F. et al. A functional CFTR assay using primary cystic fibrosis intestinal organoids. Nat. Med. 19, 939–945 (2013).

  6. 6.

    Dekkers, J. F. et al. Characterizing responses to CFTR-modulating drugs using rectal organoids derived from subjects with cystic fibrosis. Sci. Transl. Med. 8, 344ra384–344ra384 (2016).

  7. 7.

    Sato, T. et al. Long-term expansion of epithelial organoids from human colon, adenoma, adenocarcinoma, and Barrett’s epithelium. Gastroenterology 141, 1762–1772 (2011).

  8. 8.

    Lancaster, M. A. et al. Impaired Wnt-[beta]-catenin signaling disrupts adult renal homeostasis and leads to cystic kidney ciliopathy. Nat. Med. 15, 1046–1054 (2009).

  9. 9.

    Adams, D. C. et al. Follistatin-like 1 regulates renal IL-1beta expression in cisplatin nephrotoxicity. Am. J. Physiol. Renal Physiol. 299, F1320–F1327 (2010).

  10. 10.

    Poladia, D. P. et al. Role of fibroblast growth factor receptors 1 and 2 in the metanephric mesenchyme. Develop. Biol. 291, 325–339 (2006).

  11. 11.

    Cancilla, B., Davies, A., Cauchi, J. A., Risbridger, G. P. & Bertram, J. F. Fibroblast growth factor receptors and their ligands in the adult rat kidney. Kidney Int. 60, 147–155 (2001).

  12. 12.

    Xu, J., Lamouille, S. & Derynck, R. TGF-β-induced epithelial to mesenchymal transition. Cell Res. 19, 156–172 (2009).

  13. 13.

    Watanabe, K. et al. A ROCK inhibitor permits survival of dissociated human embryonic stem cells. Nat. Biotechnol. 25, 681–686 (2007).

  14. 14.

    Janda, C. Y. et al. Surrogate Wnt agonists that phenocopy canonical Wnt and β-catenin signalling. Nature 545, 234–237 (2017).

  15. 15.

    Boj, S. F. et al. Organoid models of human and mouse ductal pancreatic cancer. Cell 160, 324–338 (2015).

  16. 16.

    Baker, D. E. C. et al. Adaptation to culture of human embryonic stem cells and oncogenesis in vivo. Nat. Biotechnol. 25, 207–215 (2007).

  17. 17.

    Cheng, L. et al. Low incidence of DNA sequence variation in human induced pluripotent stem cells generated by nonintegrating plasmid expression. Cell Stem Cell 10, 337–344 (2012).

  18. 18.

    Cassio, D. Long term culture of MDCK strains alters chromosome content. BMC Res. Notes 6, 162 (2013).

  19. 19.

    Olofsson, B. et al. Vascular endothelial growth factor B (VEGF-B) binds to VEGF receptor-1 and regulates plasminogen activator activity in endothelial cells. Proc. Natl Acad. Sci. USA 95, 11709–11714 (1998).

  20. 20.

    Brunskill, E. W. et al. Atlas of gene expression in the developing kidney at microanatomic resolution. Develop. Cell 15, 781–791 (2008).

  21. 21.

    Birukov, K. G. et al. Stretch affects phenotype and proliferation of vascular smooth muscle cells. Mol. Cell. Biochem. 144, 131–139 (1995).

  22. 22.

    LeBleu, V. S. et al. Origin and function of myofibroblasts in kidney fibrosis. Nat. Med. 19, 1047–1053 (2013).

  23. 23.

    Ozcan, A. et al. PAX 8 expression in non-neoplastic tissues, primary tumors, and metastatic tumors: a comprehensive immunohistochemical study. Mod. Pathol. 24, 751–764 (2011).

  24. 24.

    Kang, H. M. et al. Sox9-positive progenitor cells play a key role in renal tubule epithelial regeneration in mice. Cell Rep. 14, 861–871 (2016).

  25. 25.

    Bussolati, B. et al. Isolation of renal progenitor cells from adult human kidney. Am. J. Pathol. 166, 545–555 (2005).

  26. 26.

    Barker, N. et al. Lgr5(+ve) stem/progenitor cells contribute to nephron formation during kidney development. Cell Rep. 2, 540–552 (2012).

  27. 27.

    Kobayashi, A. et al. Six2 defines and regulates a multipotent self-renewing nephron progenitor population throughout mammalian kidney development. Cell Stem Cell 3, 169–181 (2008).

  28. 28.

    Breiderhoff, T. et al. Deletion of claudin-10 (Cldn10) in the thick ascending limb impairs paracellular sodium permeability and leads to hypermagnesemia and nephrocalcinosis. Proc. Natl Acad. Sci. USA 109, 14241–14246 (2012).

  29. 29.

    Dimke, H. et al. Activation of the Ca 2+-sensing receptor increases renal claudin-14 expression and urinary Ca2+ excretion. Am. J. Physiol. Renal Physiol 304, F761–F769 (2013).

  30. 30.

    Ferrè, S. et al. Mutations in PCBD1 cause hypomagnesemia and renal magnesium wasting. J. Am. Soc. Nephrol. 25, 574–586 (2014).

  31. 31.

    De Baaij, J. H. et al. Identification of SLC41A3 as a novel player in magnesium homeostasis. Sci. Rep. 6, 28565 (2016).

  32. 32.

    Kriz, W. & Lehir, M. Pathways to nephron loss starting from glomerular diseases—insights from animal models. Kidney Int. 67, 404–419 (2005).

  33. 33.

    Scialdone, A. et al. Computational assignment of cell-cycle stage from single-cell transcriptome data. Methods 85, 54–61 (2015).

  34. 34.

    van den Brink, S. C. et al. Single-cell sequencing reveals dissociation-induced gene expression in tissue subpopulations. Nat. Method 14, 935 (2017).

  35. 35.

    Karafin, M. et al. Diffuse expression of PAX2 and PAX8 in the cystic epithelium of mixed epithelial stromal tumor, angiomyolipoma with epithelial cysts, and primary renal synovial sarcoma: evidence supporting renal tubular differentiation. Am.J. Surg. Pathol. 35, 1264–1273 (2011).

  36. 36.

    Chen, G. New advances in urea transporter UT-A1 membrane trafficking. Int. J. Mol. Sci. 14, 10674–10682 (2013).

  37. 37.

    Kirk, A., Campbell, S., Bass, P., Mason, J. & Collins, J. Differential expression of claudin tight junction proteins in the human cortical nephron. Nephrol. Dial. Transplant. 25, 2107–2119 (2010).

  38. 38.

    Rudnicki, M. et al. Hypoxia response and VEGF-A expression in human proximal tubular epithelial cells in stable and progressive renal disease. Lab. Invest. 89, 337–346 (2009).

  39. 39.

    Huch, M. et al. Long-term culture of genome-stable bipotent stem cells from adult human liver. Cell 160, 299–312 (2015).

  40. 40.

    Jansen, J. et al. A morphological and functional comparison of proximal tubule cell lines established from human urine and kidney tissue. Exp. Cell Res. 323, 87–99 (2014).

  41. 41.

    Barker, N., Van Oudenaarden, A. & Clevers, H. Identifying the stem cell of the intestinal crypt: strategies and pitfalls. Cell Stem Cell 11, 452–460 (2012).

  42. 42.

    Ganguly, N. et al. Low-dose cidofovir in the treatment of symptomatic BK virus infection in patients undergoing allogeneic hematopoietic stem cell transplantation: a retrospective analysis of an algorithmic approach. Transpl. Infect. Dis. 12, 406–411 (2010).

  43. 43.

    Cundy, K. C. et al. Clinical pharmacokinetics of cidofovir in human immunodeficiency virus-infected patients. Antimicrob. Agents Chemother. 39, 1247–1252 (1995).

  44. 44.

    Kalapurakal, J. A. et al. Management of Wilms’ tumour: current practice and future goals. Lancet Oncol. 5, 37–46 (2004).

  45. 45.

    Rivera, M. N. & Haber, D. A. Wilms’ tumour: connecting tumorigenesis and organ development in the kidney. Nat. Rev. Cancer 5, 699–712 (2005).

  46. 46.

    Hohenstein, P., Pritchard-Jones, K. & Charlton, J. The yin and yang of kidney development and Wilms’ tumors. Genes Develop. 29, 467–482 (2015).

  47. 47.

    Hawthorn, L. & Cowell, J. K. Analysis of wilms tumors using SNP mapping array-based comparative genomic hybridization. PloS ONE 6, e18941 (2011).

  48. 48.

    Wegert, J. et al. Mutations in the SIX1/2 pathway and the DROSHA/DGCR8 miRNA microprocessor complex underlie high-risk blastemal type Wilms tumors. Cancer Cell 27, 298–311 (2015).

  49. 49.

    Hing, S. et al. Gain of 1q is associated with adverse outcome in favorable histology Wilms’ tumors. Am. J. Pathol. 158, 393–398 (2001).

  50. 50.

    Mengelbier, L. H. et al. Deletions of 16q in Wilms tumors localize to blastemal-anaplastic cells and are associated with reduced expression of the IRXB renal tubulogenesis gene cluster. Am. J. Pathol. 177, 2609–2621 (2010).

  51. 51.

    Zhou, T. et al. Generation of human induced pluripotent stem cells from urine samples. Nat. Protoc. 7, 2080–2089 (2012).

  52. 52.

    Dekkers, J. F. et al. Potentiator synergy in rectal organoids carrying S1251N, G551D, or F508del CFTR mutations. J. Cyst. Fibros. 15, 568–578 (2016).

  53. 53.

    Homan, K. A. et al. Bioprinting of 3D convoluted renal proximal tubules on perfusable chips. Sci. Rep. 6, 34845 (2016).

  54. 54.

    Trietsch, S. J., Israëls, G. D., Joore, J., Hankemeier, T. & Vulto, P. Microfluidic titer plate for stratified 3D cell culture. Lab Chip 13, 3548–3554 (2013).

  55. 55.

    Trietsch, S. J. et al. Membrane-free culture and real-time barrier integrity assessment of perfused intestinal epithelium tubes. Nature Commun. 8, 262 (2017).

  56. 56.

    Jouan, E., Vee, M., Denizot, C., Da Violante, G. & Fardel, O. The mitochondrial fluorescent dye rhodamine 123 is a high‐affinity substrate for organic cation transporters (OCTs) 1 and 2. Fundam. Clin. Pharmacol. 28, 65–77 (2014).

  57. 57.

    Yumoto, R. et al. Transport of rhodamine 123, a P-glycoprotein substrate, across rat intestine and Caco-2 cell monolayers in the presence of cytochrome P-450 3A-related compounds. J. Pharmacol. Exp. Ther. 289, 149–155 (1999).

  58. 58.

    Hirsch, H. H. et al. Polyomavirus-associated nephropathy in renal transplantation: interdisciplinary analyses and recommendations. Transplantation 79, 1277–1286 (2005).

  59. 59.

    de Kort, H. et al. Primary human renal-derived tubular epithelial cells fail to recognize and suppress BK virus infection. Transplantation 101, 1820–1829 (2017).

  60. 60.

    Cakalagaoglu, F., Erbarut, I. & Tuglular, S. Frequency of BK virus nephropathy in graft dysfunction biopsies. Dialysis Transplant. 36, 122–126 (2007).

  61. 61.

    Bohl, D. L. & Brennan, D. C. BK virus nephropathy and kidney transplantation. Clin. J. Am. Soc. Nephrol. 2, S36–S46 (2007).

  62. 62.

    Royer-Pokora, B. et al. Wilms tumor cells with WT1 mutations have characteristic features of mesenchymal stem cells and express molecular markers of paraxial mesoderm. Hum. Mol. Genet. 19, 1651–1668 (2010).

  63. 63.

    Takasato, M. et al. Directing human embryonic stem cell differentiation towards a renal lineage generates a self-organizing kidney. Nat. Cell Biol. 16, 118–126 (2014).

  64. 64.

    Takasato, M. et al. Kidney organoids from human iPS cells contain multiple lineages and model human nephrogenesis. Nature 526, 564–568 (2015).

  65. 65.

    Freedman, B. S. et al. Modelling kidney disease with CRISPR-mutant kidney organoids derived from human pluripotent epiblast spheroids. Nat. Commun. 6, 8715 (2015).

  66. 66.

    Lam, A. Q. et al. Rapid and efficient differentiation of human pluripotent stem cells into intermediate mesoderm that forms tubules expressing kidney proximal tubular markers. J. Am. Soc. Nephrol. 25, 1211–1225 (2014).

  67. 67.

    Gutierrez-Aranda, I. et al. Human induced pluripotent stem cells develop teratoma more efficiently and faster than human embryonic stem cells regardless the site of injection. Stem Cells 28, 1568–1570 (2010).

  68. 68.

    Clevers, H. The intestinal crypt, a prototype stem cell compartment. Cell 154, 274–284 (2013).

  69. 69.

    Kim, K.-A. et al. Mitogenic influence of human R-Spondin1 on the intestinal epithelium. Science 309, 1256–1259 (2005).

  70. 70.

    Huch, M. et al. In vitro expansion of single Lgr5+liver stem cells induced by Wnt-driven regeneration. Nature 494, 247–250 (2013).

  71. 71.

    Drost, J. et al. Use of CRISPR-modified human stem cell organoids to study the origin of mutational signatures in cancer. Science 358, 234–238 (2017).

  72. 72.

    Barker, N. et al. Lgr5(+ve) stem cells drive self-renewal in the stomach and build long-lived gastric units in vitro. Cell Stem Cell 6, 25–36 (2010).

  73. 73.

    Muraro, M. J. et al. A single-cell transcriptome atlas of the human pancreas. Cell Syst 3, 385–394 (2016).

  74. 74.

    Hashimshony, T. et al. CEL-Seq2: sensitive highly-multiplexed single-cell RNA-Seq. Genome Biol. 17, 77 (2016).

  75. 75.

    Li, H. & Durbin, R. Fast and accurate short read alignment with Burrows–Wheeler transform. Bioinformatics 25, 1754–1760 (2009).

  76. 76.

    Satija, R., Farrell, J. A., Gennert, D., Schier, A. F. & Regev, A. Spatial reconstruction of single-cell gene expression data. Nat. Biotechnol. 33, 495–502 (2015).

  77. 77.

    Tirosh, I. et al. Dissecting the multicellular ecosystem of metastatic melanoma by single-cell RNA-seq. Science 352, 189–196 (2016).

  78. 78.

    Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014).

  79. 79.

    Galmes, R. et al. Vps33B is required for delivery of endocytosed cargo to lysosomes. Traffic 16, 1288–1305 (2015).

  80. 80.

    Tan, L. et al. Genetic variability among complete human respiratory syncytial virus subgroup A genomes: bridging molecular evolutionary dynamics and epidemiology. PloS ONE 7, e51439 (2012).

  81. 81.

    Coppes, M. J., Liefers, G. J., Paul, P., Yeger, H. & Williams, B. R. Homozygous somatic Wt1 point mutations in sporadic unilateral Wilms tumor. Proc. Natl Acad. Sci. USA 90, 1416–1419 (1993).

  82. 82.

    Heitzer, E. et al. Tumor-associated copy number changes in the circulation of patients with prostate cancer identified through whole-genome sequencing. Genome Med. 5, 30 (2013).

Download references

Acknowledgements

We thank the Hubrecht Imaging Center for assistance with (confocal) microscopy. We thank H. Begthel and J. Korving (both Hubrecht Institute) for preparation of histological and immunohistochemical specimens. We thank T. Nguyen (UMC Utrecht, the Netherlands) and R. de Krijger (Princess Máxima Center for Pediatric Oncology, Utrecht, the Netherlands) for help with the analysis of histological specimens. We thank the Hubrecht FACS facility for help with the sort of single (EPCAM+) cells and G. Posthuma (Department of Cell Biology and Institute of Biomembranes, UMC Utrecht, the Netherlands) for excellent support with transmission electron microscopy. We thank E. Driehuis for the photography of the VP-1 staining and for help with finalizing the manuscript. We thank J. Hoenderop (Radboud UMC, the Netherlands) for kindly providing UMOD and SLC12A1 antibodies, and NC3Rs for development of the kidney-on-a-chip assays (Nephrotube, CRACK-IT challenge). This work was supported by a grant from the Dutch Kidney Foundation (grant no. DKF14OP04), and Zwaartekracht (NWO). This work was supported by the partners of Regenerative Medicine Crossing Borders (www.regmedxb.com). Powered by Health~Holland, Top Sector Life Sciences & Health.

Author information

Affiliations

  1. Hubrecht Institute—Royal Netherlands Academy of Arts and Sciences, Utrecht, the Netherlands

    • Frans Schutgens
    • , Carola Ammerlaan
    • , Fjodor Yousef Yengej
    • , Benedetta Artegiani
    •  & Hans Clevers
  2. Department of Nephrology and Hypertension, University Medical Centre Utrecht, Utrecht, the Netherlands

    • Frans Schutgens
    • , Maarten B Rookmaaker
    • , Carola Ammerlaan
    • , Fjodor Yousef Yengej
    •  & Marianne C Verhaar
  3. Princess Máxima Center for Pediatric Oncology, Utrecht, the Netherlands

    • Thanasis Margaritis
    • , Anne Rios
    • , Sepide Derakhshan
    • , Ruben van Boxtel
    • , Marry M. van den Heuvel-Eibrink
    • , Frank Holstege
    • , Jarno Drost
    •  & Hans Clevers
  4. Division of Pharmacology, Utrecht Institute for Pharmaceutical Sciences, Utrecht University, Utrecht, the Netherlands

    • Jitske Jansen
    • , Jean-Luc Murk
    •  & Rosalinde Masereeuw
  5. Mimetas, Organ-on-a-chip Company, Leiden, the Netherlands

    • Linda Gijzen
    • , Marianne Vormann
    •  & Henriette Lanz
  6. Regenerative Medicine Center Utrecht, Utrecht, the Netherlands

    • Annelotte Vonk
    •  & Jeffrey Beekman
  7. Department of Medical Microbiology, University Medical Center Utrecht, Utrecht, the Netherlands

    • Marco Viveen
    •  & Antoni P. A. Hendrickx
  8. Wilhelmina Children’s Hospital, University Medical Center Utrecht, Utrecht, the Netherlands

    • Karin M. de Winter-de Groot
  9. Center for Molecular Medicine, Cancer Genomics Netherlands, Department of Genetics, University Medical Center Utrecht, Utrecht, the Netherlands

    • Edwin Cuppen
  10. Institute of Human Genetics, Medical University of Graz, Graz, Austria

    • Ellen Heitzer
  11. Laboratory of Medical Microbiology and Immunology, St. Elisabeth TweeSteden Ziekenhuis, Tilburg, the Netherlands

    • Jean-Luc Murk

Authors

  1. Search for Frans Schutgens in:

  2. Search for Maarten B Rookmaaker in:

  3. Search for Thanasis Margaritis in:

  4. Search for Anne Rios in:

  5. Search for Carola Ammerlaan in:

  6. Search for Jitske Jansen in:

  7. Search for Linda Gijzen in:

  8. Search for Marianne Vormann in:

  9. Search for Annelotte Vonk in:

  10. Search for Marco Viveen in:

  11. Search for Fjodor Yousef Yengej in:

  12. Search for Sepide Derakhshan in:

  13. Search for Karin M. de Winter-de Groot in:

  14. Search for Benedetta Artegiani in:

  15. Search for Ruben van Boxtel in:

  16. Search for Edwin Cuppen in:

  17. Search for Antoni P. A. Hendrickx in:

  18. Search for Marry M. van den Heuvel-Eibrink in:

  19. Search for Ellen Heitzer in:

  20. Search for Henriette Lanz in:

  21. Search for Jeffrey Beekman in:

  22. Search for Jean-Luc Murk in:

  23. Search for Rosalinde Masereeuw in:

  24. Search for Frank Holstege in:

  25. Search for Jarno Drost in:

  26. Search for Marianne C Verhaar in:

  27. Search for Hans Clevers in:

Contributions

F.S., M.B.R., M.C.V. and H.C. designed, performed, analyzed experiments and wrote the manuscript. C.A. established and maintained tubuloid cultures and performed and analyzed karyotyping experiments. A.R. performed, imaged and analyzed immunofluorescent stainings. T.M. and F.H. analyzed the single-cell sequencing data. F.S., J.J. and R.M. designed, analyzed and performed the P-gp transporter assays in tubuloids. A.P.A.H. provided support with scanning electron microscopy images. F.S., M.Vi., M.B.R. and J.M. designed, performed and analyzed the BK virus experiments. R.v.B analyzed and interpreted the WGS analysis and E.C. gave input on the interpretation. B.A. helped with the analysis of the the bulk RNA-seq. M.M.H.E. established the logistics of obtaining clinical samples of nephroblastoma tissue. F.S. and J.D. established nephroblastoma cultures and designed experiments. E.H. performed and analyzed the nephroblastoma CNV analysis. S.D. established the clonal tubuloid line and F.Y.Y. analyzed the clonal tubuloid line. K.M.W.G. obtained CF urine samples. F.S., A.V. and J.B. designed FSK-induced swelling experiments and analyzed data. F.S., L.G., M.Vo. and H.L. designed, performed and analyzed the ‘tubuloid-on-a-chip’ experiments. All authors commented on the manuscript.

Competing interests

H.C. is a holder of several patents related to organoid technology. L.G., M.Vo. and H.L. are employees of MIMETAS BV, the Netherlands, that is marketing the OrganoPlate. OrganoPlate is a trademark of MIMETAS.

Corresponding author

Correspondence to Hans Clevers.

Integrated supplementary information

  1. Supplementary Figure 1 Quantification of human metaphase spreads.

    > 40 spreads were quantified, from tubuloids in P11, P14 and P18 in 3 independent experiments, from 3 independent tubuloid cultures.

  2. Supplementary Figure 2 Set-up and mutation list of the whole-genome sequencing (WGS) analysis of short-term and long-term tubuloid culture.

    A tubuloid line was established from cortical kidney tissue after nephrectomy and tubuloids in passage 2 (P2) and passage 8 (P8) were, along with the matched tissue sample, harvested for WGS. A pie chart with types of mutations and a list of the observed missense mutations in coding regions are included. None of the reported mutations is reported in the COSMIC database or associated with kidney (dys)function. Green boxes indicate the part of the tubuloids that are used for establishing the next passage. Red boxes indicate the samples that were harvested for WGS.

  3. Supplementary Figure 3 Adult mouse kidney tubuloid culture.

    Scheme of the experimental protocol (a). The development of mouse kidney epithelial cells into folded / branching structures after seeding; image of passage 6 (representative image of at least n = 3 lines) (b). H&E stain at passage 7 (representative image of an H&E stain that was performed for at least n = 3 lines) (c). An example of a typical metaphase spread, from a tubuloid culture of passage number > 11 (d). Quantification of > 40 spreads in 3 independent experiments, from tubuloids at P11, P15 and P16 (e). Scale bars 100 µm (b, c) and 10 µm (d).

  4. Supplementary Figure 4 Kidney tubuloids are more proliferative than primary kidney epithelial cells.

    EPCAM+ kidney epithelial cells and tubuloid cells were analysed. 67% of the primary kidney epithelial cells were in GO/G1, compared to 40% of the tubuloid cells (see Methods).

  5. Supplementary Figure 5 Human kidney tubuloids are derived from kidney epithelium.

    Normalized log2 transcript counts show expression of the pan-kidney epithelium marker PAX8 across different clusters (192 kidney and 192 tubuloid cells were sequenced in one run and after quality checks (see Methods), 51 kidney cells and 149 tubuloid cells were used for clustering analysis. Cluster 1: n = 8; Cluster 2: n = 45; Cluster 3: n = 33; Cluster 4: n = 19; Cluster 5: n = 39; Cluster 6: n = 28; Cluster 7: n = 28. Tukey box plots are used to visualize the distributions. All points of the populations are shown as jittered dots.) (a). The expression of PAX8 is confirmed by staining (n = 4 tubuloid lines) (b). Scale bar: 100 µm.

  6. Supplementary Figure 6 Expression of marker genes in specific single-cell RNA-seq clusters.

    Cluster 3–7 contains only tubuloid-derived cells; cluster 2 contains only primary kidney epithelial cells and cluster 1 is a mix between tubuloid and primary kidney epithelial cells. Normalized log2 transcript counts show expression of (kidney) epithelium markers PAX8, EPCAM, KRT18 and KRT19 across different clusters, showing that all cells are epithelial in nature (a). Normalized log2 transcript counts of typical intercalated genes SLC26A7, SLC4A1, ATP6V1B1, that are expressed in cluster 1, a combination of primary kidney epithelial cells and tubuloid-derived cells (b). Normalized log2 transcript counts show expression of collecting duct principal cell genes SLC14A1 and CLDN8 in cluster 4 (c). Normalized log2 transcript counts show expression of glucose handling genes SLC2A1, ALDOC as well as VEGFA, suggestive of proximal tubule cells in cluster 5 (d). Cluster 6 does not express many segment-specific genes, except SLC22A5, which may suggest a (proximal tubule) progenitor cell phenotype (e) Normalized log2 transcript counts how higher expression of COL4A3 and COL4A4 and lower expression of KRT18 and KRT19 in cluster 7, suggesting a pro-fibrotic phenotype (f). In a-f, 192 kidney and 192 tubuloid cells were sequenced in one run and after quality checks (see methods), 51 kidney cells and 149 tubuloid cells were used for clustering analysis. Cluster 1: n = 8; Cluster 2: n = 45; Cluster 3: n = 33; Cluster 4: n = 19; Cluster 5: n = 39; Cluster 6: n = 28; Cluster 7: n = 28. Tukey box plots are used to visualize the distributions. All points of the populations are shown as jittered dots.

  7. Supplementary Figure 7 Expression of distal tubule marker CALB1 and loop of Henle marker UMOD can be induced by growth factor withdrawal.

    CALB1 expression is absent in kidney tubuloids on the protein level (a). By withdrawal of growth factors from the culture medium, CALB1 can be induced, as visualized with immunofluorescence (b). UMOD expression is absent in kidney tubuloids on the protein level (c). By withdrawal of growth factors from the culture medium, UMOD can be induced, as visualized with immunofluorescence (d). Representative images of n = 2 independent experiments. Scale bars: 75 µm.

  8. Supplementary Figure 8 Schematic representation of the proximal tubule functional assay.

    When tubuloids are exposed to calcein-AM, a substrate of P-gp (the xenobiotics efflux pump, located at the apical membrane) that diffuses freely into cells and that becomes fluorescent inside cells after cleaving the acetomethoxy group resulting in calcein, in the presence of an inhibitor (PSC-833) of P-gp, calcein accumulates (a). In absence of the inhibitor, P-gp pumps calcein from the cells, thereby preventing accumulation of fluorescent signal (b).

  9. Supplementary Figure 9 P-gp is functional in human kidney tubuloids.

    In the presence of specific P-gp-inhibitor PSC-833, calcein accumulates in tubuloids, as measured by fluorescent plate reader. Normalized quantification of n = 3 independent experiments, with each experiment n ≥ 5 plate reader measurements. Lines indicate means per experiment. * P = 0.0016 with an unpaired two-tailed t test, after pooling the individual measurements from n = 3 experiments.

  10. Supplementary Figure 10 Expression analysis of a clonal tubuloid line indicates multi-lineage potential of tubuloid cells.

    Gene expression of marker genes of the proximal tubule (ANPEP, ABCC4, SLC4A4), Loop of Henle (SLC12A1, CLDN10), distal tubule (SLC12A3, SLC41A3, PCBD1) and collecting duct (AQP3, NR3C2) was determined in the clonal tubuloid line and compared with the bulk tubuloid line that was used for establishing the clonal line. The clonal line preserves expression of markers of multiple segments: the proximal tubule genes ABCC4 and SLC4A4 are similar to the bulk tubuloid line, whereas expression of the typical Loop of Henle gene SLC12A1 is increased and the typical distal tubule marker SLC12A3 is decreased. These data indicate multi-lineage potential of a single tubuloid cell (proximal tubule and Loop of Henle). Expression levels were normalized to RPLP0, and expressed as fold change to expression levels of a colon organoid line.

  11. Supplementary Figure 11 WT1 targeted sequencing analysis for patient 1.

    Sanger sequencing shows a heterozygous 8 base pair deletion in the healthy kidney tissue (H tissue); tubuloids derived from the healthy tissue (H tubuloid); tumor tissue (T tissue) and the tubuloids derived from the tumor tissue (T tumoroid) (a). Sanger sequencing of WT1 shows a frameshift mutation in exon 10: it is a heterozygous 4 base pair insertion in the tumor tissue (T tissue) and the tumoroids derived from the tumor tissue (T tumoroid). This insertion is absent in the healthy kidney tissue (H tissue) and tubuloids derived from the healthy tissue (H tubuloid) (b). In both a and b: representative sequences of at least n = 3 independent experiments.

  12. Supplementary Figure 12 Urine-derived tubuloids from a subject with CF are kidney tubuloids.

    Assessed with a PAX8 staining (PAX8 staining performed once on this tubuloid line and on n = 3 other lines). Scale bar 100 µm.

  13. Supplementary Figure 13 The effect of VX-770 on FSK-induced swelling in intestinal organoids and urine-derived tubuloids.

    In urine-derived tubuloids and intestinal organoids from the same patient, VX-770 increased FSK-induced swelling. Average is plotted of n = 3 independent experiments that were performed in duplicate, error bars represent standard deviation.

  14. Supplementary Figure 14 Overview of the trans-epithelial transporter assay.

    Rhodamine 123 is transported into cells on the basal side by OCT-transporters and removed on the apical side by efflux pump P-gp (a). In the presence of P-gp inhibitor PSC-833, rhodamine 123 fluorescence in the lumen on the apical side, is expected to be reduced (b).

  15. Supplementary Figure 15 Filtering out severely stressed cells from the single-cell sequencing analysis.

    Dissociation-induced stress was quantified using a scoring strategy (see methods) based on the expression of heat shock genes. A score over 0.4 was used to filter out cells that were severely stressed. 192 kidney and 192 tubuloid cells were sequenced in one run and after quality checks (see methods), cells were analyzed for dissociation-induced stress (Kidney 1: n = 41; Kidney 2: n = 43; Tubloid 1: n = 76; Tubloid 2: n = 73). Tukey box plots are used to visualize the distributions. All points of the populations are shown as jittered dots.

Supplementary information

  1. Supplementary Text, Figures and Tables

    Supplementary Figures 1–15 and Supplementary Tables 1 and 2.

  2. Reporting Summary

  3. Supplementary Video 1

    Tubuloids are epithelial in nature and cells are polarized.

  4. Supplementary Video 2

    Wilms tumor–derived tumoroids (from patient 1) have a different morphology than matched normal tubuloids. This 6-day time-lapse video shows the matched normal tubuloids.

  5. Supplementary Video 3

    Wilms tumor–derived tumoroids (from patient 1) have a different morphology than matched normal tubuloids. This 6-day time-lapse video shows the tumoroids.

  6. Supplementary Video 4

    Wilms tumor–derived tumoroids (from patient 1) have a different morphology than matched normal tubuloids. This 6-day time-lapse video shows a detail of the tumoroids.

  7. Supplementary Video 5

    Forskolin-induced swelling assay of CF urine-derived tubuloids without VX-770.

  8. Supplementary Video 6

    Forskolin-induced swelling assay of CF urine-derived tubuloids with VX-770.

  9. Supplementary Video 7

    Forskolin-induced swelling assay of CF intestinal organoids without VX-770.

  10. Supplementary Video 8

    Forskolin-induced swelling assay of CF intestinal organoids with VX-770.

  11. Supplementary Dataset 1

    Bulk RNA sequencing data. CSV file of n = 3 (1, 2, 3) tissue samples (FS1) and matched tubuloid lines (FS2).

  12. Supplementary Dataset 2

    Single-cell sequencing data.

  13. Supplementary Dataset 3

    DNA sequencing data at day 1 of infection and day 30 of infection.

About this article

Publication history

Received

Accepted

Published

Issue Date

DOI

https://doi.org/10.1038/s41587-019-0048-8