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Transcriptional networks controlling stromal cell differentiation

Nature Reviews Molecular Cell Biology volume 22, pages 465–482 (2021)Cite this article

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Abstract

Stromal progenitors are found in many different tissues, where they play an important role in the maintenance of tissue homeostasis owing to their ability to differentiate into parenchymal cells. These progenitor cells are differentially pre-programmed by their tissue microenvironment but, when cultured and stimulated in vitro, these cells — commonly referred to as mesenchymal stromal cells (MSCs) — exhibit a marked plasticity to differentiate into many different cell lineages. Loss-of-function studies in vitro and in vivo have uncovered the involvement of specific signalling pathways and key transcriptional regulators that work in a sequential and coordinated fashion to activate lineage-selective gene programmes. Recent advances in omics and single-cell technologies have made it possible to obtain system-wide insights into the gene regulatory networks that drive lineage determination and cell differentiation. These insights have important implications for the understanding of cell differentiation, the contribution of stromal cells to human disease and for the development of cell-based therapeutic applications.

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Fig. 1: Culture conditions and plasticity of stromal progenitor cells in vitro.
Fig. 2: Adipocyte lineage determination and differentiation.
Fig. 3: The osteochondral lineage.
Fig. 4: The muscle cell lineage.
Fig. 5: The ability of master regulators to drive lineage determination.
Fig. 6: Chromatin-looping, lineage-selective and stem cell transcription factors in the control of lineage fate.

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References

  1. Owen, M. & Friedenstein, A. J. Stromal stem cells: marrow-derived osteogenic precursors. Ciba Found. Symp. 136, 42–60 (1988).

    CAS  PubMed  Google Scholar 

  2. Tavassoli, M. & Crosby, W. H. Transplantation of marrow to extramedullary sites. Science 161, 54–56 (1968).

    Article  CAS  PubMed  Google Scholar 

  3. Friedenstein, A. J., Chailakhjan, R. K. & Lalykina, K. S. The development of fibroblast colonies in monolayer cultures of guinea-pig bone marrow and spleen cells. Cell Tissue Kinet. 3, 393–403 (1970).

    CAS  PubMed  Google Scholar 

  4. Dominici, M. et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8, 315–317 (2006).

    Article  CAS  PubMed  Google Scholar 

  5. da Silva Meirelles, L., Chagastelles, P. C. & Nardi, N. B. Mesenchymal stem cells reside in virtually all post-natal organs and tissues. J. Cell Sci. 119, 2204–2213 (2006).

    Article  PubMed  Google Scholar 

  6. Sacchetti, B. et al. No identical “Mesenchymal Stem Cells” at different times and sites: human committed progenitors of distinct origin and differentiation potential are incorporated as adventitial cells in microvessels. Stem Cell Rep. 6, 897–913 (2016).

    Article  CAS  Google Scholar 

  7. Feng, G. et al. Imaging neuronal subsets in transgenic mice expressing multiple spectral variants of GFP. Neuron 28, 41–51 (2000).

    Article  CAS  PubMed  Google Scholar 

  8. Alemany, A., Florescu, M., Baron, C. S., Peterson-Maduro, J. & van Oudenaarden, A. Whole-organism clone tracing using single-cell sequencing. Nature 556, 108–112 (2018).

    Article  CAS  PubMed  Google Scholar 

  9. Stergachis, A. B. et al. Developmental fate and cellular maturity encoded in human regulatory DNA landscapes. Cell 154, 888–903 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  10. Thurman, R. E. et al. The accessible chromatin landscape of the human genome. Nature 489, 75–82 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  11. Neph, S. et al. An expansive human regulatory lexicon encoded in transcription factor footprints. Nature 489, 83–90 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572 e24 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Zhang, Y. et al. Chromatin connectivity maps reveal dynamic promoter-enhancer long-range associations. Nature 504, 306–310 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Dixon, J. R. et al. Chromatin architecture reorganization during stem cell differentiation. Nature 518, 331–336 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Galipeau, J. & Sensebe, L. Mesenchymal stromal cells: clinical challenges and therapeutic opportunities. Cell Stem Cell 22, 824–833 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Levy, O. et al. Shattering barriers toward clinically meaningful MSC therapies. Sci. Adv. 6, eaba6884 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. Zwolanek, D. et al. Tracking mesenchymal stem cell contributions to regeneration in an immunocompetent cartilage regeneration model. JCI Insight 2, e87322 (2017).

    Article  PubMed Central  Google Scholar 

  18. von Bahr, L. et al. Analysis of tissues following mesenchymal stromal cell therapy in humans indicates limited long-term engraftment and no ectopic tissue formation. Stem Cell 30, 1575–1578 (2012).

    Article  Google Scholar 

  19. Scheja, L. & Heeren, J. The endocrine function of adipose tissues in health and cardiometabolic disease. Nat. Rev. Endocrinol. 15, 507–524 (2019).

    Article  CAS  PubMed  Google Scholar 

  20. White, U. A. & Tchoukalova, Y. D. Sex dimorphism and depot differences in adipose tissue function. Biochim. Biophys. Acta 1842, 377–392 (2014).

    Article  CAS  PubMed  Google Scholar 

  21. Naveiras, O. et al. Bone-marrow adipocytes as negative regulators of the haematopoietic microenvironment. Nature 460, 259–263 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. Wilson, A. et al. Lack of adipocytes alters hematopoiesis in lipodystrophic mice. Front. Immunol. 9, 2573 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  23. Zhang, Z. et al. Dermal adipose tissue has high plasticity and undergoes reversible dedifferentiation in mice. J. Clin. Invest. 129, 5327–5342 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Billon, N. et al. The generation of adipocytes by the neural crest. Development 134, 2283–2292 (2007).

    Article  CAS  PubMed  Google Scholar 

  25. Sebo, Z. L. & Rodeheffer, M. S. Assembling the adipose organ: adipocyte lineage segregation and adipogenesis in vivo. Development 146, dev172098 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Sebo, Z. L., Jeffery, E., Holtrup, B. & Rodeheffer, M. S. A mesodermal fate map for adipose tissue. Development 145, dev166801 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  27. Jeffery, E. et al. The adipose tissue microenvironment regulates depot-specific adipogenesis in obesity. Cell Metab. 24, 142–150 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. Lee, Y. H., Petkova, A. P., Mottillo, E. P. & Granneman, J. G. In vivo identification of bipotential adipocyte progenitors recruited by beta3-adrenoceptor activation and high-fat feeding. Cell Metab. 15, 480–491 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Vishvanath, L. et al. Pdgfrbeta+ mural preadipocytes contribute to adipocyte hyperplasia induced by high-fat-diet feeding and prolonged cold exposure in adult mice. Cell Metab. 23, 350–359 (2016).

    Article  CAS  PubMed  Google Scholar 

  30. Hudak, C. S. et al. Pref-1 marks very early mesenchymal precursors required for adipose tissue development and expansion. Cell Rep. 8, 678–687 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Gupta, R. K. et al. Zfp423 expression identifies committed preadipocytes and localizes to adipose endothelial and perivascular cells. Cell Metab. 15, 230–239 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Vanlandewijck, M. et al. A molecular atlas of cell types and zonation in the brain vasculature. Nature 554, 475–480 (2018).

    Article  CAS  PubMed  Google Scholar 

  33. Crisan, M. et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell 3, 301–313 (2008).

    Article  CAS  PubMed  Google Scholar 

  34. Jiang, Y., Berry, D. C., Tang, W. & Graff, J. M. Independent stem cell lineages regulate adipose organogenesis and adipose homeostasis. Cell Rep. 9, 1007–1022 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. Tang, W. et al. White fat progenitor cells reside in the adipose vasculature. Science 322, 583–586 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  36. Guimaraes-Camboa, N. et al. Pericytes of multiple organs do not behave as mesenchymal stem cells in vivo. Cell Stem Cell 20, 345–359.e5 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Cano, E., Gebala, V. & Gerhardt, H. Pericytes or mesenchymal stem cells: is that the question? Cell Stem Cell 20, 296–297 (2017).

    Article  CAS  PubMed  Google Scholar 

  38. Flores, K. G., Li, J., Sempowski, G. D., Haynes, B. F. & Hale, L. P. Analysis of the human thymic perivascular space during aging. J. Clin. Invest. 104, 1031–1039 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Saisho, Y. et al. Pancreas volumes in humans from birth to age one hundred taking into account sex, obesity, and presence of type-2 diabetes. Clin. Anat. 20, 933–942 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. Burl, R. B. et al. Deconstructing adipogenesis induced by beta3-adrenergic receptor activation with single-cell expression profiling. Cell Metab. 28, 300–309.e4 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Schwalie, P. C. et al. A stromal cell population that inhibits adipogenesis in mammalian fat depots. Nature 559, 103–108 (2018).

    Article  CAS  PubMed  Google Scholar 

  42. Hepler, C. et al. Identification of functionally distinct fibro-inflammatory and adipogenic stromal subpopulations in visceral adipose tissue of adult mice. eLife 7, e39636 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  43. Merrick, D. et al. Identification of a mesenchymal progenitor cell hierarchy in adipose tissue. Science 364, eaav2501 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Benias, P. C. et al. Structure and distribution of an unrecognized interstitium in human tissues. Sci. Rep. 8, 4947 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  45. Sarvari, A. K. et al. Plasticity of epididymal adipose tissue in response to diet-induced obesity at single-nucleus resolution. Cell Metab. 33, 437–453.e5 (2021). Savari et al. used single nucleus RNA-seq to infer the in vivo developmental trajectory and the associated transcriptional changes from stromal progenitors to mature adipocytes.

    Article  CAS  PubMed  Google Scholar 

  46. Siersbaek, R. & Mandrup, S. Transcriptional networks controlling adipocyte differentiation. Cold Spring Harb. Symp. Quant. Biol. 76, 247–255 (2011).

    Article  CAS  PubMed  Google Scholar 

  47. Green, H. & Meuth, M. An established pre-adipose cell line and its differentiation in culture. Cell 3, 127–133 (1974).

    Article  CAS  PubMed  Google Scholar 

  48. Hiragun, A., Sato, M. & Mitsui, H. Preadipocyte differentiation in vitro: identification of a highly active adipogenic agent. J. Cell Physiol. 134, 124–130 (1988).

    Article  CAS  PubMed  Google Scholar 

  49. Siersbaek, R. et al. Transcription factor cooperativity in early adipogenic hotspots and super-enhancers. Cell Rep. 7, 1443–1455 (2014).

    Article  CAS  PubMed  Google Scholar 

  50. Mota de Sa, P., Richard, A. J., Hang, H. & Stephens, J. M. Transcriptional regulation of adipogenesis. Compr. Physiol. 7, 635–674 (2017).

    Article  PubMed  Google Scholar 

  51. Lefterova, M. I. et al. PPARγ and C/EBP factors orchestrate adipocyte biology via adjacent binding on a genome-wide scale. Genes Dev. 22, 2941–2952 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  52. Nielsen, R. et al. Genome-wide profiling of PPARγ:RXR and RNA polymerase II occupancy reveals temporal activation of distinct metabolic pathways and changes in RXR dimer composition during adipogenesis. Genes Dev. 22, 2953–2967 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  53. Madsen, M. S., Siersbaek, R., Boergesen, M., Nielsen, R. & Mandrup, S. Peroxisome proliferator-activated receptor γ and C/EBPα synergistically activate key metabolic adipocyte genes by assisted loading. Mol. Cell Biol. 34, 939–954 (2014).

    Article  PubMed  PubMed Central  Google Scholar 

  54. Soccio, R. E. et al. Genetic variation determines PPARγ function and anti-diabetic drug response in vivo. Cell 162, 33–44 (2015). The study by Soccio et al. highlights the impact of genetic variations in PPAR and C/EBP motifs on PPARγ binding that accounts for changes in gene expression in response to PPARγ agonists and individual disease risk.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  55. Siersbaek, M. S. et al. Genome-wide profiling of peroxisome proliferator-activated receptor gamma in primary epididymal, inguinal, and brown adipocytes reveals depot-selective binding correlated with gene expression. Mol. Cell Biol. 32, 3452–3463 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  56. Rajakumari, S. et al. EBF2 determines and maintains brown adipocyte identity. Cell Metab. 17, 562–574 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  57. Loft, A. et al. Browning of human adipocytes requires KLF11 and reprogramming of PPARγ superenhancers. Genes Dev. 29, 7–22 (2015).

    Article  PubMed  PubMed Central  Google Scholar 

  58. Hu, E., Tontonoz, P. & Spiegelman, B. M. Transdifferentiation of myoblasts by the adipogenic transcription factors PPAR gamma and C/EBP alpha. Proc. Natl Acad. Sci. USA 92, 9856–9860 (1995).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. Tontonoz, P., Hu, E. & Spiegelman, B. M. Stimulation of adipogenesis in fibroblasts by PPARγ2, a lipid-activated transcription factor. Cell 79, 1147–1156 (1994).

    Article  CAS  PubMed  Google Scholar 

  60. Linhart, H. G. et al. C/EBPα is required for differentiation of white, but not brown, adipose tissue. Proc. Natl Acad. Sci. USA 98, 12532–12537 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  61. Wang, F., Mullican, S. E., DiSpirito, J. R., Peed, L. C. & Lazar, M. A. Lipoatrophy and severe metabolic disturbance in mice with fat-specific deletion of PPARγ. Proc. Natl Acad. Sci. USA 110, 18656–18661 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  62. Wang, Q. A. et al. Peroxisome proliferator-activated receptor gamma and its role in adipocyte homeostasis and thiazolidinedione-mediated insulin sensitization. Mol. Cell. Biol. 38, e00677–17 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  63. Wang, Q. A. et al. Distinct regulatory mechanisms governing embryonic versus adult adipocyte maturation. Nat. Cell Biol. 17, 1099–1111 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. Imai, T. et al. Peroxisome proliferator-activated receptor γ is required in mature white and brown adipocytes for their survival in the mouse. Proc. Natl Acad. Sci. USA 101, 4543–4547 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  65. Mikkelsen, T. S. et al. Comparative epigenomic analysis of murine and human adipogenesis. Cell 143, 156–169 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  66. Rauch, A. et al. Osteogenesis depends on commissioning of a network of stem cell transcription factors that act as repressors of adipogenesis. Nat. Genet. 51, 716–727 (2019). Rauch et al. identified a large group of transcription factors with pro-osteogenic and anti-adipogenic activity and report that variations in their binding sites or in factor abundancy are associated with human diseases.

    Article  CAS  PubMed  Google Scholar 

  67. Siersbaek, R. et al. Extensive chromatin remodelling and establishment of transcription factor ‘hotspots’ during early adipogenesis. EMBO J. 30, 1459–1472 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Siersbaek, R. et al. Dynamic rewiring of promoter-anchored chromatin loops during adipocyte differentiation. Mol. Cell 66, 420–435.e5 (2017).

    Article  CAS  PubMed  Google Scholar 

  69. Madsen, J. G. S. et al. Integrated analysis of motif activity and gene expression changes of transcription factors. Genome Res. 28, 243–255 (2017).

    Article  PubMed  Google Scholar 

  70. Dirckx, N., Moorer, M. C., Clemens, T. L. & Riddle, R. C. The role of osteoblasts in energy homeostasis. Nat. Rev. Endocrinol. 15, 651–665 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  71. Marin-Llera, J. C., Garciadiego-Cazares, D. & Chimal-Monroy, J. Understanding the cellular and molecular mechanisms that control early cell fate decisions during appendicular skeletogenesis. Front. Genet. 10, 977 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  72. Yang, L., Tsang, K. Y., Tang, H. C., Chan, D. & Cheah, K. S. Hypertrophic chondrocytes can become osteoblasts and osteocytes in endochondral bone formation. Proc. Natl Acad. Sci. USA 111, 12097–12102 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. Ono, N., Ono, W., Nagasawa, T. & Kronenberg, H. M. A subset of chondrogenic cells provides early mesenchymal progenitors in growing bones. Nat. Cell Biol. 16, 1157–1167 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  74. Roach, H. I. Trans-differentiation of hypertrophic chondrocytes into cells capable of producing a mineralized bone matrix. Bone Miner. 19, 1–20 (1992).

    Article  CAS  PubMed  Google Scholar 

  75. Decker, R. S. et al. Cell origin, volume and arrangement are drivers of articular cartilage formation, morphogenesis and response to injury in mouse limbs. Dev. Biol. 426, 56–68 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  76. Koyama, E. et al. A distinct cohort of progenitor cells participates in synovial joint and articular cartilage formation during mouse limb skeletogenesis. Dev. Biol. 316, 62–73 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  77. Kozhemyakina, E. et al. Identification of a Prg4-expressing articular cartilage progenitor cell population in mice. Arthritis Rheumatol. 67, 1261–1273 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  78. Mizuhashi, K. et al. Resting zone of the growth plate houses a unique class of skeletal stem cells. Nature 563, 254–258 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Newton, P. T. et al. A radical switch in clonality reveals a stem cell niche in the epiphyseal growth plate. Nature 567, 234–238 (2019).

    Article  CAS  PubMed  Google Scholar 

  80. Debnath, S. et al. Discovery of a periosteal stem cell mediating intramembranous bone formation. Nature 562, 133–139 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  81. Zhou, B. O., Yue, R., Murphy, M. M., Peyer, J. G. & Morrison, S. J. Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell 15, 154–168 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Zhou, X. et al. Chondrocytes transdifferentiate into osteoblasts in endochondral bone during development, postnatal growth and fracture healing in mice. PLoS Genet. 10, e1004820 (2014). Zhou et al. demonstrate hypertrophic chondrocytes in the fracture callus and the growth plate have the ability to differentiate into the osteogenic lineage.

    Article  PubMed  PubMed Central  Google Scholar 

  83. Matsushita, Y. et al. A Wnt-mediated transformation of the bone marrow stromal cell identity orchestrates skeletal regeneration. Nat. Commun. 11, 332 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  84. Ambrosi, T. H., Longaker, M. T. & Chan, C. K. F. A revised perspective of skeletal stem cell biology. Front. Cell Dev. Biol. 7, 189 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  85. Wolock, S. L. et al. Mapping distinct bone marrow niche populations and their differentiation paths. Cell Rep. 28, 302–311.e5 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  86. Baryawno, N. et al. A cellular taxonomy of the bone marrow stroma in homeostasis and leukemia. Cell 177, 1915–1932.e16 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  87. Baccin, C. et al. Combined single-cell and spatial transcriptomics reveal the molecular, cellular and spatial bone marrow niche organization. Nat. Cell Biol. 22, 38–48 (2020).

    Article  CAS  PubMed  Google Scholar 

  88. Johnstone, B., Hering, T. M., Caplan, A. I., Goldberg, V. M. & Yoo, J. U. In vitro chondrogenesis of bone marrow-derived mesenchymal progenitor cells. Exp. Cell Res. 238, 265–272 (1998).

    Article  CAS  PubMed  Google Scholar 

  89. Sekiya, I., Larson, B. L., Vuoristo, J. T., Reger, R. L. & Prockop, D. J. Comparison of effect of BMP-2, -4, and -6 on in vitro cartilage formation of human adult stem cells from bone marrow stroma. Cell Tissue Res. 320, 269–276 (2005).

    Article  CAS  PubMed  Google Scholar 

  90. Estes, B. T., Wu, A. W. & Guilak, F. Potent induction of chondrocytic differentiation of human adipose-derived adult stem cells by bone morphogenetic protein 6. Arthritis Rheum. 54, 1222–1232 (2006).

    Article  CAS  PubMed  Google Scholar 

  91. Mackay, A. M. et al. Chondrogenic differentiation of cultured human mesenchymal stem cells from marrow. Tissue Eng. 4, 415–428 (1998).

    Article  CAS  PubMed  Google Scholar 

  92. Sart, S., Tsai, A. C., Li, Y. & Ma, T. Three-dimensional aggregates of mesenchymal stem cells: cellular mechanisms, biological properties, and applications. Tissue Eng. Part. B Rev. 20, 365–380 (2014).

    Article  PubMed  Google Scholar 

  93. Watts, A. E., Ackerman-Yost, J. C. & Nixon, A. J. A comparison of three-dimensional culture systems to evaluate in vitro chondrogenesis of equine bone marrow-derived mesenchymal stem cells. Tissue Eng. Part A 19, 2275–2283 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Ng, L. J. et al. SOX9 binds DNA, activates transcription, and coexpresses with type II collagen during chondrogenesis in the mouse. Dev. Biol. 183, 108–121 (1997).

    Article  CAS  PubMed  Google Scholar 

  95. Zhao, Q., Eberspaecher, H., Lefebvre, V. & de Crombrugghe, B. Parallel expression of Sox9 and Col2a1 in cells undergoing chondrogenesis. Dev. Dyn. 209, 377–386 (1997).

    Article  CAS  PubMed  Google Scholar 

  96. Bi, W., Deng, J. M., Zhang, Z., Behringer, R. R. & de Crombrugghe, B. Sox9 is required for cartilage formation. Nat. Genet. 22, 85–89 (1999).

    Article  CAS  PubMed  Google Scholar 

  97. Akiyama, H., Chaboissier, M. C., Martin, J. F., Schedl, A. & de Crombrugghe, B. The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev. 16, 2813–2828 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  98. Dy, P. et al. Sox9 directs hypertrophic maturation and blocks osteoblast differentiation of growth plate chondrocytes. Dev. Cell 22, 597–609 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  99. Foster, J. W. et al. Campomelic dysplasia and autosomal sex reversal caused by mutations in an SRY-related gene. Nature 372, 525–530 (1994).

    Article  CAS  PubMed  Google Scholar 

  100. Wagner, T. et al. Autosomal sex reversal and campomelic dysplasia are caused by mutations in and around the SRY-related gene SOX9. Cell 79, 1111–1120 (1994).

    Article  CAS  PubMed  Google Scholar 

  101. Smits, P. et al. The transcription factors L-Sox5 and Sox6 are essential for cartilage formation. Dev. Cell 1, 277–290 (2001).

    Article  CAS  PubMed  Google Scholar 

  102. Lefebvre, V., Li, P. & de Crombrugghe, B. A new long form of Sox5 (L-Sox5), Sox6 and Sox9 are coexpressed in chondrogenesis and cooperatively activate the type II collagen gene. EMBO J. 17, 5718–5733 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  103. Bell, D. M. et al. SOX9 directly regulates the type-II collagen gene. Nat. Genet. 16, 174–178 (1997).

    Article  CAS  PubMed  Google Scholar 

  104. Liu, C. F. & Lefebvre, V. The transcription factors SOX9 and SOX5/SOX6 cooperate genome-wide through super-enhancers to drive chondrogenesis. Nucleic Acids Res. 43, 8183–8203 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  105. Ohba, S., He, X., Hojo, H. & McMahon, A. P. Distinct transcriptional programs underlie Sox9 regulation of the mammalian chondrocyte. Cell Rep. 12, 229–243 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  106. Liu, C. F., Angelozzi, M., Haseeb, A. & Lefebvre, V. SOX9 is dispensable for the initiation of epigenetic remodeling and the activation of marker genes at the onset of chondrogenesis. Development 145, dev164459 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  107. Herlofsen, S. R. et al. Genome-wide map of quantified epigenetic changes during in vitro chondrogenic differentiation of primary human mesenchymal stem cells. BMC Genomics 14, 105 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  108. Ikeda, T. et al. The combination of SOX5, SOX6, and SOX9 (the SOX trio) provides signals sufficient for induction of permanent cartilage. Arthritis Rheum. 50, 3561–3573 (2004).

    Article  CAS  PubMed  Google Scholar 

  109. Inada, M. et al. Maturational disturbance of chondrocytes in Cbfa1-deficient mice. Dev. Dyn. 214, 279–290 (1999).

    Article  CAS  PubMed  Google Scholar 

  110. Yoshida, C. A. et al. Runx2 and Runx3 are essential for chondrocyte maturation, and Runx2 regulates limb growth through induction of Indian hedgehog. Genes Dev. 18, 952–963 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  111. Arnold, M. A. et al. MEF2C transcription factor controls chondrocyte hypertrophy and bone development. Dev. Cell 12, 377–389 (2007).

    Article  CAS  PubMed  Google Scholar 

  112. Ionescu, A. et al. FoxA family members are crucial regulators of the hypertrophic chondrocyte differentiation program. Dev. Cell 22, 927–939 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  113. Tan, Z. et al. Synergistic co-regulation and competition by a SOX9-GLI-FOXA phasic transcriptional network coordinate chondrocyte differentiation transitions. PLoS Genet. 14, e1007346 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  114. Li, J. et al. Systematic reconstruction of molecular cascades regulating GP development using single-cell RNA-Seq. Cell Rep. 15, 1467–1480 (2016). Li et al. used scRNA-seq to reveal the in vivo trajectory of chondrocytes through the growth plate and the transcriptional reprograming that occurs as cells transition through the proliferative, pre-hypertrophic and hypertrophic zones.

    Article  CAS  PubMed  Google Scholar 

  115. Maehata, Y. et al. Both direct and collagen-mediated signals are required for active vitamin D3-elicited differentiation of human osteoblastic cells: roles of osterix, an osteoblast-related transcription factor. Matrix Biol. 25, 47–58 (2006).

    Article  CAS  PubMed  Google Scholar 

  116. Jorgensen, N. R., Henriksen, Z., Sorensen, O. H. & Civitelli, R. Dexamethasone, BMP-2, and 1,25-dihydroxyvitamin D enhance a more differentiated osteoblast phenotype: validation of an in vitro model for human bone marrow-derived primary osteoblasts. Steroids 69, 219–226 (2004).

    Article  CAS  PubMed  Google Scholar 

  117. Franceschi, R. T. & Iyer, B. S. Relationship between collagen synthesis and expression of the osteoblast phenotype in MC3T3-E1 cells. J. Bone Min. Res. 7, 235–246 (1992).

    Article  CAS  Google Scholar 

  118. Tenenbaum, H. C. Role of organic phosphate in mineralization of bone in vitro. J. Dent. Res. 60 (Spec. No. C), 1586–1589 (1981).

    Article  Google Scholar 

  119. Eyckmans, J., Lin, G. L. & Chen, C. S. Adhesive and mechanical regulation of mesenchymal stem cell differentiation in human bone marrow and periosteum-derived progenitor cells. Biol. Open 1, 1058–1068 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  120. Komori, T. et al. Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89, 755–764 (1997).

    Article  CAS  PubMed  Google Scholar 

  121. Otto, F. et al. Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89, 765–771 (1997).

    Article  CAS  PubMed  Google Scholar 

  122. Adhami, M. D., Rashid, H., Chen, H. & Javed, A. Runx2 activity in committed osteoblasts is not essential for embryonic skeletogenesis. Connect. Tissue Res. 55 (Suppl. 1), 102–106 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  123. Ducy, P. et al. A Cbfa1-dependent genetic pathway controls bone formation beyond embryonic development. Genes Dev. 13, 1025–1036 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  124. Takarada, T. et al. An analysis of skeletal development in osteoblast-specific and chondrocyte-specific runt-related transcription factor-2 (Runx2) knockout mice. J. Bone Min. Res. 28, 2064–2069 (2013).

    Article  CAS  Google Scholar 

  125. Takeda, S., Bonnamy, J. P., Owen, M. J., Ducy, P. & Karsenty, G. Continuous expression of Cbfa1 in nonhypertrophic chondrocytes uncovers its ability to induce hypertrophic chondrocyte differentiation and partially rescues Cbfa1-deficient mice. Genes Dev. 15, 467–481 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  126. Lee, B. et al. Missense mutations abolishing DNA binding of the osteoblast-specific transcription factor OSF2/CBFA1 in cleidocranial dysplasia. Nat. Genet. 16, 307–310 (1997).

    Article  CAS  PubMed  Google Scholar 

  127. Yoshida, T. et al. Functional analysis of RUNX2 mutations in Japanese patients with cleidocranial dysplasia demonstrates novel genotype-phenotype correlations. Am. J. Hum. Genet. 71, 724–738 (2002).

    Article  PubMed  PubMed Central  Google Scholar 

  128. Meyer, M. B., Benkusky, N. A. & Pike, J. W. The RUNX2 cistrome in osteoblasts: characterization, down-regulation following differentiation, and relationship to gene expression. J. Biol. Chem. 289, 16016–16031 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  129. Wu, H. et al. Genomic occupancy of Runx2 with global expression profiling identifies a novel dimension to control of osteoblastogenesis. Genome Biol. 15, R52 (2014).

    Article  PubMed  PubMed Central  Google Scholar 

  130. Hakelien, A. M. et al. The regulatory landscape of osteogenic differentiation. Stem Cell 32, 2780–2793 (2014).

    Article  CAS  Google Scholar 

  131. Tarkkonen, K., Hieta, R., Kytola, V., Nykter, M. & Kiviranta, R. Comparative analysis of osteoblast gene expression profiles and Runx2 genomic occupancy of mouse and human osteoblasts in vitro. Gene 626, 119–131 (2017).

    Article  CAS  PubMed  Google Scholar 

  132. Yan, J. et al. Smad4 deficiency impairs chondrocyte hypertrophy via the Runx2 transcription factor in mouse skeletal development. J. Biol. Chem. 293, 9162–9175 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  133. Kawane, T. et al. Dlx5 and mef2 regulate a novel runx2 enhancer for osteoblast-specific expression. J. Bone Min. Res. 29, 1960–1969 (2014).

    Article  CAS  Google Scholar 

  134. Robledo, R. F., Rajan, L., Li, X. & Lufkin, T. The Dlx5 and Dlx6 homeobox genes are essential for craniofacial, axial, and appendicular skeletal development. Genes Dev. 16, 1089–1101 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  135. Hojo, H., Ohba, S., He, X., Lai, L. P. & McMahon, A. P. Sp7/Osterix is restricted to bone-forming vertebrates where it acts as a Dlx co-factor in osteoblast specification. Dev. Cell 37, 238–253 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Nakashima, K. et al. The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell 108, 17–29 (2002).

    Article  CAS  PubMed  Google Scholar 

  137. Day, T. F., Guo, X., Garrett-Beal, L. & Yang, Y. Wnt/beta-catenin signaling in mesenchymal progenitors controls osteoblast and chondrocyte differentiation during vertebrate skeletogenesis. Dev. Cell 8, 739–750 (2005).

    Article  CAS  PubMed  Google Scholar 

  138. Hu, H. et al. Sequential roles of Hedgehog and Wnt signaling in osteoblast development. Development 132, 49–60 (2005).

    Article  CAS  PubMed  Google Scholar 

  139. Hill, T. P., Spater, D., Taketo, M. M., Birchmeier, W. & Hartmann, C. Canonical Wnt/beta-catenin signaling prevents osteoblasts from differentiating into chondrocytes. Dev. Cell 8, 727–738 (2005).

    Article  CAS  PubMed  Google Scholar 

  140. Akiyama, H. et al. Interactions between Sox9 and beta-catenin control chondrocyte differentiation. Genes Dev. 18, 1072–1087 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  141. Wu, H. et al. Chromatin dynamics regulate mesenchymal stem cell lineage specification and differentiation to osteogenesis. Biochim. Biophys. Acta Gene Regul. Mech. 1860, 438–449 (2017).

    Article  CAS  PubMed  Google Scholar 

  142. Meyer, M. B., Benkusky, N. A., Sen, B., Rubin, J. & Pike, J. W. Epigenetic plasticity drives adipogenic and osteogenic differentiation of marrow-derived mesenchymal stem cells. J. Biol. Chem. 291, 17829–17847 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  143. Asfour, H. A., Allouh, M. Z. & Said, R. S. Myogenic regulatory factors: the orchestrators of myogenesis after 30 years of discovery. Exp. Biol. Med. 243, 118–128 (2018).

    Article  CAS  Google Scholar 

  144. Chal, J. & Pourquie, O. Making muscle: skeletal myogenesis in vivo and in vitro. Development 144, 2104–2122 (2017).

    Article  CAS  PubMed  Google Scholar 

  145. Gokhin, D. S., Ward, S. R., Bremner, S. N. & Lieber, R. L. Quantitative analysis of neonatal skeletal muscle functional improvement in the mouse. J. Exp. Biol. 211, 837–843 (2008).

    Article  CAS  PubMed  Google Scholar 

  146. Mauro, A. Satellite cell of skeletal muscle fibers. J. Biophys. Biochem. Cytol. 9, 493–495 (1961).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  147. Snow, M. H. Myogenic cell formation in regenerating rat skeletal muscle injured by mincing. II. An autoradiographic study. Anat. Rec. 188, 201–217 (1977).

    Article  CAS  PubMed  Google Scholar 

  148. Biressi, S. et al. Myf5 expression during fetal myogenesis defines the developmental progenitors of adult satellite cells. Dev. Biol. 379, 195–207 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  149. Kostallari, E. et al. Pericytes in the myovascular niche promote post-natal myofiber growth and satellite cell quiescence. Development 142, 1242–1253 (2015).

    CAS  PubMed  Google Scholar 

  150. Dellavalle, A. et al. Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat. Cell Biol. 9, 255–267 (2007).

    Article  CAS  PubMed  Google Scholar 

  151. Mitchell, K. J. et al. Identification and characterization of a non-satellite cell muscle resident progenitor during postnatal development. Nat. Cell Biol. 12, 257–266 (2010).

    Article  CAS  PubMed  Google Scholar 

  152. Dellavalle, A. et al. Pericytes resident in postnatal skeletal muscle differentiate into muscle fibres and generate satellite cells. Nat. Commun. 2, 499 (2011).

    Article  CAS  PubMed  Google Scholar 

  153. Kuang, S., Kuroda, K., Le Grand, F. & Rudnicki, M. A. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell 129, 999–1010 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  154. Webster, M. T., Manor, U., Lippincott-Schwartz, J. & Fan, C. M. Intravital imaging reveals ghost fibers as architectural units guiding myogenic progenitors during regeneration. Cell Stem Cell 18, 243–252 (2016).

    Article  CAS  PubMed  Google Scholar 

  155. Tierney, M. T., Stec, M. J., Rulands, S., Simons, B. D. & Sacco, A. Muscle stem cells exhibit distinct clonal dynamics in response to tissue repair and homeostatic aging. Cell Stem Cell 22, 119–127.e3 (2018).

    Article  CAS  PubMed  Google Scholar 

  156. Dumont, N. A. et al. Dystrophin expression in muscle stem cells regulates their polarity and asymmetric division. Nat. Med. 21, 1455–1463 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  157. Dell’Orso, S. et al. Single cell analysis of adult mouse skeletal muscle stem cells in homeostatic and regenerative conditions. Development 146, dev174177 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  158. De Micheli, A. J. et al. Single-cell analysis of the muscle stem cell hierarchy identifies heterotypic communication signals involved in skeletal muscle regeneration. Cell Rep. 30, 3583–3595.e5 (2020). In this study by De Micheli et al., scRNA-seq of cells from injured muscle revealed the myogenic trajectory during muscle regenertion.

    Article  PubMed  PubMed Central  Google Scholar 

  159. Porpiglia, E. et al. High-resolution myogenic lineage mapping by single-cell mass cytometry. Nat. Cell Biol. 19, 558–567 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  160. Gilbert, P. M. et al. Substrate elasticity regulates skeletal muscle stem cell self-renewal in culture. Science 329, 1078–1081 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  161. Danoviz, M. E. & Yablonka-Reuveni, Z. Skeletal muscle satellite cells: background and methods for isolation and analysis in a primary culture system. Methods Mol. Biol. 798, 21–52 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  162. Magli, A., Schnettler, E., Rinaldi, F., Bremer, P. & Perlingeiro, R. C. Functional dissection of Pax3 in paraxial mesoderm development and myogenesis. Stem Cell 31, 59–70 (2013).

    Article  CAS  Google Scholar 

  163. Hutcheson, D. A., Zhao, J., Merrell, A., Haldar, M. & Kardon, G. Embryonic and fetal limb myogenic cells are derived from developmentally distinct progenitors and have different requirements for beta-catenin. Genes Dev. 23, 997–1013 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  164. Lepper, C., Conway, S. J. & Fan, C. M. Adult satellite cells and embryonic muscle progenitors have distinct genetic requirements. Nature 460, 627–631 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  165. Seale, P. et al. Pax7 is required for the specification of myogenic satellite cells. Cell 102, 777–786 (2000).

    Article  CAS  PubMed  Google Scholar 

  166. Zammit, P. S. et al. Pax7 and myogenic progression in skeletal muscle satellite cells. J. Cell Sci. 119, 1824–1832 (2006).

    Article  CAS  PubMed  Google Scholar 

  167. Blum, R., Vethantham, V., Bowman, C., Rudnicki, M. & Dynlacht, B. D. Genome-wide identification of enhancers in skeletal muscle: the role of MyoD1. Genes Dev. 26, 2763–2779 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  168. Cao, Y. et al. Global and gene-specific analyses show distinct roles for Myod and Myog at a common set of promoters. EMBO J. 25, 502–511 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  169. Conerly, M. L., Yao, Z., Zhong, J. W., Groudine, M. & Tapscott, S. J. Distinct activities of Myf5 and MyoD indicate separate roles in skeletal muscle lineage specification and differentiation. Dev. Cell 36, 375–385 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  170. Fajas, L. et al. The organization, promoter analysis, and expression of the human PPARgamma gene. J. Biol. Chem. 272, 18779–18789 (1997).

    Article  CAS  PubMed  Google Scholar 

  171. Soleimani, V. D. et al. Cis-regulatory determinants of MyoD function. Nucleic Acids Res. 46, 7221–7235 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Soleimani, V. D. et al. Snail regulates MyoD binding-site occupancy to direct enhancer switching and differentiation-specific transcription in myogenesis. Mol. Cell 47, 457–468 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  173. Mok, G. F., Mohammed, R. H. & Sweetman, D. Expression of myogenic regulatory factors in chicken embryos during somite and limb development. J. Anat. 227, 352–360 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  174. Davis, R. L., Weintraub, H. & Lassar, A. B. Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell 51, 987–1000 (1987).

    Article  CAS  PubMed  Google Scholar 

  175. Braun, T., Buschhausen-Denker, G., Bober, E., Tannich, E. & Arnold, H. H. A novel human muscle factor related to but distinct from MyoD1 induces myogenic conversion in 10T1/2 fibroblasts. EMBO J. 8, 701–709 (1989).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  176. Wright, W. E., Sassoon, D. A. & Lin, V. K. Myogenin, a factor regulating myogenesis, has a domain homologous to MyoD. Cell 56, 607–617 (1989).

    Article  CAS  PubMed  Google Scholar 

  177. Rhodes, S. J. & Konieczny, S. F. Identification of MRF4: a new member of the muscle regulatory factor gene family. Genes Dev. 3, 2050–2061 (1989).

    Article  CAS  PubMed  Google Scholar 

  178. Nabeshima, Y. et al. Myogenin gene disruption results in perinatal lethality because of severe muscle defect. Nature 364, 532–535 (1993).

    Article  CAS  PubMed  Google Scholar 

  179. Hasty, P. et al. Muscle deficiency and neonatal death in mice with a targeted mutation in the myogenin gene. Nature 364, 501–506 (1993).

    Article  CAS  PubMed  Google Scholar 

  180. Rudnicki, M. A., Braun, T., Hinuma, S. & Jaenisch, R. Inactivation of MyoD in mice leads to up-regulation of the myogenic HLH gene Myf-5 and results in apparently normal muscle development. Cell 71, 383–390 (1992).

    Article  CAS  PubMed  Google Scholar 

  181. Braun, T., Rudnicki, M. A., Arnold, H. H. & Jaenisch, R. Targeted inactivation of the muscle regulatory gene Myf-5 results in abnormal rib development and perinatal death. Cell 71, 369–382 (1992).

    Article  CAS  PubMed  Google Scholar 

  182. Braun, T. & Arnold, H. H. Inactivation of Myf-6 and Myf-5 genes in mice leads to alterations in skeletal muscle development. EMBO J. 14, 1176–1186 (1995).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  183. Rawls, A. et al. Overlapping functions of the myogenic bHLH genes MRF4 and MyoD revealed in double mutant mice. Development 125, 2349–2358 (1998).

    Article  CAS  PubMed  Google Scholar 

  184. Kassar-Duchossoy, L. et al. Mrf4 determines skeletal muscle identity in Myf5:Myod double-mutant mice. Nature 431, 466–471 (2004).

    Article  CAS  PubMed  Google Scholar 

  185. Valdez, M. R., Richardson, J. A., Klein, W. H. & Olson, E. N. Failure of Myf5 to support myogenic differentiation without myogenin, MyoD, and MRF4. Dev. Biol. 219, 287–298 (2000).

    Article  CAS  PubMed  Google Scholar 

  186. Rudnicki, M. A. et al. MyoD or Myf-5 is required for the formation of skeletal muscle. Cell 75, 1351–1359 (1993).

    Article  CAS  PubMed  Google Scholar 

  187. Tajbakhsh, S. et al. A population of myogenic cells derived from the mouse neural tube. Neuron 13, 813–821 (1994).

    Article  CAS  PubMed  Google Scholar 

  188. Kablar, B. et al. MyoD and Myf-5 define the specification of musculature of distinct embryonic origin. Biochem. Cell Biol. 76, 1079–1091 (1998).

    Article  CAS  PubMed  Google Scholar 

  189. Gensch, N., Borchardt, T., Schneider, A., Riethmacher, D. & Braun, T. Different autonomous myogenic cell populations revealed by ablation of Myf5-expressing cells during mouse embryogenesis. Development 135, 1597–1604 (2008).

    Article  CAS  PubMed  Google Scholar 

  190. Chen, J. C., Mortimer, J., Marley, J. & Goldhamer, D. J. MyoD-cre transgenic mice: a model for conditional mutagenesis and lineage tracing of skeletal muscle. Genesis 41, 116–121 (2005).

    Article  PubMed  Google Scholar 

  191. Haldar, M., Karan, G., Tvrdik, P. & Capecchi, M. R. Two cell lineages, myf5 and myf5-independent, participate in mouse skeletal myogenesis. Dev. Cell 14, 437–445 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  192. Blackburn, D. M. et al. High-resolution genome-wide expression analysis of single myofibers using SMART-Seq. J. Biol. Chem. 294, 20097–20108 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  193. Kanisicak, O., Mendez, J. J., Yamamoto, S., Yamamoto, M. & Goldhamer, D. J. Progenitors of skeletal muscle satellite cells express the muscle determination gene, MyoD. Dev. Biol. 332, 131–141 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  194. Crist, C. G., Montarras, D. & Buckingham, M. Muscle satellite cells are primed for myogenesis but maintain quiescence with sequestration of Myf5 mRNA targeted by microRNA-31 in mRNP granules. Cell Stem Cell 11, 118–126 (2012).

    Article  CAS  PubMed  Google Scholar 

  195. Yamamoto, M. et al. Loss of MyoD and Myf5 in skeletal muscle stem cells results in altered myogenic programming and failed regeneration. Stem Cell Rep. 10, 956–969 (2018).

    Article  CAS  Google Scholar 

  196. Ustanina, S., Carvajal, J., Rigby, P. & Braun, T. The myogenic factor Myf5 supports efficient skeletal muscle regeneration by enabling transient myoblast amplification. Stem Cell 25, 2006–2016 (2007).

    Article  CAS  Google Scholar 

  197. Megeney, L. A., Kablar, B., Garrett, K., Anderson, J. E. & Rudnicki, M. A. MyoD is required for myogenic stem cell function in adult skeletal muscle. Genes Dev. 10, 1173–1183 (1996).

    Article  CAS  PubMed  Google Scholar 

  198. Wang, C. et al. Loss of MyoD promotes fate transdifferentiation of myoblasts into brown adipocytes. EBioMedicine 16, 212–223 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  199. An, Y. et al. A molecular switch regulating cell fate choice between muscle progenitor cells and brown adipocytes. Dev. Cell 41, 382–391.e5 (2017).

    Article  CAS  PubMed  Google Scholar 

  200. Cui, S. et al. β-Catenin is essential for differentiation of primary myoblasts via cooperation with MyoD and α-catenin. Development 146, dev167080 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  201. Peng, X. L. et al. MyoD- and FoxO3-mediated hotspot interaction orchestrates super-enhancer activity during myogenic differentiation. Nucleic Acids Res. 45, 8785–8805 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  202. Lazure, F. et al. Myf6/MRF4 is a myogenic niche regulator required for the maintenance of the muscle stem cell pool. EMBO Rep. 21, e49499 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  203. Constantinides, P. G., Jones, P. A. & Gevers, W. Functional striated muscle cells from non-myoblast precursors following 5-azacytidine treatment. Nature 267, 364–366 (1977).

    Article  CAS  PubMed  Google Scholar 

  204. Blau, H. M. et al. Plasticity of the differentiated state. Science 230, 758–766 (1985).

    Article  CAS  PubMed  Google Scholar 

  205. Weintraub, H. et al. Activation of muscle-specific genes in pigment, nerve, fat, liver, and fibroblast cell lines by forced expression of MyoD. Proc. Natl Acad. Sci. USA 86, 5434–5438 (1989).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  206. Zheng, H., Guo, Z., Ma, Q., Jia, H. & Dang, G. Cbfa1/osf2 transduced bone marrow stromal cells facilitate bone formation in vitro and in vivo. Calcif. Tissue Int. 74, 194–203 (2004).

    Article  CAS  PubMed  Google Scholar 

  207. Gersbach, C. A., Byers, B. A., Pavlath, G. K. & Garcia, A. J. Runx2/Cbfa1 stimulates transdifferentiation of primary skeletal myoblasts into a mineralizing osteoblastic phenotype. Exp. Cell Res. 300, 406–417 (2004).

    Article  CAS  PubMed  Google Scholar 

  208. Byers, B. A., Pavlath, G. K., Murphy, T. J., Karsenty, G. & Garcia, A. J. Cell-type-dependent up-regulation of in vitro mineralization after overexpression of the osteoblast-specific transcription factor Runx2/Cbfal. J. Bone Min. Res. 17, 1931–1944 (2002).

    Article  CAS  Google Scholar 

  209. Lecka-Czernik, B. et al. Inhibition of Osf2/Cbfa1 expression and terminal osteoblast differentiation by PPARγ2. J. Cell Biochem. 74, 357–371 (1999).

    Article  CAS  PubMed  Google Scholar 

  210. Ishay-Ronen, D. et al. Gain fat-lose metastasis: converting invasive breast cancer cells into adipocytes inhibits cancer metastasis. Cancer Cell 35, 17–32.e6 (2019).

    Article  CAS  PubMed  Google Scholar 

  211. Strand, D. W. et al. PPARγ isoforms differentially regulate metabolic networks to mediate mouse prostatic epithelial differentiation. Cell Death Dis. 3, e361 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  212. de la Serna, I. L., Carlson, K. A. & Imbalzano, A. N. Mammalian SWI/SNF complexes promote MyoD-mediated muscle differentiation. Nat. Genet. 27, 187–190 (2001).

    Article  PubMed  Google Scholar 

  213. Gerber, A. N., Klesert, T. R., Bergstrom, D. A. & Tapscott, S. J. Two domains of MyoD mediate transcriptional activation of genes in repressive chromatin: a mechanism for lineage determination in myogenesis. Genes Dev. 11, 436–450 (1997).

    Article  CAS  PubMed  Google Scholar 

  214. Adam, R. C. et al. Pioneer factors govern super-enhancer dynamics in stem cell plasticity and lineage choice. Nature 521, 366–370 (2015). The study by Adam et al. indicates that cell fate switches are associated with the decomissioning of lineage-specifying super-enhancers by the substitution of master regulators, a process that is enhanced by the pioneering activity of the overtaking master regulator.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  215. Kanakasabai, S. et al. PPARγ agonists promote oligodendrocyte differentiation of neural stem cells by modulating stemness and differentiation genes. PLoS ONE 7, e50500 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  216. Mao-Qiang, M. et al. Peroxisome-proliferator-activated receptor (PPAR)-γ activation stimulates keratinocyte differentiation. J. Invest. Dermatol. 123, 305–312 (2004).

    Article  CAS  PubMed  Google Scholar 

  217. Liu, C. et al. Pparg promotes differentiation and regulates mitochondrial gene expression in bladder epithelial cells. Nat. Commun. 10, 4589 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  218. Monemdjou, R. et al. Association of cartilage-specific deletion of peroxisome proliferator-activated receptor gamma with abnormal endochondral ossification and impaired cartilage growth and development in a murine model. Arthritis Rheum. 64, 1551–1561 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  219. Vasheghani, F. et al. PPARγ deficiency results in severe, accelerated osteoarthritis associated with aberrant mTOR signalling in the articular cartilage. Ann. Rheum. Dis. 74, 569–578 (2015).

    Article  CAS  PubMed  Google Scholar 

  220. Norris, A. W. et al. Endogenous peroxisome proliferator-activated receptor-γ augments fatty acid uptake in oxidative muscle. Endocrinology 149, 5374–5383 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  221. Hevener, A. L. et al. Muscle-specific Pparg deletion causes insulin resistance. Nat. Med. 9, 1491–1497 (2003).

    Article  CAS  PubMed  Google Scholar 

  222. Norris, A. W. et al. Muscle-specific PPARγ-deficient mice develop increased adiposity and insulin resistance but respond to thiazolidinediones. J. Clin. Invest. 112, 608–618 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  223. Ren, D., Collingwood, T. N., Rebar, E. J., Wolffe, A. P. & Camp, H. S. PPARγ knockdown by engineered transcription factors: exogenous PPARγ2 but not PPARgamma1 reactivates adipogenesis. Genes Dev. 16, 27–32 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  224. Maruyama, Z. et al. Runx2 determines bone maturity and turnover rate in postnatal bone development and is involved in bone loss in estrogen deficiency. Dev. Dyn. 236, 1876–1890 (2007).

    Article  CAS  PubMed  Google Scholar 

  225. Sugii, S. et al. PPARγ activation in adipocytes is sufficient for systemic insulin sensitization. Proc. Natl Acad. Sci. USA 106, 22504–22509 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  226. Gundersen, K., Rabben, I., Klocke, B. J. & Merlie, J. P. Overexpression of myogenin in muscles of transgenic mice: interaction with Id-1, negative crossregulation of myogenic factors, and induction of extrasynaptic acetylcholine receptor expression. Mol. Cell Biol. 15, 7127–7134 (1995).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  227. Liu, W. et al. Overexpression of Cbfa1 in osteoblasts inhibits osteoblast maturation and causes osteopenia with multiple fractures. J. Cell Biol. 155, 157–166 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  228. Hattori, T. et al. SOX9 is a major negative regulator of cartilage vascularization, bone marrow formation and endochondral ossification. Development 137, 901–911 (2010).

    Article  CAS  PubMed  Google Scholar 

  229. Parfitt, A. M. Osteonal and hemi-osteonal remodeling: the spatial and temporal framework for signal traffic in adult human bone. J. Cell Biochem. 55, 273–286 (1994).

    Article  CAS  PubMed  Google Scholar 

  230. Jilka, R. L., Weinstein, R. S., Bellido, T., Parfitt, A. M. & Manolagas, S. C. Osteoblast programmed cell death (apoptosis): modulation by growth factors and cytokines. J. Bone Min. Res. 13, 793–802 (1998).

    Article  CAS  Google Scholar 

  231. Aigner, T. et al. Apoptotic cell death is not a widespread phenomenon in normal aging and osteoarthritis human articular knee cartilage: a study of proliferation, programmed cell death (apoptosis), and viability of chondrocytes in normal and osteoarthritic human knee cartilage. Arthritis Rheum. 44, 1304–1312 (2001).

    Article  CAS  PubMed  Google Scholar 

  232. Spalding, K. L. et al. Dynamics of fat cell turnover in humans. Nature 453, 783–787 (2008).

    Article  CAS  PubMed  Google Scholar 

  233. Spalding, K. L., Bhardwaj, R. D., Buchholz, B. A., Druid, H. & Frisen, J. Retrospective birth dating of cells in humans. Cell 122, 133–143 (2005).

    Article  CAS  PubMed  Google Scholar 

  234. Madsen, J. G. S. et al. Highly interconnected enhancer communities control lineage-determining genes in human mesenchymal stem cells. Nat. Genet. 52, 1227–1238 (2020). Madsen et al. showed that communities of highly interacting transcriptional enhancers activate lineage-determining genes in human stromal progenitors and that these communities are already formed in the progenitor state.

    Article  CAS  PubMed  Google Scholar 

  235. Kern, S., Eichler, H., Stoeve, J., Kluter, H. & Bieback, K. Comparative analysis of mesenchymal stem cells from bone marrow, umbilical cord blood, or adipose tissue. Stem Cell 24, 1294–1301 (2006).

    Article  CAS  Google Scholar 

  236. Fennema, E. M. et al. Ectopic bone formation by aggregated mesenchymal stem cells from bone marrow and adipose tissue: a comparative study. J. Tissue Eng. Regen. Med. 12, e150–e158 (2018).

    Article  CAS  PubMed  Google Scholar 

  237. Xu, J. et al. Comparison of skeletal and soft tissue pericytes identifies CXCR4+ bone forming mural cells in human tissues. Bone Res. 8, 22 (2020). Xu et al. found that CXCR4-signalling is a niche-specific signal that contributes to pre-programming the in vivo potency of stromal progenitors and how its manipulation shifts lineage potential in transplantation assays.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  238. Chan, C. K. et al. Identification and specification of the mouse skeletal stem cell. Cell 160, 285–298 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  239. Chan, C. K. F. et al. Identification of the human skeletal stem cell. Cell 175, 43–56.e21 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  240. Breschi, A. et al. A limited set of transcriptional programs define major cell types. Genome Res. 30, 1047–1059 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  241. Reinisch, A. et al. Epigenetic and in vivo comparison of diverse MSC sources reveals an endochondral signature for human hematopoietic niche formation. Blood 125, 249–260 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  242. Roson-Burgo, B., Sanchez-Guijo, F., Del Canizo, C. & De Las Rivas, J. Insights into the human mesenchymal stromal/stem cell identity through integrative transcriptomic profiling. BMC Genomics 17, 944 (2016).

    Article  PubMed  PubMed Central  Google Scholar 

  243. Barrio-Hernandez, I. et al. Phosphoproteomic profiling reveals a defined genetic program for osteoblastic lineage commitment of human bone marrow-derived stromal stem cells. Genome Res 30, 127–137 (2019).

    Article  PubMed  Google Scholar 

  244. Bernstein, B. E. et al. A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell 125, 315–326 (2006).

    Article  CAS  PubMed  Google Scholar 

  245. Azuara, V. et al. Chromatin signatures of pluripotent cell lines. Nat. Cell Biol. 8, 532–538 (2006).

    Article  CAS  PubMed  Google Scholar 

  246. Eng, C. L. et al. Transcriptome-scale super-resolved imaging in tissues by RNA seqFISH. Nature 568, 235–239 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  247. Vickovic, S. et al. High-definition spatial transcriptomics for in situ tissue profiling. Nat. Methods 16, 987–990 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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Acknowledgements

The authors would like to thank members from the Mandrup group and the Kassem group for valuable discussions on the topic of this review as well as Rasmus Siersbæk for constructive feedback on the manuscript. The work was supported by grants to A.R. from the Novo Nordisk Foundation (NNF17OC0029290), the Independent Research Fund Denmark (0134-00292B), and the Lundbeck Foundation (R335-2019-2195) and to S.M. from The Danish National Research Foundation to the Center for Functional Genomics and Tissue Plasticity (ATLAS) (Project grant: 141), and the Independent Research Fund Denmark, the Lundbeck Foundation and the Novo Nordisk Foundation.

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Authors and Affiliations

  1. Molecular Endocrinology & Stem Cell Research Unit (KMEB), Department of Endocrinology and Metabolism, Odense University Hospital and Department of Clinical Research, University of Southern Denmark, Odense, Denmark

    Alexander Rauch

  2. Steno Diabetes Center Odense, Odense University Hospital, Odense, Denmark

    Alexander Rauch

  3. Center for Functional Genomics and Tissue Plasticity, Functional Genomics & Metabolism Research Unit, Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark

    Susanne Mandrup

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The authors contributed equally to the discussion, writing and revision of the article.

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Nature Reviews Molecular Cell Biology thanks Jacques Galipeau, Armand Keating and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

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Glossary

Immortalized stromal progenitors

Manipulation of cells to overcome proliferative senescence but does not have to result in transformation.

Genetic driver

Regulatory element of a gene used to control the expression of a transgene in order to recapitulate the expression of the endogenous target.

Neural crest cells

Temporary group of cells that leave from the dorsal part of the neurol tube after its formation.

Hypertrophy

Increase in tissue mass due to the enlargement of cells.

Hyperplasia

Increase in tissue mass due to the proliferation of cells.

Reticular interstitium

A fluid-filled small space in tissues with a specialized extracellular matrix and that is drained by lymph nodes.

Droplet-based single-cell sequencing

Microfluidic device-based compartmentalization of single cells into lysis buffer and barcoded microbeads containing droplets that allows the parallel analysis of thousands of cells.

Single-nucleus RNA-seq

(snRNA-seq). Adaptation of single-cell RNA-seq procedure with isolated nuclei, which offers the recovery of cell types that evade droplet-based sequencing techniques.

Super-enhancers

Groups of enhancers that are in close linear genomic proximity with high levels of enhancer activity. These enhancers control many genes important for lineage determination and cell-type specification.

IMAGE machine learning tool

Modelling approach to predict the contribution of transcription factors to the overall enhancer and gene activity based on the occurrence of all known motifs.

Cartilage anlagen

The initial clustering of cells during embryonic development from which the bones develop.

Growth plate

A disc-like structure of cartilage located near the ends of the bones that permit growth in length.

Articular cartilage

Smooth tissue layer that covers the ends of the bones where they form the joints.

Periosteum

A dense, fibrous cell-containing membrane covering the surface of bones, which is not covered by articular cartilage and which serves as attachment for tendons and muscles.

Epithelial-to-mesenchymal transition

A process by which epithelial cells lose their cell polarity and cell–cell adhesion in order to gain migratory and invasive properties characteristic of the mesenchymal lineage.

Closed chromatin

State of chromatin in which nucleosome density reduces DNA accessibility for interacting proteins, for example, transcription factors.

Lipid droplet

Lipid-rich cellular organelles, also called oil bodies, for the storage and hydrolysis of neutral lipids.

Enhancer capture Hi-C

A method to specifically enrich highly complex Hi-C libraries of DNA fragments containing sequences of the enhancer of interest using biotinylated RNA baits with complementary sequence. Hi-C is a high throughput method to measure pairwise contacts between virtually any genomic region.

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Rauch, A., Mandrup, S. Transcriptional networks controlling stromal cell differentiation. Nat Rev Mol Cell Biol 22, 465–482 (2021). https://doi.org/10.1038/s41580-021-00357-7

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