Review Article | Published:

Emerging views of the nucleus as a cellular mechanosensor

Nature Cell Biologyvolume 20pages373381 (2018) | Download Citation

Abstract

The ability of cells to respond to mechanical forces is critical for numerous biological processes. Emerging evidence indicates that external mechanical forces trigger changes in nuclear envelope structure and composition, chromatin organization and gene expression. However, it remains unclear if these processes originate in the nucleus or are downstream of cytoplasmic signals. Here we discuss recent findings that support a direct role of the nucleus in cellular mechanosensing and highlight novel tools to study nuclear mechanotransduction.

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References

  1. 1.

    Dewey, C. F. Jr, Bussolari, S. R., Gimbrone, M. A. Jr & Davies, P. F. The dynamic response of vascular endothelial cells to fluid shear stress. J. Biomech. Eng. 103, 177–185 (1981).

  2. 2.

    Freeman, P. M., Natarajan, R. N., Kimura, J. H. & Andriacchi, T. P. Chondrocyte cells respond mechanically to compressive loads. J. Orthop. Res. 12, 311–320 (1994).

  3. 3.

    Magid, A. & Law, D. J. Myofibrils bear most of the resting tension in frog skeletal muscle. Science 230, 1280–1282 (1985).

  4. 4.

    Le, H. Q. et al. Mechanical regulation of transcription controls Polycomb-mediated gene silencing during lineage commitment. Nat. Cell Biol. 18, 864–875 (2016).

  5. 5.

    Lecuit, T. & Lenne, P. F. Cell surface mechanics and the control of cell shape, tissue patterns and morphogenesis. Nat. Rev. Mol. Cell Biol. 8, 633–644 (2007).

  6. 6.

    Barnes, J. M., Przybyla, L. & Weaver, V. M. Tissue mechanics regulate brain development, homeostasis and disease. J. Cell Sci. 130, 71–82 (2017).

  7. 7.

    Matthews, B. D., Overby, D. R., Mannix, R. & Ingber, D. E. Cellular adaptation to mechanical stress: role of integrins, Rho, cytoskeletal tension and mechanosensitive ion channels. J. Cell Sci. 119, 508–518 (2006).

  8. 8.

    Reidy, M. A. & Bowyer, D. E. Scanning electron microscopy of arteries. The morphology of aortic endothelium in haemodynamically stressed areas associated with branches. Atherosclerosis 26, 181–194 (1977).

  9. 9.

    Tucker, J. B. & Meats, M. Microtubules and control of insect egg shape. J. Cell Biol. 71, 207–217 (1976).

  10. 10.

    Ingber, D. E. Mechanobiology and diseases of mechanotransduction. Ann. Med. 35, 564–577 (2003).

  11. 11.

    Jaalouk, D. E. & Lammerding, J. Mechanotransduction gone awry. Nat. Rev. Mol. Cell Biol. 10, 63–73 (2009).

  12. 12.

    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).

  13. 13.

    Humphrey, J. D., Dufresne, E. R. & Schwartz, M. A. Mechanotransduction and extracellular matrix homeostasis. Nat. Rev. Mol. Cell Biol. 15, 802–812 (2014).

  14. 14.

    Muehlich, S., Hermanns, C., Meier, M. A., Kircher, P. & Gudermann, T. Unravelling a new mechanism linking actin polymerization and gene transcription. Nucleus 7, 121–125 (2016).

  15. 15.

    Panciera, T., Azzolin, L., Cordenonsi, M. & Piccolo, S. Mechanobiology of YAP and TAZ in physiology and disease. Nat. Rev. Mol. Cell Biol. 18, 758–770 (2017).

  16. 16.

    Ross, T. D. et al. Integrins in mechanotransduction. Curr. Opin. Cell Biol. 25, 613–618 (2013).

  17. 17.

    Sun, Z., Guo, S. S. & Fassler, R. Integrin-mediated mechanotransduction. J. Cell Biol. 215, 445–456 (2016).

  18. 18.

    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).

  19. 19.

    Fedorchak, G. R., Kaminski, A. & Lammerding, J. Cellular mechanosensing: getting to the nucleus of it all. Prog. Biophys. Mol. Biol. 115, 76–92 (2014).

  20. 20.

    Maniotis, A. J., Chen, C. S. & Ingber, D. E. Demonstration of mechanical connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc. Natl Acad. Sci. USA 94, 894–854 (1997).

  21. 21.

    Chambliss, A. B. et al. The LINC-anchored actin cap connects the extracellular milieu to the nucleus for ultrafast mechanotransduction. Sci. Rep. 3, 1087 (2013).

  22. 22.

    Wang, N., Tytell, J. D. & Ingber, D. E. Mechanotransduction at a distance: mechanically coupling the extracellular matrix with the nucleus. Nat. Rev. Mol. Cell Biol. 10, 75–82 (2009).

  23. 23.

    Cho, S., Irianto, J. & Discher, D. E. Mechanosensing by the nucleus: from pathways to scaling relationships. J. Cell Biol. 216, 305–315 (2017).

  24. 24.

    Miroshnikova, Y. A., Nava, M. M. & Wickstrom, S. A. Emerging roles of mechanical forces in chromatin regulation. J. Cell Sci. 130, 2243–2250 (2017).

  25. 25.

    Arsenovic, P. T. et al. Nesprin-2G, a component of the nuclear LINC complex, is subject to myosin-dependent tension. Biophys. J. 110, 34–43 (2016).

  26. 26.

    Guilluy, C. et al. Isolated nuclei adapt to force and reveal a mechanotransduction pathway in the nucleus. Nat. Cell Biol. 16, 376–381 (2014).

  27. 27.

    Lombardi, M. L. et al. The interaction between nesprins and sun proteins at the nuclear envelope is critical for force transmission between the nucleus and cytoskeleton. J. Biol. Chem. 286, 26743–26753 (2011).

  28. 28.

    Tajik, A. et al. Transcription upregulation via force-induced direct stretching of chromatin. Nat. Mater. 15, 1287–1296 (2016).

  29. 29.

    Caille, N., Thoumine, O., Tardy, Y. & Meister, J. J. Contribution of the nucleus to the mechanical properties of endothelial cells. J. Biomech. 35, 177–187 (2002).

  30. 30.

    Lammerding, J. Mechanics of the nucleus. Compr. Physiol. 1, 783–807 (2011).

  31. 31.

    Dreger, M., Bengtsson, L., Schoneberg, T., Otto, H. & Hucho, F. Nuclear envelope proteomics: novel integral membrane proteins of the inner nuclear membrane. Proc. Natl Acad. Sci. USA 98, 11943–11948 (2001).

  32. 32.

    Worman, H. J. & Schirmer, E. C. Nuclear membrane diversity: underlying tissue-specific pathologies in disease? Curr. Opin. Cell Biol. 34, 101–112 (2015).

  33. 33.

    Jovanovic-Talisman, T. & Zilman, A. Protein transport by the nuclear pore complex: simple biophysics of a complex biomachine. Biophys. J. 113, 6–14 (2017).

  34. 34.

    de Leeuw, R., Gruenbaum, Y. & Medalia, O. Nuclear lamins: thin filaments with major functions. Trends Cell Biol. 28, 34–45 (2018).

  35. 35.

    Ho, C. Y. & Lammerding, J. Lamins at a glance. J. Cell Sci. 125, 2087–2093 (2012).

  36. 36.

    Genschel, J. & Schmidt, H. H. Mutations in the LMNA gene encoding lamin A/C. Hum. Mutat. 16, 451–459 (2000).

  37. 37.

    Bonne, G. et al. Mutations in the gene encoding lamin A/C cause autosomal dominant Emery–Dreifuss muscular dystrophy. Nat. Genet. 21, 285–288 (1999).

  38. 38.

    Fatkin, D. et al. Missense mutations in the rod domain of the lamin A/C gene as causes of dilated cardiomyopathy and conduction-system disease. N. Engl. J. Med. 341, 1715–1724 (1999).

  39. 39.

    Lammerding, J. et al. Lamins A and C but not lamin B1 regulate nuclear mechanics. J. Biol. Chem. 281, 25768–25780 (2006).

  40. 40.

    Zwerger, M. et al. Altering lamina assembly reveals lamina-dependent and -independent functions for A-type lamins. J. Cell Sci. 128, 3607–3620 (2015).

  41. 41.

    Broers, J. L. et al. Decreased mechanical stiffness in LMNA −/− cells is caused by defective nucleo-cytoskeletal integrity: implications for the development of laminopathies. Hum. Mol. Genet. 13, 2567–2580 (2004).

  42. 42.

    Dahl, K. N., Kahn, S. M., Wilson, K. L. & Discher, D. E. The nuclear envelope lamina network has elasticity and a compressibility limit suggestive of a molecular shock absorber. J. Cell Sci. 117, 4779–4786 (2004).

  43. 43.

    Lammerding, J. et al. Lamin A/C deficiency causes defective nuclear mechanics and mechanotransduction. J. Clin. Invest. 113, 370–378 (2004).

  44. 44.

    Ivorra, C. et al. A mechanism of AP-1 suppression through interaction of c-Fos with lamin A/C. Genes Dev. 20, 307–320 (2006).

  45. 45.

    Osmanagic-Myers, S., Dechat, T. & Foisner, R. Lamins at the crossroads of mechanosignaling. Genes Dev. 29, 225–237 (2015).

  46. 46.

    Dorner, D. et al. Lamina-associated polypeptide 2α regulates cell cycle progression and differentiation via the retinoblastoma–E2F pathway. J. Cell Biol. 173, 83–93 (2006).

  47. 47.

    Ho, C. Y., Jaalouk, D. E., Vartiainen, M. K. & Lammerding, J. Lamin A/C and emerin regulate MKL1–SRF activity by modulating actin dynamics. Nature 497, 507–511 (2013).

  48. 48.

    Harr, J. C. et al. Directed targeting of chromatin to the nuclear lamina is mediated by chromatin state and A-type lamins. J. Cell Biol. 208, 33–52 (2015).

  49. 49.

    Solovei, I. et al. LBR and lamin A/C sequentially tether peripheral heterochromatin and inversely regulate differentiation. Cell 152, 584–598 (2013).

  50. 50.

    Harada, T. et al. Nuclear lamin stiffness is a barrier to 3D migration, but softness can limit survival. J. Cell Biol. 204, 669–682 (2014).

  51. 51.

    Lee, J. S. et al. Nuclear lamin A/C deficiency induces defects in cell mechanics, polarization, and migration. Biophys. J. 93, 2542–2552 (2007).

  52. 52.

    Davidson, P. M., Denais, C., Bakshi, M. C. & Lammerding, J. Nuclear deformability constitutes a rate-limiting step during cell migration in 3-D environments. Cell. Mol. Bioeng. 7, 293–306 (2014).

  53. 53.

    Robijns, J. et al. In silico synchronization reveals regulators of nuclear ruptures in lamin A/C deficient model cells. Sci. Rep. 6, 30325 (2016).

  54. 54.

    Vargas, J. D., Hatch, E. M., Anderson, D. J. & Hetzer, M. W. Transient nuclear envelope rupturing during interphase in human cancer cells. Nucleus 3, 88–100 (2012).

  55. 55.

    Zwerger, M. et al. Myopathic lamin mutations impair nuclear stability in cells and tissue and disrupt nucleo-cytoskeletal coupling. Hum. Mol. Genet. 22, 2335–2349 (2013).

  56. 56.

    Folker, E. S., Ostlund, C., Luxton, G. W., Worman, H. J. & Gundersen, G. G. Lamin A variants that cause striated muscle disease are defective in anchoring transmembrane actin-associated nuclear lines for nuclear movement. Proc. Natl Acad. Sci. USA 108, 131–136 (2011).

  57. 57.

    Bertrand, A. T. et al. Cellular microenvironments reveal defective mechanosensing responses and elevated YAP signaling in LMNA-mutated muscle precursors. J. Cell Sci. 127, 2873–2884 (2014).

  58. 58.

    Cupesi, M. et al. Attenuated hypertrophic response to pressure overload in a lamin A/C haploinsufficiency mouse. J. Mol. Cell. Cardiol. 48, 1290–1297 (2010).

  59. 59.

    Buxboim, A. et al. Coordinated increase of nuclear tension and lamin-A with matrix stiffness out-competes lamin-B receptor which favors soft tissue phenotypes. Mol. Biol. Cell 28, 3333–3348 (2017).

  60. 60.

    Ihalainen, T. O. et al. Differential basal-to-apical accessibility of lamin A/C epitopes in the nuclear lamina regulated by changes in cytoskeletal tension. Nat. Mater. 14, 1252–1261 (2015).

  61. 61.

    Swift, J. et al. Nuclear lamin-A scales with tissue stiffness and enhances matrix-directed differentiation. Science 341, 1240104 (2013).

  62. 62.

    Sullivan, T. et al. Loss of A-type lamin expression compromises nuclear envelope integrity leading to muscular dystrophy. J. Cell Biol. 147, 913–920 (1999).

  63. 63.

    Kubben, N. et al. Post-natal myogenic and adipogenic developmental: defects and metabolic impairment upon loss of A-type lamins. Nucleus 2, 195–207 (2011).

  64. 64.

    Alam, S. G. et al. The nucleus is an intracellular propagator of tensile forces in NIH 3T3 fibroblasts. J. Cell Sci. 128, 1901–1911 (2015).

  65. 65.

    Crisp, M. & Burke, B. The nuclear envelope as an integrator of nuclear and cytoplasmic architecture. FEBS Lett. 582, 2023–2032 (2008).

  66. 66.

    Chang, W., Worman, H. J. & Gundersen, G. G. Accessorizing and anchoring the LINC complex for multifunctionality. J. Cell Biol. 208, 11–22 (2015).

  67. 67.

    Rajgor, D. & Shanahan, C. M. Nesprins: from the nuclear envelope and beyond. Expert Rev. Mol. Med. 15, e5 (2013).

  68. 68.

    Wilson, M. H. & Holzbaur, E. L. Nesprins anchor kinesin-1 motors to the nucleus to drive nuclear distribution in muscle cells. Development 142, 218–228 (2015).

  69. 69.

    Fridolfsson, H. N., Ly, N., Meyerzon, M. & Starr, D. A. UNC-83 coordinates kinesin-1 and dynein activities at the nuclear envelope during nuclear migration. Dev. Biol. 338, 237–250 (2010).

  70. 70.

    Wilhelmsen, K. et al. Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. J. Cell Biol. 171, 799–810 (2005).

  71. 71.

    Roux, K. J. et al. Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization. Proc. Natl Acad. Sci. USA 106, 2194–2199 (2009).

  72. 72.

    Horn, H. F. LINC complex proteins in development and disease. Curr. Top. Dev. Biol. 109, 287–321 (2014).

  73. 73.

    Elosegui-Artola, A. et al. Force triggers YAP nuclear entry by regulating transport across nuclear pores. Cell 171, 1397–1410 (2017).

  74. 74.

    Thiam, H. R. et al. Perinuclear Arp2/3-driven actin polymerization enables nuclear deformation to facilitate cell migration through complex environments. Nat. Commun. 7, 10997 (2016).

  75. 75.

    Banerjee, I. et al. Targeted ablation of nesprin 1 and nesprin 2 from murine myocardium results in cardiomyopathy, altered nuclear morphology and inhibition of the biomechanical gene response. PLoS Genet. 10, e1004114 (2014).

  76. 76.

    Lottersberger, F., Karssemeijer, R. A., Dimitrova, N. & de Lange, T. 53BP1 and the LINC complex promote microtubule-dependent DSB mobility and DNA repair. Cell 163, 880–893 (2015).

  77. 77.

    Makhija, E., Jokhun, D. S. & Shivashankar, G. V. Nuclear deformability and telomere dynamics are regulated by cell geometric constraints. Proc. Natl Acad. Sci. USA 113, 32–40 (2016).

  78. 78.

    Keeling, M. C., Flores, L. R., Dodhy, A. H., Murray, E. R. & Gavara, N. Actomyosin and vimentin cytoskeletal networks regulate nuclear shape, mechanics and chromatin organization. Sci. Rep. 7, 5219 (2017).

  79. 79.

    Hatch, E. M. & Hetzer, M. W. Nuclear envelope rupture is induced by actin-based nucleus confinement. J. Cell Biol. 215, 27–36 (2016).

  80. 80.

    Yeung, T. et al. Effects of substrate stiffness on cell morphology, cytoskeletal structure, and adhesion. Cell Motil. Cytoskelet. 60, 24–34 (2005).

  81. 81.

    Swift, J. & Discher, D. E. The nuclear lamina is mechano-responsive to ECM elasticity in mature tissue. J. Cell Sci. 127, 3005–3015 (2014).

  82. 82.

    Staszewska, I., Fischer, I. & Wiche, G. Plectin isoform 1-dependent nuclear docking of desmin networks affects myonuclear architecture and expression of mechanotransducers. Hum. Mol. Genet. 24, 7373–7389 (2015).

  83. 83.

    Konieczny, P. et al. Myofiber integrity depends on desmin network targeting to Z-disks and costameres via distinct plectin isoforms. J. Cell Biol. 181, 667–681 (2008).

  84. 84.

    Stroud, M. J. et al. Nesprin 1α2 is essential for mouse postnatal viability and nuclear positioning in skeletal muscle. J. Cell Biol. 216, 1915–1924 (2017).

  85. 85.

    Bone, C. R. & Starr, D. A. Nuclear migration events throughout development. J. Cell Sci. 129, 1951–1961 (2016).

  86. 86.

    Collins, M. A. et al. Emery–Dreifuss muscular dystrophy-linked genes and centronuclear myopathy-linked genes regulate myonuclear movement by distinct mechanisms. Mol. Biol. Cell 28, 2303–2317 (2017).

  87. 87.

    Roman, W. et al. Myofibril contraction and crosslinking drive nuclear movement to the periphery of skeletal muscle. Nat. Cell Biol. 19, 1189–1201 (2017).

  88. 88.

    Zhang, Q. et al. Nesprin-1 and -2 are involved in the pathogenesis of Emery Dreifuss muscular dystrophy and are critical for nuclear envelope integrity. Hum. Mol. Genet. 16, 2816–2833 (2007).

  89. 89.

    Puckelwartz, M. J. et al. Disruption of nesprin-1 produces an Emery Dreifuss muscular dystrophy-like phenotype in mice. Hum. Mol. Genet. 18, 607–620 (2009).

  90. 90.

    Stroud, M. J., Banerjee, I., Veevers, J. & Chen, J. Linker of nucleoskeleton and cytoskeleton complex proteins in cardiac structure, function, and disease. Circ. Res. 114, 538–548 (2014).

  91. 91.

    De Vos, W. H. et al. Repetitive disruptions of the nuclear envelope invoke temporary loss of cellular compartmentalization in laminopathies. Hum. Mol. Genet. 20, 4175–4186 (2011).

  92. 92.

    Zuela, N., Zwerger, M., Levin, T., Medalia, O. & Gruenbaum, Y. Impaired mechanical response of an EDMD mutation leads to motility phenotypes that are repaired by loss of prenylation. J. Cell Sci. 129, 1781–1791 (2016).

  93. 93.

    Gonzalez-Granado, J. M. et al. Nuclear envelope lamin-A couples actin dynamics with immunological synapse architecture and T cell activation. Sci. Signal. 7, ra37 (2014).

  94. 94.

    Khatau, S. B. et al. The distinct roles of the nucleus and nucleus-cytoskeleton connections in three-dimensional cell migration. Sci. Rep. 2, 488 (2012).

  95. 95.

    Chang, W., Antoku, S., Ostlund, C., Worman, H. J. & Gundersen, G. G. Linker of nucleoskeleton and cytoskeleton (LINC) complex-mediated actin-dependent nuclear positioning orients centrosomes in migrating myoblasts. Nucleus 6, 77–88 (2015).

  96. 96.

    King, S. J. et al. Nesprin-1 and nesprin-2 regulate endothelial cell shape and migration. Cytoskeleton 71, 423–434 (2014).

  97. 97.

    Horn, H. F. et al. The LINC complex is essential for hearing. J. Clin. Invest. 123, 740–750 (2013).

  98. 98.

    Potter, C. et al. Multiple isoforms of nesprin1 are integral components of ciliary rootlets. Curr. Biol. 27, 2014–2022 (2017).

  99. 99.

    Stewart, R. M. et al. Nuclear–cytoskeletal linkages facilitate cross talk between the nucleus and intercellular adhesions. J. Cell Biol. 209, 403–418 (2015).

  100. 100.

    Zhang, X. et al. SUN1/2 and syne/nesprin-1/2 complexes connect centrosome to the nucleus during neurogenesis and neuronal migration in mice. Neuron 64, 173–187 (2009).

  101. 101.

    Khatau, S. B. et al. A perinuclear actin cap regulates nuclear shape. Proc. Natl Acad. Sci. USA 106, 19017–19022 (2009).

  102. 102.

    Driscoll, T. P., Cosgrove, B. D., Heo, S. J., Shurden, Z. E. & Mauck, R. L. Cytoskeletal to nuclear strain transfer regulates YAP signaling in mesenchymal stem cells. Biophys. J. 108, 2783–2793 (2015).

  103. 103.

    Ramdas, N. M. & Shivashankar, G. V. Cytoskeletal control of nuclear morphology and chromatin organization. J. Mol. Biol. 427, 695–706 (2015).

  104. 104.

    Geyer, P. K., Vitalini, M. W. & Wallrath, L. L. Nuclear organization: taking a position on gene expression. Curr. Opin. Cell Biol. 23, 354–359 (2011).

  105. 105.

    Chen, H. et al. Functional organization of the human 4D Nucleome. Proc. Natl Acad. Sci. USA 112, 8002–8007 (2015).

  106. 106.

    Misteli, T. Beyond the sequence: cellular organization of genome function. Cell 128, 787–800 (2007).

  107. 107.

    Sexton, T., Schober, H., Fraser, P. & Gasser, S. M. Gene regulation through nuclear organization. Nat. Struct. Mol. Biol. 14, 1049–1055 (2007).

  108. 108.

    Zullo, J. M. et al. DNA sequence-dependent compartmentalization and silencing of chromatin at the nuclear lamina. Cell 149, 1474–1487 (2012).

  109. 109.

    Wang, Y., Nagarajan, M., Uhler, C. & Shivashankar, G. V. Orientation and repositioning of chromosomes correlate with cell geometry-dependent gene expression. Mol. Biol. Cell 28, 1997–2009 (2017).

  110. 110.

    Spagnol, S. T. & Dahl, K. N. Spatially resolved quantification of chromatin condensation through differential local rheology in cell nuclei fluorescence lifetime imaging. PLoS ONE 11, e0146244 (2016).

  111. 111.

    Booth-Gauthier, E. A., Alcoser, T. A., Yang, G. & Dahl, K. N. Force-induced changes in subnuclear movement and rheology. Biophys. J. 103, 2423–2431 (2012).

  112. 112.

    Poh, Y. C. et al. Dynamic force-induced direct dissociation of protein complexes in a nuclear body in living cells. Nat. Commun. 3, 866 (2012).

  113. 113.

    Spagnol, S. T., Armiger, T. J. & Dahl, K. N. Mechanobiology of chromatin and the nuclear interior. Cell. Mol. Bioeng. 9, 268–276 (2016).

  114. 114.

    Chalut, K. J. et al. Chromatin decondensation and nuclear softening accompany Nanog downregulation in embryonic stem cells. Biophys. J. 103, 2060–2070 (2012).

  115. 115.

    Dahl, K. N., Engler, A. J., Pajerowski, J. D. & Discher, D. E. Power-law rheology of isolated nuclei with deformation mapping of nuclear substructures. Biophys. J. 89, 2855–2864 (2005).

  116. 116.

    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).

  117. 117.

    Stephens, A. D. et al. Chromatin histone modifications and rigidity affect nuclear morphology independent of lamins. Mol. Biol. Cell 29, 220–233 (2018).

  118. 118.

    Cui, Y. & Bustamante, C. Pulling a single chromatin fiber reveals the forces that maintain its higher-order structure. Proc. Natl Acad. Sci. USA 97, 127–132 (2000).

  119. 119.

    Bennett, R. R. et al. Elastic-fluid model for DNA damage and mutation from nuclear fluid segregation due to cell migration. Biophys. J. 112, 2271–2279 (2017).

  120. 120.

    Irianto, J. et al. DNA damage follows repair factor depletion and portends genome variation in cancer cells after pore migration. Curr. Biol. 27, 210–223 (2017).

  121. 121.

    Buxboim, A. et al. Matrix elasticity regulates lamin-A, C phosphorylation and turnover with feedback to actomyosin. Curr. Biol. 24, 1909–1917 (2014).

  122. 122.

    Tifft, K. E., Bradbury, K. A. & Wilson, K. L. Tyrosine phosphorylation of nuclear-membrane protein emerin by Src, Abl and other kinases. J. Cell Sci. 122, 3780–3790 (2009).

  123. 123.

    Enyedi, B., Jelcic, M. & Niethammer, P. The cell nucleus serves as a mechanotransducer of tissue damage-induced inflammation. Cell 165, 1160–1170 (2016).

  124. 124.

    Enyedi, B. & Niethammer, P. Nuclear membrane stretch and its role in mechanotransduction. Nucleus 8, 156–161 (2017).

  125. 125.

    Heald, R. & Cohen-Fix, O. Morphology and function of membrane-bound organelles. Curr. Opin. Cell Biol. 26, 79–86 (2014).

  126. 126.

    Shibata, Y. et al. Mechanisms determining the morphology of the peripheral ER. Cell 143, 774–788 (2010).

  127. 127.

    Isermann, P. & Lammerding, J. Consequences of a tight squeeze: nuclear envelope rupture and repair. Nucleus 8, 268–274 (2017).

  128. 128.

    Denais, C. M. et al. Nuclear envelope rupture and repair during cancer cell migration. Science 352, 353–358 (2016).

  129. 129.

    Raab, M. et al. ESCRT III repairs nuclear envelope ruptures during cell migration to limit DNA damage and cell death. Science 352, 359–362 (2016).

  130. 130.

    Takaki, T. et al. Actomyosin drives cancer cell nuclear dysmorphia and threatens genome stability. Nat. Commun. 8, 16013 (2017).

  131. 131.

    Lammerding, J. & Wolf, K. Nuclear envelope rupture: actin fibers are putting the squeeze on the nucleus. J. Cell Biol. 215, 5–8 (2016).

  132. 132.

    Le Berre, M., Aubertin, J. & Piel, M. Fine control of nuclear confinement identifies a threshold deformation leading to lamina rupture and induction of specific genes. Integr. Biol. 4, 1406–1414 (2012).

  133. 133.

    Sun, L., Wu, J., Du, F., Chen, X. & Chen, Z. J. Cyclic GMP–AMP synthase is a cytosolic DNA sensor that activates the type I interferon pathway. Science 339, 786–791 (2013).

  134. 134.

    Gluck, S. et al. Innate immune sensing of cytosolic chromatin fragments through cGAS promotes senescence. Nat. Cell Biol. 19, 1061–1070 (2017).

  135. 135.

    Harding, S. M. et al. Mitotic progression following DNA damage enables pattern recognition within micronuclei. Nature 548, 466–470 (2017).

  136. 136.

    Mackenzie, K. J. et al. cGAS surveillance of micronuclei links genome instability to innate immunity. Nature 548, 461–465 (2017).

  137. 137.

    Dou, Z. et al. Cytoplasmic chromatin triggers inflammation in senescence and cancer. Nature 550, 402–406 (2017).

  138. 138.

    Chug, H., Trakhanov, S., Hulsmann, B. B., Pleiner, T. & Gorlich, D. Crystal structure of the metazoan Nup62•Nup58•Nup54 nucleoporin complex. Science 350, 106–110 (2015).

  139. 139.

    Stuwe, T. et al. Architecture of the fungal nuclear pore inner ring complex. Science 350, 56–64 (2015).

  140. 140.

    Solmaz, S. R., Blobel, G. & Melcak, I. Ring cycle for dilating and constricting the nuclear pore. Proc. Natl Acad. Sci. USA 110, 5858–5863 (2013).

  141. 141.

    Liu, Q. et al. Functional association of Sun1 with nuclear pore complexes. J. Cell Biol. 178, 785–798 (2007).

  142. 142.

    Jahed, Z., Soheilypour, M., Peyro, M. & Mofrad, M. R. The LINC and NPC relationship—it’s complicated! J. Cell Sci. 129, 3219–3229 (2016).

  143. 143.

    Fahrenkrog, B. & Aebi, U. The nuclear pore complex: nucleocytoplasmic transport and beyond. Nat. Rev. Mol. Cell Biol. 4, 757–766 (2003).

  144. 144.

    Dupont, S. et al. Role of YAP/TAZ in mechanotransduction. Nature 474, 179–183 (2011).

  145. 145.

    Dorner, D., Gotzmann, J. & Foisner, R. Nucleoplasmic lamins and their interaction partners, LAP2α, Rb, and BAF, in transcriptional regulation. FEBS J. 274, 1362–1373 (2007).

  146. 146.

    Markiewicz, E., Dechat, T., Foisner, R., Quinlan, R. A. & Hutchison, C. J. Lamin A/C binding protein LAP2α is required for nuclear anchorage of retinoblastoma protein. Mol. Biol. Cell 13, 4401–4413 (2002).

  147. 147.

    Ma, H. et al. Multicolor CRISPR labeling of chromosomal loci in human cells. Proc. Natl Acad. Sci. USA 112, 3002–3007 (2015).

  148. 148.

    Shao, S. et al. Long-term dual-color tracking of genomic loci by modified sgRNAs of the CRISPR/Cas9 system. Nucleic Acids Res. 44, e86 (2016).

  149. 149.

    Donnert, G. et al. Macromolecular-scale resolution in biological fluorescence microscopy. Proc. Natl Acad. Sci. USA 103, 11440–11445 (2006).

  150. 150.

    Belton, J. M. et al. Hi-C: a comprehensive technique to capture the conformation of genomes. Methods 58, 268–276 (2012).

  151. 151.

    Nagano, T. et al. Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502, 59–64 (2013).

  152. 152.

    Buenrostro, J. D., Wu, B., Chang, H. Y. & Greenleaf, W. J. ATAC-seq: a method for assaying chromatin accessibility genome-wide. Curr. Protoc. Mol. Biol. 109, 21.29.1–21.29.9 (2015).

  153. 153.

    Neelam, S. et al. Direct force probe reveals the mechanics of nuclear homeostasis in the mammalian cell. Proc. Natl Acad. Sci. USA 112, 5720–5725 (2015).

  154. 154.

    Krause, M., Te Riet, J. & Wolf, K. Probing the compressibility of tumor cell nuclei by combined atomic force-confocal microscopy. Phys. Biol. 10, 065002 (2013).

  155. 155.

    Arsenovic, P. T., Bathula, K. & Conway, D. E. A protocol for using Förster resonance energy transfer (FRET)-force biosensors to measure mechanical forces across the nuclear LINC complex. J. Vis. Exp. 122, e54902 (2017).

  156. 156.

    Grashoff, C. et al. Measuring mechanical tension across vinculin reveals regulation of focal adhesion dynamics. Nature 466, 263–266 (2010).

  157. 157.

    Meng, F., Suchyna, T. M., Lazakovitch, E., Gronostajski, R. M. & Sachs, F. Real time FRET based detection of mechanical stress in cytoskeletal and extracellular matrix proteins. Cell. Mol. Bioeng. 4, 148–159 (2011).

  158. 158.

    Fedorchak, G. & Lammerding, J. Cell Microharpooning to study nucleo-cytoskeletal coupling. Methods Mol. Biol. 1411, 241–254 (2016).

  159. 159.

    Lombardi, M. L., Zwerger, M. & Lammerding, J. Biophysical assays to probe the mechanical properties of the interphase cell nucleus: substrate strain application and microneedle manipulation. J. Vis. Exp. 55, e3087 (2011).

  160. 160.

    Britton, S., Coates, J. & Jackson, S. P. A new method for high-resolution imaging of Ku foci to decipher mechanisms of DNA double-strand break repair. J. Cell Biol. 202, 579–595 (2013).

  161. 161.

    Bianchini, P., Cardarelli, F., Di Luca, M., Diaspro, A. & Bizzarri, R. Nanoscale protein diffusion by STED-based pair correlation analysis. PLoS ONE 9, e99619 (2014).

  162. 162.

    Failla, A. V., Spoeri, U., Albrecht, B., Kroll, A. & Cremer, C. Nanosizing of fluorescent objects by spatially modulated illumination microscopy. Appl. Opt. 41, 7275–7283 (2002).

  163. 163.

    Hildenbrand, G. et al. Nano-sizing of specific gene domains in intact human cell nuclei by spatially modulated illumination light microscopy. Biophys. J. 88, 4312–4318 (2005).

  164. 164.

    Reymann, J. et al. High-precision structural analysis of subnuclear complexes in fixed and live cells via spatially modulated illumination (SMI) microscopy. Chromosome Res. 16, 367–382 (2008).

  165. 165.

    Cremer, C., Szczurek, A., Schock, F., Gourram, A. & Birk, U. Super-resolution microscopy approaches to nuclear nanostructure imaging. Methods 123, 11–32 (2017).

  166. 166.

    Lesterlin, C., Ball, G., Schermelleh, L. & Sherratt, D. J. RecA bundles mediate homology pairing between distant sisters during DNA break repair. Nature 506, 249–253 (2014).

  167. 167.

    Schermelleh, L. et al. Subdiffraction multicolor imaging of the nuclear periphery with 3D structured illumination microscopy. Science 320, 1332–1336 (2008).

  168. 168.

    Henriques, R., Griffiths, C., Hesper Rego, E. & Mhlanga, M. M. PALM and STORM: unlocking live-cell super-resolution. Biopolymers 95, 322–331 (2011).

  169. 169.

    Boettiger, A. N. et al. Super-resolution imaging reveals distinct chromatin folding for different epigenetic states. Nature 529, 418–422 (2016).

  170. 170.

    Ricci, M. A., Manzo, C., Garcia-Parajo, M. F., Lakadamyali, M. & Cosma, M. P. Chromatin fibers are formed by heterogeneous groups of nucleosomes in vivo. Cell 160, 1145–1158 (2015).

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Acknowledgements

We apologize to all authors whose work could not be cited due to space constraints. This work was supported by awards from the National Institutes of Health (R01 HL082792 and U54 CA210184), the Department of Defense Breast Cancer Research Program (Breakthrough Award BC150580), the National Science Foundation (CAREER Award CBET-1254846 and MCB-1715606) and a Fleming Postdoctoral Fellowship to T.J.K.

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  1. Nancy E. and Peter C. Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA

    • Tyler J. Kirby
    •  & Jan Lammerding
  2. Weill Institute for Cell and Molecular Biology, Cornell University, Ithaca, NY, USA

    • Tyler J. Kirby
    •  & Jan Lammerding

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The authors declare no competing interests.

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Correspondence to Jan Lammerding.

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https://doi.org/10.1038/s41556-018-0038-y