Cells and tissues generate and are exposed to various mechanical forces that act across a range of scales, from tissues to cells to organelles. Forces provide crucial signals to inform cell behaviour during development and adult tissue homeostasis, and alterations in forces and in their downstream mechanotransduction pathways can influence disease progression. Recent advances have been made in our understanding of the mechanisms by which forces regulate chromatin organization and state, and of the mechanosensitive transcription factors that respond to the physical properties of the cell microenvironment to coordinate gene expression, cell states and behaviours. These insights highlight the relevance of mechanosensitive transcriptional regulation to physiology, disease and emerging therapies.
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Collinet, C. & Lecuit, T. Programmed and self-organized flow of information during morphogenesis. Nat. Rev. Mol. Cell Biol. 22, 245–265 (2021).
Mishra, N. & Heisenberg, C. P. Dissecting organismal morphogenesis by bridging genetics and biophysics. Annu. Rev. Genet. 55, 209–233 (2021).
Stooke-Vaughan, G. A. & Campas, O. Physical control of tissue morphogenesis across scales. Curr. Opin. Genet. Dev. 51, 111–119 (2018).
Janmey, P. A., Fletcher, D. A. & Reinhart-King, C. A. Stiffness sensing by cells. Physiol. Rev. 100, 695–724 (2020).
Murrell, M., Oakes, P. W., Lenz, M. & Gardel, M. L. Forcing cells into shape: the mechanics of actomyosin contractility. Nat. Rev. Mol. Cell Biol. 16, 486–498 (2015).
Vining, K. H. & Mooney, D. J. Mechanical forces direct stem cell behaviour in development and regeneration. Nat. Rev. Mol. Cell Biol. 18, 728–742 (2017).
Vogel, V. Unraveling the mechanobiology of extracellular matrix. Annu. Rev. Physiol. 80, 353–387 (2018).
Tschumperlin, D. J., Ligresti, G., Hilscher, M. B. & Shah, V. H. Mechanosensing and fibrosis. J. Clin. Invest. 128, 74–84 (2018).
Humphrey, J. D. & Schwartz, M. A. Vascular mechanobiology: homeostasis, adaptation, and disease. Annu. Rev. Biomed. Eng. 23, 1–27 (2021).
Maurer, M. & Lammerding, J. The driving force: nuclear mechanotransduction in cellular function, fate, and disease. Annu. Rev. Biomed. Eng. 21, 443–468 (2019).
Charras, G. & Yap, A. S. Tensile forces and mechanotransduction at cell-cell junctions. Curr. Biol. 28, R445–R457 (2018).
Iskratsch, T., Wolfenson, H. & Sheetz, M. P. Appreciating force and shape — the rise of mechanotransduction in cell biology. Nat. Rev. Mol. Cell Biol. 15, 825–833 (2014).
Kechagia, J. Z., Ivaska, J. & Roca-Cusachs, P. Integrins as biomechanical sensors of the microenvironment. Nat. Rev. Mol. Cell Biol. 20, 457–473 (2019).
Hu, X., Margadant, F. M., Yao, M. & Sheetz, M. P. Molecular stretching modulates mechanosensing pathways. Protein Sci. 26, 1337–1351 (2017).
Saini, K. & Discher, D. E. Forced unfolding of proteins directs biochemical cascades. Biochemistry 58, 4893–4902 (2019).
Cox, C. D., Bavi, N. & Martinac, B. Origin of the force: the force-from-lipids principle applied to piezo channels. Curr. Top. Membr. 79, 59–96 (2017).
Douguet, D. & Honore, E. Mammalian mechanoelectrical transduction: structure and function of force-gated ion channels. Cell 179, 340–354 (2019).
Murthy, S. E., Dubin, A. E. & Patapoutian, A. Piezos thrive under pressure: mechanically activated ion channels in health and disease. Nat. Rev. Mol. Cell Biol. 18, 771–783 (2017).
De Belly, H. et al. Membrane tension gates ERK-mediated regulation of pluripotent cell fate. Cell Stem Cell 28, 273–284.e276 (2021).
Sinha, B. et al. Cells respond to mechanical stress by rapid disassembly of caveolae. Cell 144, 402–413 (2011).
Torrino, S. et al. EHD2 is a mechanotransducer connecting caveolae dynamics with gene transcription. J. Cell Biol. 217, 4092–4105 (2018).
Discher, D. E. et al. Matrix mechanosensing: from scaling concepts in ‘omics data to mechanisms in the nucleus, regeneration, and cancer. Annu. Rev. Biophys. 46, 295–315 (2017).
Miroshnikova, Y. A. & Wickstrom, S. A. Mechanical forces in nuclear organization. Cold Spring Harb. Perspect. Biol. 14, a039685 (2022).
Buxboim, A. et al. Matrix elasticity regulates lamin-A,C phosphorylation and turnover with feedback to actomyosin. Curr. Biol. 24, 1909–1917 (2014).
Cho, S. et al. Mechanosensing by the lamina protects against nuclear rupture, DNA damage, and cell-cycle arrest. Dev. Cell 49, 920–935 e925 (2019).
Lammerding, J. et al. Abnormal nuclear shape and impaired mechanotransduction in emerin-deficient cells. J. Cell Biol. 170, 781–791 (2005).
Lammerding, J. et al. Lamin A/C deficiency causes defective nuclear mechanics and mechanotransduction. J. Clin. Invest. 113, 370–378 (2004).
Swift, J. et al. Nuclear lamin-A scales with tissue stiffness and enhances matrix-directed differentiation. Science 341, 1240104 (2013).
Guelen, L. et al. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453, 948–951 (2008).
van Steensel, B. & Belmont, A. S. Lamina-associated domains: links with chromosome architecture, heterochromatin, and gene repression. Cell 169, 780–791 (2017).
Gesson, K. et al. A-type lamins bind both hetero- and euchromatin, the latter being regulated by lamina-associated polypeptide 2 alpha. Genome Res. 26, 462–473 (2016).
Gruenbaum, Y. & Foisner, R. Lamins: nuclear intermediate filament proteins with fundamental functions in nuclear mechanics and genome regulation. Annu. Rev. Biochem. 84, 131–164 (2015).
Lund, E. et al. Lamin A/C-promoter interactions specify chromatin state-dependent transcription outcomes. Genome Res. 23, 1580–1589 (2013).
Buxboim, A. et al. Coordinated increase of nuclear tension and lamin-A with matrix stiffness outcompetes lamin-B receptor that favors soft tissue phenotypes. Mol. Biol. Cell 28, 3333–3348 (2017).
Ivanovska, I. L. et al. Cross-linked matrix rigidity and soluble retinoids synergize in nuclear lamina regulation of stem cell differentiation. Mol. Biol. Cell 28, 2010–2022 (2017).
Nava, M. M. et al. Heterochromatin-driven nuclear softening protects the genome against mechanical stress-induced damage. Cell 181, 800–817.e822 (2020). This study shows that chromatin dynamically adapts to changing force environments by altering nuclear mechanical properties and that this has a role in genome mechanoprotection.
Stephens, A. D., Banigan, E. J., Adam, S. A., Goldman, R. D. & Marko, J. F. Chromatin and lamin A determine two different mechanical response regimes of the cell nucleus. Mol. Biol. Cell 28, 1984–1996 (2017).
Rheinlaender, J. et al. Cortical cell stiffness is independent of substrate mechanics. Nat. Mater. 19, 1019–1025 (2020).
Enyedi, B., Jelcic, M. & Niethammer, P. The cell nucleus serves as a mechanotransducer of tissue damage-induced inflammation. Cell 165, 1160–1170 (2016). This study was among the first to demonstrate that the nucleus is a mechanosensor and to identify calcium as a downstream signalling mediator.
Itano, N., Okamoto, S., Zhang, D., Lipton, S. A. & Ruoslahti, E. Cell spreading controls endoplasmic and nuclear calcium: a physical gene regulation pathway from the cell surface to the nucleus. Proc. Natl Acad. Sci. USA 100, 5181–5186 (2003).
Lomakin, A. J et al. The nucleus acts as a ruler tailoring cell responses to spatial constraints. Science 370, eaba2894 (2020).
Venturini, V. et al. The nucleus measures shape changes for cellular proprioception to control dynamic cell behavior. Science 370, eaba2644 (2020).
Aureille, J. et al. Nuclear envelope deformation controls cell cycle progression in response to mechanical force. EMBO Rep. 20, e48084 (2019).
Elosegui-Artola, A. et al. Force triggers YAP nuclear entry by regulating transport across nuclear pores. Cell 171, 1397–1410 e1314 (2017).
Hoffman, L. M. et al. Mechanical stress triggers nuclear remodeling and the formation of transmembrane actin nuclear lines with associated nuclear pore complexes. Mol. Biol. Cell 31, 1774–1787 (2020).
Zimmerli, C. E. et al. Nuclear pores dilate and constrict in cellulo. Science 374, eabd9776 (2021).
Guet, D. et al. Mechanical role of actin dynamics in the rheology of the Golgi complex and in Golgi-associated trafficking events. Curr. Biol. 24, 1700–1711 (2014).
Helle, S. C. J. et al. Mechanical force induces mitochondrial fission. eLife 6, e30292 (2017).
Romani, P. et al. Extracellular matrix mechanical cues regulate lipid metabolism through lipin-1 and SREBP. Nat. Cell Biol. 21, 338–347 (2019).
Romani, P. et al. Mitochondrial fission links ECM mechanotransduction to metabolic redox homeostasis and metastatic chemotherapy resistance. Nat. Cell Biol. 24, 168–180 (2022).
Tharp, K. M. et al. Adhesion-mediated mechanosignaling forces mitohormesis. Cell Metab. 33, 1322–1341 e1313 (2021).
Spasic, M. & Jacobs, C. R. Primary cilia: cell and molecular mechanosensors directing whole tissue function. Semin. Cell Dev. Biol. 71, 42–52 (2017).
Sun, J., Chen, J., Mohagheghian, E. & Wang, N. Force-induced gene up-regulation does not follow the weak power law but depends on H3K9 demethylation. Sci. Adv. 6, eaay9095 (2020).
Tajik, A. et al. Transcription upregulation via force-induced direct stretching of chromatin. Nat. Mater. 15, 1287–1296 (2016).
Mardaryev, A. N. et al. p63 and Brg1 control developmentally regulated higher-order chromatin remodelling at the epidermal differentiation complex locus in epidermal progenitor cells. Development 141, 101–111 (2014).
Carley, E. et al. The LINC complex transmits integrin-dependent tension to the nuclear lamina and represses epidermal differentiation. eLife 10, e58541 (2021).
Heo, S. J. et al. Differentiation alters stem cell nuclear architecture, mechanics, and mechano-sensitivity. eLife 5, e18207 (2016).
Heo, S. J. et al. Mechanically induced chromatin condensation requires cellular contractility in mesenchymal stem cells. Biophys. J. 111, 864–874 (2016).
Le, H. Q. et al. Mechanical regulation of transcription controls Polycomb-mediated gene silencing during lineage commitment. Nat. Cell Biol. 18, 864–875 (2016). This pioneering study shows how mechanical stress alters chromatin architecture with direct impact on lineage gene expression and stem cell differentiation.
Miroshnikova, Y. A., Cohen, I., Ezhkova, E. & Wickstrom, S. A. Epigenetic gene regulation, chromatin structure, and force-induced chromatin remodelling in epidermal development and homeostasis. Curr. Opin. Genet. Dev. 55, 46–51 (2019).
Stephens, A. D. et al. Physicochemical mechanotransduction alters nuclear shape and mechanics via heterochromatin formation. Mol. Biol. Cell 30, 2320–2330 (2019).
Le Beyec, J. et al. Cell shape regulates global histone acetylation in human mammary epithelial cells. Exp. Cell Res. 313, 3066–3075 (2007).
Li, Y. et al. Biophysical regulation of histone acetylation in mesenchymal stem cells. Biophys. J. 100, 1902–1909 (2011).
Jain, N., Iyer, K. V., Kumar, A. & Shivashankar, G. V. Cell geometric constraints induce modular gene-expression patterns via redistribution of HDAC3 regulated by actomyosin contractility. Proc. Natl Acad. Sci. USA 110, 11349–11354 (2013).
Lee, J. et al. Geometric regulation of histone state directs melanoma reprogramming. Commun. Biol. 3, 341 (2020).
Fasciani, A. et al. MLL4-associated condensates counterbalance Polycomb-mediated nuclear mechanical stress in Kabuki syndrome. Nat. Genet. 52, 1397–1411 (2020).
Shao, X., Li, Q., Mogilner, A., Bershadsky, A. D. & Shivashankar, G. V. Mechanical stimulation induces formin-dependent assembly of a perinuclear actin rim. Proc. Natl Acad. Sci. USA 112, E2595–E2601 (2015).
Wales, P. et al. Calcium-mediated actin reset (CaAR) mediates acute cell adaptations. eLife 5, e19850 (2016).
Jung, H. J. et al. An absence of nuclear lamins in keratinocytes leads to ichthyosis, defective epidermal barrier function, and intrusion of nuclear membranes and endoplasmic reticulum into the nuclear chromatin. Mol. Cell Biol. 34, 4534–4544 (2014).
Nastaly, P. et al. Role of the nuclear membrane protein emerin in front-rear polarity of the nucleus. Nat. Commun. 11, 2122 (2020).
Johnson, N. et al. Actin-filled nuclear invaginations indicate degree of cell de-differentiation. Differentiation 71, 414–424 (2003).
Biedzinski, S. et al. Microtubules control nuclear shape and gene expression during early stages of hematopoietic differentiation. EMBO J. 39, e103957 (2020).
Procter, D. J., Furey, C., Garza-Gongora, A. G., Kosak, S. T. & Walsh, D. Cytoplasmic control of intranuclear polarity by human cytomegalovirus. Nature 587, 109–114 (2020).
Ulferts, S., Prajapati, B., Grosse, R. & Vartiainen, M. K. Emerging properties and functions of actin and actin filaments inside the nucleus. Cold Spring Harb. Perspect. Biol. 13, a040121 (2021).
Dopie, J., Skarp, K. P., Rajakyla, E. K., Tanhuanpaa, K. & Vartiainen, M. K. Active maintenance of nuclear actin by importin 9 supports transcription. Proc. Natl Acad. Sci. USA 109, E544–E552 (2012).
Plessner, M., Melak, M., Chinchilla, P., Baarlink, C. & Grosse, R. Nuclear F-actin formation and reorganization upon cell spreading. J. Biol. Chem. 290, 11209–11216 (2015).
Fiore, A. et al. Laminin-111 and the level of nuclear actin regulate epithelial quiescence via exportin-6. Cell Rep. 19, 2102–2115 (2017).
Hofmann, W. A. et al. Actin is part of pre-initiation complexes and is necessary for transcription by RNA polymerase II. Nat. Cell Biol. 6, 1094–1101 (2004).
Philimonenko, V. V. et al. Nuclear actin and myosin I are required for RNA polymerase I transcription. Nat. Cell Biol. 6, 1165–1172 (2004).
Qi, T. et al. G-actin participates in RNA polymerase II-dependent transcription elongation by recruiting positive transcription elongation factor b (P-TEFb). J. Biol. Chem. 286, 15171–15181 (2011).
Sokolova, M. et al. Nuclear actin is required for transcription during drosophila oogenesis. iScience 9, 63–70 (2018).
Viita, T. et al. Nuclear actin interactome analysis links actin to KAT14 histone acetyl transferase and mRNA splicing. J. Cell Sci. 132, jcs226852 (2019).
Spencer, V. A. et al. Depletion of nuclear actin is a key mediator of quiescence in epithelial cells. J. Cell Sci. 124, 123–132 (2011).
Hurst, V., Shimada, K. & Gasser, S. M. Nuclear actin and actin-binding proteins in DNA repair. Trends Cell Biol. 29, 462–476 (2019).
Jungblut, A., Hopfner, K. P. & Eustermann, S. Megadalton chromatin remodelers: common principles for versatile functions. Curr. Opin. Struct. Biol. 64, 134–144 (2020).
Xie, X. et al. β-Actin-dependent global chromatin organization and gene expression programs control cellular identity. FASEB J. 32, 1296–1314 (2018).
Xie, X., Jankauskas, R., Mazari, A. M. A., Drou, N. & Percipalle, P. β-Actin regulates a heterochromatin landscape essential for optimal induction of neuronal programs during direct reprograming. PLoS Genet. 14, e1007846 (2018).
Zhao, K. et al. Rapid and phosphoinositol-dependent binding of the SWI/SNF-like BAF complex to chromatin after T lymphocyte receptor signaling. Cell 95, 625–636 (1998).
Hari-Gupta, Y. et al. Myosin VI regulates the spatial organisation of mammalian transcription initiation. Nat. Commun. 13, 1346 (2022).
Vreugde, S. et al. Nuclear myosin VI enhances RNA polymerase II-dependent transcription. Mol. Cell 23, 749–755 (2006).
Wu, X. et al. Regulation of RNA-polymerase-II-dependent transcription by N-WASP and its nuclear-binding partners. Nat. Cell Biol. 8, 756–763 (2006).
Xia, P. et al. WASH is required for the differentiation commitment of hematopoietic stem cells in a c-Myc-dependent manner. J. Exp. Med. 211, 2119–2134 (2014).
Yoo, Y., Wu, X. & Guan, J. L. A novel role of the actin-nucleating Arp2/3 complex in the regulation of RNA polymerase II-dependent transcription. J. Biol. Chem. 282, 7616–7623 (2007).
Zorca, C. E. et al. Myosin VI regulates gene pairing and transcriptional pause release in T cells. Proc. Natl Acad. Sci. USA 112, E1587–E1593 (2015).
Miyamoto, K., Pasque, V., Jullien, J. & Gurdon, J. B. Nuclear actin polymerization is required for transcriptional reprogramming of Oct4 by oocytes. Genes Dev. 25, 946–958 (2011).
Miyamoto, K. et al. Nuclear Wave1 is required for reprogramming transcription in oocytes and for normal development. Science 341, 1002–1005 (2013). This work provides evidence that nuclear actin polymerization is key for transcription in the context of pluripotency reprogramming.
Yamazaki, S. et al. The actin-family protein arp4 is a novel suppressor for the formation and functions of nuclear F-actin. Cells 9, 758 (2020).
Dalton, S. & Treisman, R. Characterization of SAP-1, a protein recruited by serum response factor to the c-fos serum response element. Cell 68, 597–612 (1992).
Esnault, C. et al. Rho-actin signaling to the MRTF coactivators dominates the immediate transcriptional response to serum in fibroblasts. Genes Dev. 28, 943–958 (2014).
Hill, C. S., Wynne, J. & Treisman, R. The Rho family GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell 81, 1159–1170 (1995). This landmark study shows that SRF activity is regulated in response to cytoskeletal dynamics.
Norman, C., Runswick, M., Pollock, R. & Treisman, R. Isolation and properties of cDNA clones encoding SRF, a transcription factor that binds to the c-fos serum response element. Cell 55, 989–1003 (1988).
Wang, D. Z. et al. Potentiation of serum response factor activity by a family of myocardin-related transcription factors. Proc. Natl Acad. Sci. USA 99, 14855–14860 (2002).
Wang, Z. et al. Myocardin and ternary complex factors compete for SRF to control smooth muscle gene expression. Nature 428, 185–189 (2004).
Miralles, F., Posern, G., Zaromytidou, A. I. & Treisman, R. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113, 329–342 (2003).
Baarlink, C., Wang, H. & Grosse, R. Nuclear actin network assembly by formins regulates the SRF coactivator MAL. Science 340, 864–867 (2013).
Vartiainen, M. K., Guettler, S., Larijani, B. & Treisman, R. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science 316, 1749–1752 (2007).
Cui, Y. et al. Cyclic stretching of soft substrates induces spreading and growth. Nat. Commun. 6, 6333 (2015).
Fearing, B. V. et al. Mechanosensitive transcriptional coactivators MRTF-A and YAP/TAZ regulate nucleus pulposus cell phenotype through cell shape. FASEB J. 33, 14022–14035 (2019).
Foster, C. T., Gualdrini, F. & Treisman, R. Mutual dependence of the MRTF-SRF and YAP-TEAD pathways in cancer-associated fibroblasts is indirect and mediated by cytoskeletal dynamics. Genes. Dev. 31, 2361–2375 (2017).
O’Connor, J. W. & Gomez, E. W. Cell adhesion and shape regulate TGF-beta1-induced epithelial-myofibroblast transition via MRTF-A signaling. PLoS ONE 8, e83188 (2013).
O’Connor, J. W., Riley, P. N., Nalluri, S. M., Ashar, P. K. & Gomez, E. W. Matrix rigidity mediates TGFbeta1-induced epithelial-myofibroblast transition by controlling cytoskeletal organization and MRTF-A localization. J. Cell Physiol. 230, 1829–1839 (2015).
Olson, E. N. & Nordheim, A. Linking actin dynamics and gene transcription to drive cellular motile functions. Nat. Rev. Mol. Cell Biol. 11, 353–365 (2010).
Taskinen, M. E. et al. MASTL promotes cell contractility and motility through kinase- independent signaling. J. Cell Biol. 219, e201906204 (2020).
Aragona, M. et al. Mechanisms of stretch-mediated skin expansion at single-cell resolution. Nature 584, 268–273 (2020). This is one of the first studies to address the in vivo response of a tissue to mechanical perturbation using single-cell sequencing analyses and lineage tracing.
Koegel, H. et al. Loss of serum response factor in keratinocytes results in hyperproliferative skin disease in mice. J. Clin. Invest. 119, 899–910 (2009).
Luxenburg, C., Pasolli, H. A., Williams, S. E. & Fuchs, E. Developmental roles for Srf, cortical cytoskeleton and cell shape in epidermal spindle orientation. Nat. Cell Biol. 13, 203–214 (2011).
Verdoni, A. M., Ikeda, S. & Ikeda, A. Serum response factor is essential for the proper development of skin epithelium. Mamm. Genome 21, 64–76 (2010).
Verdoni, A. M. et al. A pathogenic relationship between a regulator of the actin cytoskeleton and serum response factor. Genetics 186, 147–157 (2010).
Hinkel, R. et al. MRTF-A controls vessel growth and maturation by increasing the expression of CCN1 and CCN2. Nat. Commun. 5, 3970 (2014).
Weinl, C. et al. Endothelial depletion of murine SRF/MRTF provokes intracerebral hemorrhagic stroke. Proc. Natl Acad. Sci. USA 112, 9914–9919 (2015).
Weinl, C. et al. Endothelial SRF/MRTF ablation causes vascular disease phenotypes in murine retinae. J. Clin. Invest. 123, 2193–2206 (2013).
Connelly, J. T. et al. Actin and serum response factor transduce physical cues from the microenvironment to regulate epidermal stem cell fate decisions. Nat. Cell Biol. 12, 711–718 (2010).
Gualdrini, F. et al. SRF co-factors control the balance between cell proliferation and contractility. Mol. Cell 64, 1048–1061 (2016).
Watt, F. M., Jordan, P. W. & O’Neill, C. H. Cell shape controls terminal differentiation of human epidermal keratinocytes. Proc. Natl Acad. Sci. USA 85, 5576–5580 (1988). This work pioneers micropatterning to control cell geometry and mechanics and makes a strong case for mechanical control of cell differentiation, 10 years ahead of the field.
Wozniak, M. A. et al. Adhesion regulates MAP kinase/ternary complex factor exchange to control a proliferative transcriptional switch. Curr. Biol. 22, 2017–2026 (2012).
Gaut, L. et al. EGR1 regulates transcription downstream of mechanical signals during tendon formation and healing. PLoS ONE 11, e0166237 (2016).
Joshi, B. et al. Phosphocaveolin-1 is a mechanotransducer that induces caveola biogenesis via Egr1 transcriptional regulation. J. Cell Biol. 199, 425–435 (2012).
Morawietz, H. et al. Rapid induction and translocation of Egr-1 in response to mechanical strain in vascular smooth muscle cells. Circ. Res. 84, 678–687 (1999).
Dupont, S. Role of YAP/TAZ in cell-matrix adhesion-mediated signalling and mechanotransduction. Exp. Cell Res. 343, 42–53 (2016).
Dupont, S. et al. Role of YAP/TAZ in mechanotransduction. Nature 474, 179–183 (2011). This article describes discovery of what turned out to be a universal mechanoresponsive nuclear relay system.
Pocaterra, A., Romani, P. & Dupont, S. YAP/TAZ functions and their regulation at a glance. J. Cell Sci. 133, jcs230425 (2020).
Aragona, M. et al. A mechanical checkpoint controls multicellular growth through YAP/TAZ regulation by actin-processing factors. Cell 154, 1047–1059 (2013).
Benham-Pyle, B. W., Pruitt, B. L. & Nelson, W. J. Cell adhesion. Mechanical strain induces E-cadherin-dependent Yap1 and beta-catenin activation to drive cell cycle entry. Science 348, 1024–1027 (2015).
Zhao, B. et al. Inactivation of YAP oncoprotein by the Hippo pathway is involved in cell contact inhibition and tissue growth control. Genes Dev. 21, 2747–2761 (2007).
Baker, N. E. Emerging mechanisms of cell competition. Nat. Rev. Genet. 21, 683–697 (2020).
Chen, C. L., Schroeder, M. C., Kango-Singh, M., Tao, C. & Halder, G. Tumor suppression by cell competition through regulation of the Hippo pathway. Proc. Natl Acad. Sci. USA 109, 484–489 (2012).
Ishihara, E. et al. Prostaglandin E2 and its receptor EP2 trigger signaling that contributes to YAP-mediated cell competition. Genes. Cell 25, 197–214 (2020).
Levayer, R. Solid stress, competition for space and cancer: The opposing roles of mechanical cell competition in tumour initiation and growth. Semin. Cancer Biol. 63, 69–80 (2020).
Liu, Z. et al. Differential YAP expression in glioma cells induces cell competition and promotes tumorigenesis. J. Cell Sci. 132, jcs225714 (2019).
Moya, I. M. et al. Peritumoral activation of the Hippo pathway effectors YAP and TAZ suppresses liver cancer in mice. Science 366, 1029–1034 (2019).
Nishio, M. et al. Hippo pathway controls cell adhesion and context-dependent cell competition to influence skin engraftment efficiency. FASEB J. 33, 5548–5560 (2019).
Price, C. J. et al. Genetically variant human pluripotent stem cells selectively eliminate wild-type counterparts through YAP-mediated cell competition. Dev. Cell 56, 2455–2470 e2410 (2021).
Dupont, S. & Morsut, L. Tissue patterning: the winner takes it all, the losers standing small. Curr. Biol. 29, R334–R337 (2019).
Xia, P., Gutl, D., Zheden, V. & Heisenberg, C. P. Lateral inhibition in cell specification mediated by mechanical signals modulating TAZ activity. Cell 176, 1379–1392 e1314 (2019).
Elosegui-Artola, A. et al. Mechanical regulation of a molecular clutch defines force transmission and transduction in response to matrix rigidity. Nat. Cell Biol. 18, 540–548 (2016).
Wada, K., Itoga, K., Okano, T., Yonemura, S. & Sasaki, H. Hippo pathway regulation by cell morphology and stress fibers. Development 138, 3907–3914 (2011).
Meng, Z. et al. RAP2 mediates mechanoresponses of the Hippo pathway. Nature 560, 655–660 (2018).
Chang, L. et al. The SWI/SNF complex is a mechanoregulated inhibitor of YAP and TAZ. Nature 563, 265–269 (2018).
Shiu, J. Y., Aires, L., Lin, Z. & Vogel, V. Nanopillar force measurements reveal actin-cap-mediated YAP mechanotransduction. Nat. Cell Biol. 20, 262–271 (2018).
Engler, A. J. et al. Myotubes differentiate optimally on substrates with tissue-like stiffness: pathological implications for soft or stiff microenvironments. J. Cell Biol. 166, 877–887 (2004).
Engler, A. J., Sen, S., Sweeney, H. L. & Discher, D. E. Matrix elasticity directs stem cell lineage specification. Cell 126, 677–689 (2006). This article describes how tissue mechanical properties can serve as a template to match stem cell differentiation with tissue function.
Gilbert, P. M. et al. Substrate elasticity regulates skeletal muscle stem cell self-renewal in culture. Science 329, 1078–1081 (2010).
Urciuolo, A. et al. Collagen VI regulates satellite cell self-renewal and muscle regeneration. Nat. Commun. 4, 1964 (2013).
Cantini, M., Donnelly, H., Dalby, M. J. & Salmeron-Sanchez, M. The plot thickens: the emerging role of matrix viscosity in cell mechanotransduction. Adv. Healthc. Mater. 9, e1901259 (2020).
Chaudhuri, O. et al. Substrate stress relaxation regulates cell spreading. Nat. Commun. 6, 6364 (2015).
Elosegui-Artola, A. The extracellular matrix viscoelasticity as a regulator of cell and tissue dynamics. Curr. Opin. Cell Biol. 72, 10–18 (2021).
Donat, S. et al. Heg1 and Ccm1/2 proteins control endocardial mechanosensitivity during zebrafish valvulogenesis. eLife 7, e28939 (2018).
Lerche, M. et al. Integrin binding dynamics modulate ligand-specific mechanosensing in mammary gland fibroblasts. iScience 23, 100907 (2020).
Pocaterra, A. et al. F-actin dynamics regulates mammalian organ growth and cell fate maintenance. J. Hepatol. 71, 130–142 (2019). The study reports a physiological role for actomyosin contractility in regulating adult organ size homeostasis, cell fate maintenance and metabolism in a mammalian system.
Pocaterra, A. et al. Fascin1 empowers YAP mechanotransduction and promotes cholangiocarcinoma development. Commun. Biol. 4, 763 (2021).
Sansores-Garcia, L. et al. Modulating F-actin organization induces organ growth by affecting the Hippo pathway. EMBO J. 30, 2325–2335 (2011).
Wang, S. et al. CCM3 is a gatekeeper in focal adhesions regulating mechanotransduction and YAP/TAZ signalling. Nat. Cell Biol. 23, 758–770 (2021).
Calvo, F. et al. Mechanotransduction and YAP-dependent matrix remodelling is required for the generation and maintenance of cancer-associated fibroblasts. Nat. Cell Biol. 15, 637–646 (2013). This work demonstrates that YAP mechanotransduction feeds back on ECM remodelling and how this is relevant in the context of cancer-associated fibrosis.
Link, P. A. et al. Combined control of the fibroblast contractile program by YAP and TAZ. Am. J. Physiol. Lung Cell Mol. Physiol. 322, L23–L32 (2022).
Pagliari, S. et al. YAP-TEAD1 control of cytoskeleton dynamics and intracellular tension guides human pluripotent stem cell mesoderm specification. Cell Death Differ. 28, 1193–1207 (2021).
Porazinski, S. et al. YAP is essential for tissue tension to ensure vertebrate 3D body shape. Nature 521, 217–221 (2015).
Romani, P., Valcarcel-Jimenez, L., Frezza, C. & Dupont, S. Crosstalk between mechanotransduction and metabolism. Nat. Rev. Mol. Cell Biol. 22, 22–38 (2021).
Farge, E. Mechanical induction of Twist in the Drosophila foregut/stomodeal primordium. Curr. Biol. 13, 1365–1377 (2003).
Desprat, N., Supatto, W., Pouille, P. A., Beaurepaire, E. & Farge, E. Tissue deformation modulates twist expression to determine anterior midgut differentiation in Drosophila embryos. Dev. Cell 15, 470–477 (2008). The study demonstrates how tissue deformation can inform cell fate by regulating a TF during embryogenesis.
Roper, J. C. et al. The major beta-catenin/E-cadherin junctional binding site is a primary molecular mechano-transductor of differentiation in vivo. eLife 7, e33381 (2018).
Nusse, R. & Clevers, H. Wnt/beta-catenin signaling, disease, and emerging therapeutic modalities. Cell 169, 985–999 (2017).
Doumpas, N. et al. TCF/LEF dependent and independent transcriptional regulation of Wnt/beta-catenin target genes. EMBO J. 38, e98873 (2019).
Stadeli, R., Hoffmans, R. & Basler, K. Transcription under the control of nuclear Arm/beta-catenin. Curr. Biol. 16, R378–R385 (2006).
Brunet, T. et al. Evolutionary conservation of early mesoderm specification by mechanotransduction in Bilateria. Nat. Commun. 4, 2821 (2013).
Fernandez-Sanchez, M. E. et al. Mechanical induction of the tumorigenic beta-catenin pathway by tumour growth pressure. Nature 523, 92–95 (2015).
Samuel, M. S. et al. Actomyosin-mediated cellular tension drives increased tissue stiffness and beta-catenin activation to induce epidermal hyperplasia and tumor growth. Cancer Cell 19, 776–791 (2011).
Muncie, J. M. et al. Mechanical tension promotes formation of gastrulation-like nodes and patterns mesoderm specification in human embryonic stem cells. Dev. Cell 55, 679–694 e611 (2020).
Przybyla, L., Lakins, J. N. & Weaver, V. M. Tissue mechanics orchestrate wnt-dependent human embryonic stem cell differentiation. Cell Stem Cell 19, 462–475 (2016).
Ray, S., Foote, H. P. & Lechler, T. β-Catenin protects the epidermis from mechanical stresses. J. Cell Biol. 202, 45–52 (2013).
Gayrard, C., Bernaudin, C., Dejardin, T., Seiler, C. & Borghi, N. Src- and confinement-dependent FAK activation causes E-cadherin relaxation and beta-catenin activity. J. Cell Biol. 217, 1063–1077 (2018).
Miroshnikova, Y. A. et al. Calcium signaling mediates a biphasic mechanoadaptive response of endothelial cells to cyclic mechanical stretch. Mol. Biol. Cell 32, 1724–1736 (2021).
Kono, K., Tamashiro, D. A. & Alarcon, V. B. Inhibition of RHO-ROCK signaling enhances ICM and suppresses TE characteristics through activation of Hippo signaling in the mouse blastocyst. Dev. Biol. 394, 142–155 (2014).
Maitre, J. L. et al. Asymmetric division of contractile domains couples cell positioning and fate specification. Nature 536, 344–348 (2016). This study shows how YAP/TAZ activity is patterned by forces and the relevance of contractile asymmetry during early mammalian embryogenesis.
Mihajlovic, A. I. & Bruce, A. W. Rho-associated protein kinase regulates subcellular localisation of Angiomotin and Hippo-signalling during preimplantation mouse embryo development. Reprod. Biomed. Online 33, 381–390 (2016).
Nishioka, N. et al. The Hippo signaling pathway components Lats and Yap pattern Tead4 activity to distinguish mouse trophectoderm from inner cell mass. Dev. Cell 16, 398–410 (2009).
Royer, C. et al. Establishment of a relationship between blastomere geometry and YAP localisation during compaction. Development 147, dev189449 (2020).
Wang, X. et al. Characterizing inner pressure and stiffness of trophoblast and inner cell mass of blastocysts. Biophys. J. 115, 2443–2450 (2018).
Morin-Kensicki, E. M. et al. Defects in yolk sac vasculogenesis, chorioallantoic fusion, and embryonic axis elongation in mice with targeted disruption of Yap65. Mol. Cell Biol. 26, 77–87 (2006).
Hossain, Z. et al. Glomerulocystic kidney disease in mice with a targeted inactivation of Wwtr1. Proc. Natl Acad. Sci. USA 104, 1631–1636 (2007).
Makita, R. et al. Multiple renal cysts, urinary concentration defects, and pulmonary emphysematous changes in mice lacking TAZ. Am. J. Physiol. Ren. Physiol. 294, F542–F553 (2008).
Tian, Y. et al. TAZ promotes PC2 degradation through a SCFbeta-Trcp E3 ligase complex. Mol. Cell Biol. 27, 6383–6395 (2007).
Dingare, C. et al. The Hippo pathway effector Taz is required for cell morphogenesis and fertilization in zebrafish. Development 145, dev167023 (2018).
Jaslove, J. M. et al. Transmural pressure signals through retinoic acid to regulate lung branching. Development 149, dev199726 (2022).
Lin, C. et al. YAP is essential for mechanical force production and epithelial cell proliferation during lung branching morphogenesis. eLife 6, e21130 (2017).
Han, W. M. et al. Synthetic matrix enhances transplanted satellite cell engraftment in dystrophic and aged skeletal muscle with comorbid trauma. Sci. Adv. 4, eaar4008 (2018).
Moyle, L. A. et al. Three-dimensional niche stiffness synergizes with Wnt7a to modulate the extent of satellite cell symmetric self-renewal divisions. Mol. Biol. Cell 31, 1703–1713 (2020).
Gjorevski, N. et al. Designer matrices for intestinal stem cell and organoid culture. Nature 539, 560–564 (2016).
Yui, S. et al. YAP/TAZ-dependent reprogramming of colonic epithelium links ECM remodeling to tissue regeneration. Cell Stem Cell 22, 35–49 e37 (2018).
Bertolio, R. et al. Sterol regulatory element binding protein 1 couples mechanical cues and lipid metabolism. Nat. Commun. 10, 1326 (2019).
Hong, J. H. et al. TAZ, a transcriptional modulator of mesenchymal stem cell differentiation. Science 309, 1074–1078 (2005).
McBeath, R., Pirone, D. M., Nelson, C. M., Bhadriraju, K. & Chen, C. S. Cell shape, cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev. Cell 6, 483–495 (2004). This is one of the founding studies for the field of mechanobiology, showing mechanical control of stem cell commitment.
Sordella, R., Jiang, W., Chen, G. C., Curto, M. & Settleman, J. Modulation of Rho GTPase signaling regulates a switch between adipogenesis and myogenesis. Cell 113, 147–158 (2003).
Tang, Y. et al. MT1-MMP-dependent control of skeletal stem cell commitment via a beta1-integrin/YAP/TAZ signaling axis. Dev. Cell 25, 402–416 (2013).
Chung, H. et al. Yap1 is dispensable for self-renewal but required for proper differentiation of mouse embryonic stem (ES) cells. EMBO Rep. 17, 519–52 (2016).
Lian, I. et al. The role of YAP transcription coactivator in regulating stem cell self-renewal and differentiation. Genes Dev. 24, 1106–1118 (2010).
Tamm, C., Bower, N. & Anneren, C. Regulation of mouse embryonic stem cell self-renewal by a Yes-YAP-TEAD2 signaling pathway downstream of LIF. J. Cell Sci. 124, 1136–1144 (2011).
Qin, H. et al. Transcriptional analysis of pluripotency reveals the Hippo pathway as a barrier to reprogramming. Hum. Mol. Genet. 21, 2054–2067 (2012).
Qin, H. et al. YAP induces human naive pluripotency. Cell Rep. 14, 2301–2312 (2016).
Cebola, I. et al. TEAD and YAP regulate the enhancer network of human embryonic pancreatic progenitors. Nat. Cell Biol. 17, 615–626 (2015).
Hogrebe, N. J., Augsornworawat, P., Maxwell, K. G., Velazco-Cruz, L. & Millman, J. R. Targeting the cytoskeleton to direct pancreatic differentiation of human pluripotent stem cells. Nat. Biotechnol. 38, 460–470 (2020).
Mamidi, A. et al. Mechanosignalling via integrins directs fate decisions of pancreatic progenitors. Nature 564, 114–118 (2018).
Rosado-Olivieri, E. A., Anderson, K., Kenty, J. H. & Melton, D. A. YAP inhibition enhances the differentiation of functional stem cell-derived insulin-producing beta cells. Nat. Commun. 10, 1464 (2019).
Fan, F. et al. Pharmacological targeting of kinases MST1 and MST2 augments tissue repair and regeneration. Sci. Transl. Med. 8, 352ra108 (2016).
Kastan, N. et al. Small-molecule inhibition of Lats kinases may promote Yap-dependent proliferation in postmitotic mammalian tissues. Nat. Commun. 12, 3100 (2021).
Pobbati, A. V. et al. Identification of quinolinols as activators of TEAD-dependent transcription. ACS Chem. Biol. 14, 2909–2921 (2019).
De Rosa, L. et al. Laminin 332-dependent YAP dysregulation depletes epidermal stem cells in junctional epidermolysis bullosa. Cell Rep. 27, 2036–2049 e2036 (2019).
Morikawa, Y., Heallen, T., Leach, J., Xiao, Y. & Martin, J. F. Dystrophin-glycoprotein complex sequesters Yap to inhibit cardiomyocyte proliferation. Nature 547, 227–231 (2017).
Morikawa, Y. et al. Actin cytoskeletal remodeling with protrusion formation is essential for heart regeneration in Hippo-deficient mice. Sci. Signal. 8, ra41 (2015).
Tzima, E. et al. A mechanosensory complex that mediates the endothelial cell response to fluid shear stress. Nature 437, 426–431 (2005).
Shay-Salit, A. et al. VEGF receptor 2 and the adherens junction as a mechanical transducer in vascular endothelial cells. Proc. Natl Acad. Sci. USA 99, 9462–9467 (2002).
Min, E. & Schwartz, M. A. Translocating transcription factors in fluid shear stress-mediated vascular remodeling and disease. Exp. Cell Res. 376, 92–97 (2019).
Chiu, J. J. & Chien, S. Effects of disturbed flow on vascular endothelium: pathophysiological basis and clinical perspectives. Physiol. Rev. 91, 327–387 (2011).
Rojo de la Vega, M., Chapman, E. & Zhang, D. D. NRF2 and the hallmarks of cancer. Cancer Cell 34, 21–43 (2018).
Tonelli, C., Chio, I. I. C. & Tuveson, D. A. Transcriptional regulation by Nrf2. Antioxid. Redox Signal. 29, 1727–1745 (2018).
Chen, X. L. et al. Laminar flow induction of antioxidant response element-mediated genes in endothelial cells. A novel anti-inflammatory mechanism. J. Biol. Chem. 278, 703–711 (2003).
Dai, G. et al. Biomechanical forces in atherosclerosis-resistant vascular regions regulate endothelial redox balance via phosphoinositol 3-kinase/Akt-dependent activation of Nrf2. Circ. Res. 101, 723–733 (2007).
Hosoya, T. et al. Differential responses of the Nrf2-Keap1 system to laminar and oscillatory shear stresses in endothelial cells. J. Biol. Chem. 280, 27244–27250 (2005).
Hsieh, C. Y. et al. Regulation of shear-induced nuclear translocation of the Nrf2 transcription factor in endothelial cells. J. Biomed. Sci. 16, 12 (2009).
Jones, C. I. III et al. Regulation of antioxidants and phase 2 enzymes by shear-induced reactive oxygen species in endothelial cells. Ann. Biomed. Eng. 35, 683–693 (2007).
Psefteli, P. M. et al. Glycocalyx sialic acids regulate Nrf2-mediated signaling by fluid shear stress in human endothelial cells. Redox Biol. 38, 101816 (2021).
Warabi, E. et al. Shear stress stabilizes NF-E2-related factor 2 and induces antioxidant genes in endothelial cells: role of reactive oxygen/nitrogen species. Free. Radic. Biol. Med. 42, 260–269 (2007).
Baeyens, N. et al. Defective fluid shear stress mechanotransduction mediates hereditary hemorrhagic telangiectasia. J. Cell Biol. 214, 807–816 (2016).
Poduri, A. et al. Endothelial cells respond to the direction of mechanical stimuli through SMAD signaling to regulate coronary artery size. Development 144, 3241–3252 (2017).
Zhou, J. et al. Force-specific activation of Smad1/5 regulates vascular endothelial cell cycle progression in response to disturbed flow. Proc. Natl Acad. Sci. USA 109, 7770–7775 (2012).
Goetz, J. G. et al. Endothelial cilia mediate low flow sensing during zebrafish vascular development. Cell Rep. 6, 799–808 (2014).
Vion, A. C. et al. Primary cilia sensitize endothelial cells to BMP and prevent excessive vascular regression. J. Cell Biol. 217, 1651–1665 (2018).
Huang, J. et al. KLF2 mediates the suppressive effect of laminar flow on vascular calcification by inhibiting endothelial BMP/SMAD1/5 signaling. Circ. Res. 129, e87–e100 (2021).
Ahimou, F., Mok, L. P., Bardot, B. & Wesley, C. The adhesion force of Notch with Delta and the rate of Notch signaling. J. Cell Biol. 167, 1217–1229 (2004).
Gordon, W. R. et al. Mechanical allostery: evidence for a force requirement in the proteolytic activation of Notch. Dev. Cell 33, 729–736 (2015).
Langridge, P. D. & Struhl, G Epsin-dependent ligand endocytosis activates Notch by force. Cell 171, 1383–1396.e1312 (2017).
Meloty-Kapella, L., Shergill, B., Kuon, J., Botvinick, E. & Weinmaster, G. Notch ligand endocytosis generates mechanical pulling force dependent on dynamin, epsins, and actin. Dev. Cell 22, 1299–1312 (2012).
Sprinzak, D. & Blacklow, S. C. Biophysics of Notch signaling. Annu. Rev. Biophys. 50, 157–189 (2021).
Wang, X. & Ha, T. Defining single molecular forces required to activate integrin and Notch signaling. Science 340, 991–994 (2013).
Caolo, V. et al. Shear stress activates ADAM10 sheddase to regulate Notch1 via the Piezo1 force sensor in endothelial cells. eLife 9, e50684 (2020).
Fang, J. S. et al. Shear-induced Notch-Cx37-p27 axis arrests endothelial cell cycle to enable arterial specification. Nat. Commun. 8, 2149 (2017).
Lee, J. et al. 4-Dimensional light-sheet microscopy to elucidate shear stress modulation of cardiac trabeculation. J. Clin. Invest. 126, 1679–1690 (2016).
Mack, J. J. et al. NOTCH1 is a mechanosensor in adult arteries. Nat. Commun. 8, 1620 (2017).
Masumura, T., Yamamoto, K., Shimizu, N., Obi, S. & Ando, J. Shear stress increases expression of the arterial endothelial marker ephrinB2 in murine ES cells via the VEGF-Notch signaling pathways. Arterioscler. Thromb. Vasc. Biol. 29, 2125–2131 (2009).
Polacheck, W. J. et al. A non-canonical Notch complex regulates adherens junctions and vascular barrier function. Nature 552, 258–262 (2017).
van Engeland, N. C. A. et al. Vimentin regulates Notch signaling strength and arterial remodeling in response to hemodynamic stress. Sci. Rep. 9, 12415 (2019).
Chu, Y. S. et al. Force measurements in E-cadherin-mediated cell doublets reveal rapid adhesion strengthened by actin cytoskeleton remodeling through Rac and Cdc42. J. Cell Biol. 167, 1183–1194 (2004).
Hunter, G. L. et al. A role for actomyosin contractility in Notch signaling. BMC Biol. 17, 12 (2019).
Liu, Z. et al. Mechanical tugging force regulates the size of cell-cell junctions. Proc. Natl Acad. Sci. USA 107, 9944–9949 (2010).
Shaya, O. et al. Cell-cell contact area affects notch signaling and notch-dependent patterning. Dev. Cell 40, 505–511 e506 (2017).
Chen, C. S., Mrksich, M., Huang, S., Whitesides, G. M. & Ingber, D. E. Geometric control of cell life and death. Science 276, 1425–1428 (1997). This study demonstrates that spatial arrangement of the ECM, and not the amount of ECM ligand, can drive a complex cellular phenotype.
Folkman, J. & Moscona, A. Role of cell shape in growth control. Nature 273, 345–349 (1978).
Moro, A. et al. MicroRNA-dependent regulation of biomechanical genes establishes tissue stiffness homeostasis. Nat. Cell Biol. 21, 348–358 (2019).
Astone, M. et al. Zebrafish mutants and TEAD reporters reveal essential functions for Yap and Taz in posterior cardinal vein development. Sci. Rep. 8, 10189 (2018).
Kim, J. et al. YAP/TAZ regulates sprouting angiogenesis and vascular barrier maturation. J. Clin. Invest. 127, 3441–3461 (2017).
Neto, F. et al. YAP and TAZ regulate adherens junction dynamics and endothelial cell distribution during vascular development. eLife 7, e31037 (2018).
van der Stoel, M. et al. DLC1 is a direct target of activated YAP/TAZ that drives collective migration and sprouting angiogenesis. J. Cell Sci. 133, jcs239947 (2020).
Wang, X. et al. YAP/TAZ Orchestrate VEGF signaling during developmental angiogenesis. Dev. Cell 42, 462–478 e467 (2017).
Mammoto, A. et al. A mechanosensitive transcriptional mechanism that controls angiogenesis. Nature 457, 1103–1108 (2009).
Deng, H. et al. Activation of Smad2/3 signaling by low fluid shear stress mediates artery inward remodeling. Proc. Natl Acad. Sci. USA 118, e2105339118 (2021).
Hahn, C. & Schwartz, M. A. Mechanotransduction in vascular physiology and atherogenesis. Nat. Rev. Mol. Cell Biol. 10, 53–62 (2009).
Nakajima, H. & Mochizuki, N. Flow pattern-dependent endothelial cell responses through transcriptional regulation. Cell Cycle 16, 1893–1901 (2017).
Ishida, T., Takahashi, M., Corson, M. A. & Berk, B. C. Fluid shear stress-mediated signal transduction: how do endothelial cells transduce mechanical force into biological responses? Ann. N. Y. Acad. Sci. 811, 12–23 (1997).
Khachigian, L. M., Resnick, N., Gimbrone, M. A. Jr. & Collins, T. Nuclear factor-kappa B interacts functionally with the platelet-derived growth factor B-chain shear-stress response element in vascular endothelial cells exposed to fluid shear stress. J. Clin. Invest. 96, 1169–1175 (1995).
Lan, Q., Mercurius, K. O. & Davies, P. F. Stimulation of transcription factors NF kappa B and AP1 in endothelial cells subjected to shear stress. Biochem. Biophys. Res. Commun. 201, 950–956 (1994). This study is one of the earliest demonstrations of a mechanoresponsive TF.
Mohan, S. et al. Low shear stress preferentially enhances IKK activity through selective sources of ROS for persistent activation of NF-kappaB in endothelial cells. Am. J. Physiol. Cell Physiol. 292, C362–C371 (2007).
Nagel, T., Resnick, N., Dewey, C. F. Jr. & Gimbrone, M. A. Jr. Vascular endothelial cells respond to spatial gradients in fluid shear stress by enhanced activation of transcription factors. Arterioscler. Thromb. Vasc. Biol. 19, 1825–1834 (1999).
Tzima, E. et al. Activation of Rac1 by shear stress in endothelial cells mediates both cytoskeletal reorganization and effects on gene expression. EMBO J. 21, 6791–6800 (2002).
Liu, D. et al. Atheroprotective effects of methotrexate via the inhibition of YAP/TAZ under disturbed flow. J. Transl. Med. 17, 378 (2019).
Park, H. et al. Defective flow-migration coupling causes arteriovenous malformations in hereditary hemorrhagic telangiectasia. Circulation 144, 805–822 (2021).
Orr, A. W. et al. The subendothelial extracellular matrix modulates NF-kappaB activation by flow: a potential role in atherosclerosis. J. Cell Biol. 169, 191–202 (2005).
Drain, A. P. et al. Matrix compliance permits NF-κB activation to drive therapy resistance in breast cancer. J. Exp. Med. 218, e20191360 (2021).
Ishihara, S. et al. Substrate stiffness regulates temporary NF-kappaB activation via actomyosin contractions. Exp. Cell Res. 319, 2916–2927 (2013).
Liang, F. & Gardner, D. G. Mechanical strain activates BNP gene transcription through a p38/NF-kappaB-dependent mechanism. J. Clin. Invest. 104, 1603–1612 (1999).
Yamamoto, K. et al. Induction of tenascin-C in cardiac myocytes by mechanical deformation. Role of reactive oxygen species. J. Biol. Chem. 274, 21840–21846 (1999).
Mochitate, K., Pawelek, P. & Grinnell, F. Stress relaxation of contracted collagen gels: disruption of actin filament bundles, release of cell surface fibronectin, and down-regulation of DNA and protein synthesis. Exp. Cell Res. 193, 198–207 (1991).
Aramaki-Hattori, N., Okabe, K., Hamada, M., Takato, T. & Kishi, K. Relationship between keloid formation and YAP/TAZ signaling. Plast. Reconstr. Surg. Glob. Open. 5, e1357 (2017).
Huang, C. et al. Keloid progression: a stiffness gap hypothesis. Int. Wound J. 14, 764–771 (2017).
Chen, K. et al. Disrupting biological sensors of force promotes tissue regeneration in large organisms. Nat. Commun. 12, 5256 (2021).
Mascharak, S. et al. Preventing Engrailed-1 activation in fibroblasts yields wound regeneration without scarring. Science https://doi.org/10.1126/science.aba2374 (2021).
Mascharak, S. et al. Multi-omic analysis reveals divergent molecular events in scarring and regenerative wound healing. Cell Stem Cell 29, 315–327 e316 (2022).
Bertero, T. et al. Tumor-stroma mechanics coordinate amino acid availability to sustain tumor growth and malignancy. Cell Metab. 29, 124–140 e110 (2019).
Li, R., Li, X., Hagood, J., Zhu, M. S. & Sun, X. Myofibroblast contraction is essential for generating and regenerating the gas-exchange surface. J. Clin. Invest. 130, 2859–2871 (2020).
Bell, J. L. et al. Optimization of novel nipecotic bis(amide) inhibitors of the Rho/MKL1/SRF transcriptional pathway as potential anti-metastasis agents. Bioorg. Med. Chem. Lett. 23, 3826–3832 (2013).
Fan, L. et al. Cell contact-dependent regulation of epithelial-myofibroblast transition via the rho-rho kinase-phospho-myosin pathway. Mol. Biol. Cell 18, 1083–1097 (2007).
Haak, A. J. et al. Targeting the myofibroblast genetic switch: inhibitors of myocardin-related transcription factor/serum response factor-regulated gene transcription prevent fibrosis in a murine model of skin injury. J. Pharmacol. Exp. Ther. 349, 480–486 (2014).
Hutchings, K. M. et al. Pharmacokinetic optimitzation of CCG-203971: novel inhibitors of the Rho/MRTF/SRF transcriptional pathway as potential antifibrotic therapeutics for systemic scleroderma. Bioorg. Med. Chem. Lett. 27, 1744–1749 (2017).
Johnson, L. A. et al. Novel Rho/MRTF/SRF inhibitors block matrix-stiffness and TGF-beta-induced fibrogenesis in human colonic myofibroblasts. Inflamm. Bowel Dis. 20, 154–165 (2014).
Luchsinger, L. L., Patenaude, C. A., Smith, B. D. & Layne, M. D. Myocardin-related transcription factor-A complexes activate type I collagen expression in lung fibroblasts. J. Biol. Chem. 286, 44116–44125 (2011).
Sandbo, N., Kregel, S., Taurin, S., Bhorade, S. & Dulin, N. O. Critical role of serum response factor in pulmonary myofibroblast differentiation induced by TGF-beta. Am. J. Respir. Cell Mol. Biol. 41, 332–338 (2009).
Shi, Z., Ren, M. & Rockey, D. C. Myocardin and myocardin-related transcription factor-A synergistically mediate actin cytoskeletal-dependent inhibition of liver fibrogenesis. Am. J. Physiol. Gastrointest. Liver Physiol. 318, G504–G517 (2020).
Shiwen, X. et al. A role of myocardin related transcription factor-A (MRTF-A) in scleroderma related fibrosis. PLoS ONE 10, e0126015 (2015).
Sisson, T. H. et al. Inhibition of myocardin-related transcription factor/serum response factor signaling decreases lung fibrosis and promotes mesenchymal cell apoptosis. Am. J. Pathol. 185, 969–986 (2015).
Small, E. M. et al. Myocardin-related transcription factor-a controls myofibroblast activation and fibrosis in response to myocardial infarction. Circ. Res. 107, 294–304 (2010).
Tian, W. et al. Myocardin related transcription factor A programs epigenetic activation of hepatic stellate cells. J. Hepatol. 62, 165–174 (2015).
Velasquez, L. S. et al. Activation of MRTF-A-dependent gene expression with a small molecule promotes myofibroblast differentiation and wound healing. Proc. Natl Acad. Sci. USA 110, 16850–16855 (2013).
Xu, H. et al. Myocardin-related transcription factor a epigenetically regulates renal fibrosis in diabetic nephropathy. J. Am. Soc. Nephrol. 26, 1648–1660 (2015).
Francisco, J. et al. Blockade of fibroblast YAP attenuates cardiac fibrosis and dysfunction through MRTF-A inhibition. JACC Basic. Transl. Sci. 5, 931–945 (2020).
Er, E. E. et al. Pericyte-like spreading by disseminated cancer cells activates YAP and MRTF for metastatic colonization. Nat. Cell Biol. 20, 966–978 (2018).
Girard, C. A. et al. A feed-forward mechanosignaling loop confers resistance to therapies targeting the MAPK pathway in BRAF-mutant melanoma. Cancer Res. 80, 1927–1941 (2020).
Kim, T. et al. MRTF potentiates TEAD-YAP transcriptional activity causing metastasis. EMBO J. 36, 520–535 (2017).
Kim, T. et al. A basal-like breast cancer-specific role for SRF-IL6 in YAP-induced cancer stemness. Nat. Commun. 6, 10186 (2015).
Yu, O. M. et al. YAP and MRTF-A, transcriptional co-activators of RhoA-mediated gene expression, are critical for glioblastoma tumorigenicity. Oncogene 37, 5492–5507 (2018).
Budi, E. H., Schaub, J. R., Decaris, M., Turner, S. & Derynck, R. TGF-beta as a driver of fibrosis: physiological roles and therapeutic opportunities. J. Pathol. 254, 358–373 (2021).
Ahamed, J., Janczak, C. A., Wittkowski, K. M. & Coller, B. S. In vitro and in vivo evidence that thrombospondin-1 (TSP-1) contributes to stirring- and shear-dependent activation of platelet-derived TGF-beta1. PLoS ONE 4, e6608 (2009).
Buscemi, L. et al. The single-molecule mechanics of the latent TGF-beta1 complex. Curr. Biol. 21, 2046–2054 (2011).
Froese, A. R. et al. Stretch-induced activation of transforming growth factor-beta1 in pulmonary fibrosis. Am. J. Respir. Crit. Care Med. 194, 84–96 (2016).
Klingberg, F. et al. Prestress in the extracellular matrix sensitizes latent TGF-beta1 for activation. J. Cell Biol. 207, 283–297 (2014).
Munger, J. S., Harpel, J. G., Giancotti, F. G. & Rifkin, D. B. Interactions between growth factors and integrins: latent forms of transforming growth factor-beta are ligands for the integrin alphavbeta1. Mol. Biol. Cell 9, 2627–2638 (1998). This paper describes the discovery that paved the way to the idea of force-mediated liberation of TGFβ1 ligands from the ECM.
Shi, M. et al. Latent TGF-beta structure and activation. Nature 474, 343–349 (2011).
Wipff, P. J., Rifkin, D. B., Meister, J. J. & Hinz, B. Myofibroblast contraction activates latent TGF-beta1 from the extracellular matrix. J. Cell Biol. 179, 1311–1323 (2007).
Zhou, Y., Hagood, J. S., Lu, B., Merryman, W. D. & Murphy-Ullrich, J. E. Thy-1-integrin alphav beta5 interactions inhibit lung fibroblast contraction-induced latent transforming growth factor-beta1 activation and myofibroblast differentiation. J. Biol. Chem. 285, 22382–22393 (2010).
Jones, D. L. et al. ZNF416 is a pivotal transcriptional regulator of fibroblast mechanoactivation. J. Cell Biol. https://doi.org/10.1083/jcb.202007152 (2021).
Zhang, K. et al. Mechanical signals regulate and activate SNAIL1 protein to control the fibrogenic response of cancer-associated fibroblasts. J. Cell Sci. 129, 1989–2002 (2016).
Butcher, D. T., Alliston, T. & Weaver, V. M. A tense situation: forcing tumour progression. Nat. Rev. Cancer 9, 108–122 (2009).
Gehler, S., Ponik, S. M., Riching, K. M. & Keely, P. J. Bi-directional signaling: extracellular matrix and integrin regulation of breast tumor progression. Crit. Rev. Eukaryot. Gene Expr. 23, 139–157 (2013).
Mohammadi, H. & Sahai, E. Mechanisms and impact of altered tumour mechanics. Nat. Cell Biol. 20, 766–774 (2018).
Chen, D. et al. LIFR is a breast cancer metastasis suppressor upstream of the Hippo-YAP pathway and a prognostic marker. Nat. Med. 18, 1511–1517 (2012).
Cordenonsi, M. et al. The Hippo transducer TAZ confers cancer stem cell-related traits on breast cancer cells. Cell 147, 759–772 (2011).
Hiemer, S. E., Szymaniak, A. D. & Varelas, X. The transcriptional regulators TAZ and YAP direct transforming growth factor beta-induced tumorigenic phenotypes in breast cancer cells. J. Biol. Chem. 289, 13461–13474 (2014).
Lamar, J. M. et al. SRC tyrosine kinase activates the YAP/TAZ axis and thereby drives tumor growth and metastasis. J. Biol. Chem. 294, 2302–2317 (2019).
Overholtzer, M. et al. Transforming properties of YAP, a candidate oncogene on the chromosome 11q22 amplicon. Proc. Natl Acad. Sci. USA 103, 12405–12410 (2006).
Rashidian, J. et al. Ski regulates Hippo and TAZ signaling to suppress breast cancer progression. Sci. Signal. 8, ra14 (2015).
Yang, C. S. et al. Glutamine-utilizing transaminases are a metabolic vulnerability of TAZ/YAP-activated cancer cells. EMBO Rep. 19, e43577 (2018).
Zanconato, F. et al. Transcriptional addiction in cancer cells is mediated by YAP/TAZ through BRD4. Nat. Med. 24, 1599–1610 (2018).
Zanconato, F. et al. Genome-wide association between YAP/TAZ/TEAD and AP-1 at enhancers drives oncogenic growth. Nat. Cell Biol. 17, 1218–1227 (2015).
Lin, C. H. et al. Microenvironment rigidity modulates responses to the HER2 receptor tyrosine kinase inhibitor lapatinib via YAP and TAZ transcription factors. Mol. Biol. Cell 26, 3946–3953 (2015).
Panciera, T. et al. Reprogramming normal cells into tumour precursors requires ECM stiffness and oncogene-mediated changes of cell mechanical properties. Nat. Mater. 19, 797–806 (2020).
Ding, X., Park, S. I., McCauley, L. K. & Wang, C. Y. Signaling between transforming growth factor beta (TGF-beta) and transcription factor SNAI2 represses expression of microRNA miR-203 to promote epithelial-mesenchymal transition and tumor metastasis. J. Biol. Chem. 288, 10241–10253 (2013).
Moes, M. et al. A novel network integrating a miRNA-203/SNAI1 feedback loop which regulates epithelial to mesenchymal transition. PLoS ONE 7, e35440 (2012).
Northey, J. J. et al. Stiff stroma increases breast cancer risk by inducing the oncogene ZNF217. J. Clin. Invest. 130, 5721–5737 (2020).
Wei, S. C. et al. Matrix stiffness drives epithelial-mesenchymal transition and tumour metastasis through a TWIST1-G3BP2 mechanotransduction pathway. Nat. Cell Biol. 17, 678–688 (2015).
Barry-Hamilton, V. et al. Allosteric inhibition of lysyl oxidase-like-2 impedes the development of a pathologic microenvironment. Nat. Med. 16, 1009–1017 (2010).
Bui, T. et al. Functional redundancy between beta1 and beta3 Integrin in activating the IR/Akt/mTORC1 signaling axis to promote ErbB2-driven breast cancer. Cell Rep. 29, 589–602.e586 (2019).
Chan, S. W. et al. A role for TAZ in migration, invasion, and tumorigenesis of breast cancer cells. Cancer Res. 68, 2592–2598 (2008).
Chen, Q. et al. A temporal requirement for Hippo signaling in mammary gland differentiation, growth, and tumorigenesis. Genes Dev. 28, 432–437 (2014).
Levental, K. R. et al. Matrix crosslinking forces tumor progression by enhancing integrin signaling. Cell 139, 891–906 (2009). This pioneering study shows the relevance of matrix crosslinking and ECM mechanics for breast cancer tumorigenesis in vivo.
Mooney, D. et al. Switching from differentiation to growth in hepatocytes: control by extracellular matrix. J. Cell Physiol. 151, 497–505 (1992).
Driskill, J. H. & Pan, D. The hippo pathway in liver homeostasis and pathophysiology. Annu. Rev. Pathol. 16, 299–322 (2021).
Ishikawa, J. et al. Mechanical homeostasis of liver sinusoid is involved in the initiation and termination of liver regeneration. Commun. Biol. 4, 409 (2021).
Ye, J. Transcription factors activated through RIP (regulated intramembrane proteolysis) and RAT (regulated alternative translocation). J. Biol. Chem. 295, 10271–10280 (2020).
Basu, H. et al. FHL2 anchors mitochondria to actin and adapts mitochondrial dynamics to glucose supply. J. Cell Biol. 220, e201912077 (2021).
De Vos, K. J., Allan, V. J., Grierson, A. J. & Sheetz, M. P. Mitochondrial function and actin regulate dynamin-related protein 1-dependent mitochondrial fission. Curr. Biol. 15, 678–683 (2005).
Korobova, F., Ramabhadran, V. & Higgs, H. N. An actin-dependent step in mitochondrial fission mediated by the ER-associated formin INF2. Science 339, 464–467 (2013).
Majstrowicz, K. et al. Coordination of mitochondrial and cellular dynamics by the actin-based motor Myo19. J. Cell Sci. 134, jcs255844 (2021).
Manor, U. et al. A mitochondria-anchored isoform of the actin-nucleating spire protein regulates mitochondrial division. eLife 4, e08828 (2015).
Moore, A. S. et al. Actin cables and comet tails organize mitochondrial networks in mitosis. Nature 591, 659–664 (2021).
Moore, A. S., Wong, Y. C., Simpson, C. L. & Holzbaur, E. L. Dynamic actin cycling through mitochondrial subpopulations locally regulates the fission-fusion balance within mitochondrial networks. Nat. Commun. 7, 12886 (2016).
Yang, H. et al. Materials stiffness-dependent redox metabolic reprogramming of mesenchymal stem cells for secretome-based therapeutic angiogenesis. Adv. Healthc. Mater. 8, e1900929 (2019).
Khacho, M. et al. Mitochondrial dynamics impacts stem cell identity and fate decisions by regulating a nuclear transcriptional program. Cell Stem Cell 19, 232–247 (2016).
Khacho, M. et al. Mitochondrial dysfunction underlies cognitive defects as a result of neural stem cell depletion and impaired neurogenesis. Hum. Mol. Genet. 26, 3327–3341 (2017).
The authors are indebted to Y. Miroshnikova for reading the manuscript and providing thoughtful advice. They apologize to colleagues whose work they have inadvertently failed to cite. Work on nuclear mechanotransduction in the Dupont lab is supported by Worldwide Cancer Research grant no. 21-0156 and AIRC Foundation Investigator grant no. 21392, and in the Wickström lab by the Academy of Finland and Max Planck Society.
The authors declare no competing interests.
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- Mechanical force
A force that is caused by contact with another object and that induces a change in state of rest or motion.
Occurs when a fluid applies a tangential force that pushes one part of the cell in one direction, and the rest of the cell is dragged in the opposite direction by adhesion to the extracellular matrix or to other cells.
- Tensile stress
The action–reaction forces acting at the cell–extracellular matrix or cell–cell contact sites to stretch cells and to resist deformation. Tensile stress is the opposite of compression.
- Hydrostatic pressure
A pressure exerted by a fluid onto a contact surface as a result of gravity.
The ability of a material to return to its original shape after a deformation-inducing force has been removed.
The ability of a fluid to resist gradual deformation. Most biological materials are considered viscoelastic, that is, they display properties of both elastic and viscous materials. Viscoelastic materials have a strain (deformation) rate that is dependent on time and they dissipate energy when a load is applied and removed whereas a purely elastic material does not.
A resistance that a surface encounters when it is moving over another surface, for example, in joints, tendons, eye or skin.
- LINC complex
A multi-protein complex that crosses the nuclear envelope and provides a physical link between cytoplasmic and nuclear cytoskeletal structures.
- Nuclear lamina
Structure between the inner nuclear membrane and peripheral chromatin, composed mainly of intermediate filament proteins, the lamins, and lamin-associated proteins.
- Epidermal differentiation complex
(EDC). A gene complex of >50 genes that encode proteins involved in terminal differentiation and cornification of skin epidermal keratinocytes.
The process by which a cell subjected to mechanical forces reinforces its force-bearing structures.
- Mediator complex
A multisubunit protein complex that bridges transcription factors and the basal RNA polymerase II transcriptional machinery.
- Contact inhibition of growth
The process by which cell crowding inhibits cell proliferation through the establishment of cell–cell contacts and the reduction of cell size.
- Cell competition
The active elimination of a viable but undesirable cell population by competitive interactions within a tissue.
The tissue of the pre-implantation mammalian embryo that will contribute to formation of the placenta.
- Inner cell mass
The tissue of the pre-implantation mammalian embryo that will contribute to formation of the tissues of the fetus.
The somatic cells of the female gonad that surround oocytes and support their growth.
- Branching morphogenesis
The developmental process by which a growing epithelium buds branches in the surrounding mesenchyme to form a tree-like structure.
- Naive pluripotent state
Pluripotent stem cells equivalent to those found in the early inner cell mass, with largely unmethylated genome, and still able to differentiate into germ cells.
- Regenerative medicine
The clinical use of stem cells to stimulate repair mechanisms and restore function in damaged body tissues or organs.
- Laminar flow
When particles in a moving liquid follow linear paths without lateral mixing.
- Sprouting angiogenesis
The growth of new capillary vessels out of pre-existing blood vessels.
- Fibrotic response
Tissue remodelling characterized by the deposition of collagenous extracellular matrix, which can have a physiological function during wound healing (scarring) or a pathological function that can interfere with or totally inhibit the normal architecture and function of the underlying organ or tissue.
- Cancer-associated fibroblasts
A population of cells, likely deriving from the fibroblast lineages, that are found in tumours and have an elongated morphology, are negative for epithelial, endothelial and leukocyte markers, and lack the mutations found in cancer cells.
- Epithelial to mesenchymal transition
(EMT). The differentiation process by which cells lose epithelial identity and the ability to form stable cell–cell adhesions, and gain expression of mesenchymal markers associated with increased migratory ability.
- Cholangiocellular transdifferentiation
The differentiation of hepatocytes into cells that express markers typically found in bile duct cells and in bipotent liver cell progenitors.
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Dupont, S., Wickström, S.A. Mechanical regulation of chromatin and transcription. Nat Rev Genet (2022). https://doi.org/10.1038/s41576-022-00493-6