Mechanics and functional consequences of nuclear deformations.

[en] As the home of cellular genetic information, the nucleus has a critical role in determining cell fate and function in response to various signals and stimuli. In addition to biochemical inputs, the nucleus is constantly exposed to intrinsic and extrinsic mechanical forces that trigger dynamic changes in nuclear structure and morphology. Emerging data suggest that the physical deformation of the nucleus modulates many cellular and nuclear functions. These functions have long been considered to be downstream of cytoplasmic signalling pathways and dictated by gene expression. In this Review, we discuss an emerging perspective on the mechanoregulation of the nucleus that considers the physical connections from chromatin to nuclear lamina and cytoskeletal filaments as a single mechanical unit. We describe key mechanisms of nuclear deformations in time and space and provide a critical review of the structural and functional adaptive responses of the nucleus to deformations. We then consider the contribution of nuclear deformations to the regulation of important cellular functions, including muscle contraction, cell migration and human disease pathogenesis. Collectively, these emerging insights shed new light on the dynamics of nuclear deformations and their roles in cellular mechanobiology.

Disciplines :

Physical, chemical, mathematical & earth Sciences: Multidisciplinary, general & others

Author, co-author :

Kalukula, Yohalie ; Université de Mons - UMONS > Faculté des Sciences > Service du Laboratoire Interfaces et Fluides Complexes

Stephens, Andrew D; Biology Department, University of Massachusetts Amherst, Amherst, MA, USA

Lammerding, Jan ^✱; Weill Institute for Cell and Molecular Biology, Cornell University, Ithaca, NY, USA. jan.lammerding@cornell.edu ; Nancy E. and Peter C. Meinig School of Biomedical Engineering, Cornell University, Ithaca, NY, USA. jan.lammerding@cornell.edu

Gabriele, Sylvain ^✱; Université de Mons - UMONS > Faculté des Sciences > Service du Laboratoire Interfaces et Fluides Complexes

^✱ These authors have contributed equally to this work.

Language :

English

Title :

Mechanics and functional consequences of nuclear deformations.

Publication date :

05 May 2022

Journal title :

Nature Reviews. Molecular Cell Biology

ISSN :

1471-0072

eISSN :

1471-0080

Publisher :

Nature Research, England

Peer reviewed :

Peer Reviewed verified by ORBi

Additional URL :

https://www.nature.com/articles/s41580-022-00480-z.pdf

Research institute :

R100 - Institut des Biosciences

Funding text :

The authors apologize to all authors whose work could not be included owing to space constraints. A.D.S. is supported by the Pathway to Independence Award (R00GM123195) and 4D Nucleome 2 centre grant (1UM1HG011536). S.G. acknowledges funding from FEDER Prostem Research Project no. 1510614 (Wallonia DG06), the F.R.S.-FNRS Epiforce Project no. T.0092.21 and the Interreg MAT(T)ISSE project, which is financially supported by Interreg France-Wallonie-Vlaanderen (Fonds Européen de Développement Régional, FEDER-ERDF). Y.K. is financially supported by FRIA (F.R.S.-FNRS) and FRMH (Fonds pour la Recherche Médicale dans le Hainaut). J.L. is supported by awards from the National Institutes of Health (R01HL082792, R01GM137605, U54CA210184), the National Science Foundation (URoL-2022048) and the VolkswagenStiftung (Az. 96733).

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Bibliography

Lammerding, J. Mechanics of the nucleus. Compr. Physiol. 1, 783–807 (2013).
Szczesny, S. E. & Mauck, R. L. The nuclear option: evidence implicating the cell nucleus in mechanotransduction. J. Biomech. Eng. 139, 0210061–02100616 (2017). DOI: 10.1115/1.4035350
Long, J. T. & Lammerding, J. Nuclear deformation lets cells gauge their physical confinement. Dev. Cell 56, 156–158 (2021). DOI: 10.1016/j.devcel.2021.01.002
Thomas, C. H., Collier, J. H., Sfeir, C. S. & Healy, K. E. Engineering gene expression and protein synthesis by modulation of nuclear shape. Proc. Natl Acad. Sci. USA 99, 1972–1977 (2002). DOI: 10.1073/pnas.032668799
Skinner, B. M. & Johnson, E. E. P. Nuclear morphologies: their diversity and functional relevance. Chromosoma 126, 195–212 (2017). DOI: 10.1007/s00412-016-0614-5
Gupta, S., Marcel, N., Sarin, A. & Shivashankar, G. V. Role of actin dependent nuclear deformation in regulating early gene expression. PLoS ONE 7, e53031 (2012). DOI: 10.1371/journal.pone.0053031
Miroshnikova, Y. A., Nava, M. M. & Wickström, S. A. Emerging roles of mechanical forces in chromatin regulation. J. Cell Sci. 130, 2243–2250 (2017).
Zink, D., Fischer, A. H. & Nickerson, J. A. Nuclear structure in cancer cells. Nat. Rev. Cancer 4, 677–687 (2004). DOI: 10.1038/nrc1430
Clippinger, S. R. et al. Disrupted mechanobiology links the molecular and cellular phenotypes in familial dilated cardiomyopathy. Proc. Natl Acad. Sci. USA 116, 17831–17840 (2019). DOI: 10.1073/pnas.1910962116
Franke, W. W., Scheer, U., Krohne, G. & Jarasch, E. D. The nuclear envelope and the architecture of the nuclear periphery. J. Cell Biol. 91, 39s–50s (1981). DOI: 10.1083/jcb.91.3.39s
Kim, D.-H. et al. Volume regulation and shape bifurcation in the cell nucleus. J. Cell Sci. 128, 3375–3385 (2015).
Jevtić, P. et al. The nucleoporin ELYS regulates nuclear size by controlling NPC number and nuclear import capacity. EMBO Rep. 20, e47283 (2019). DOI: 10.15252/embr.201847283
García-González, A. et al. The effect of cell morphology on the permeability of the nuclear envelope to diffusive factors. Front. Physiol. 9, 925 (2018). DOI: 10.3389/fphys.2018.00925
Donnaloja, F., Jacchetti, E., Soncini, M. & Raimondi, M. T. Mechanosensing at the nuclear envelope by nuclear pore complex stretch activation and its effect in physiology and pathology. Front. Physiol. 10, 896 (2019). DOI: 10.3389/fphys.2019.00896
Schuller, A. P. et al. The cellular environment shapes the nuclear pore complex architecture. Nature 598, 667–671 (2021). DOI: 10.1038/s41586-021-03985-3
Nava, M. M. et al. Heterochromatin-driven nuclear softening protects the genome against mechanical stress-induced damage. Cell 181, 800–817.e22 (2020). This article demonstrates how cyclic strain application can induce transient chromatin modifications, which, together with slower realignment of cells perpendicular to the stretch direction, help protect the cells from mechanically induced DNA damage. DOI: 10.1016/j.cell.2020.03.052
Lomakin, A. J. et al. The nucleus acts as a ruler tailoring cell responses to spatial constraints. Science 370, eaba2894 (2020). This article, together with concurrently published work by Venturini et al. (ref.127), demonstrates how physical confinement that compresses the nucleus triggers increased cell cortical contractility via recruitment of cPLA2 to the stretched nuclear membranes. DOI: 10.1126/science.aba2894
Turgay, Y. et al. The molecular architecture of lamins in somatic cells. Nature 543, 261–264 (2017). DOI: 10.1038/nature21382
Tenga, R. & Medalia, O. Structure and unique mechanical aspects of nuclear lamin filaments. Curr. Opin. Struct. Biol. 64, 152–159 (2020). DOI: 10.1016/j.sbi.2020.06.017
de Leeuw, R., Gruenbaum, Y. & Medalia, O. Nuclear lamins: thin filaments with major functions. Trends Cell Biol. 28, 34–45 (2018). DOI: 10.1016/j.tcb.2017.08.004
Shimi, T. et al. Structural organization of nuclear lamins A, C, B1, and B2 revealed by superresolution microscopy. Mol. Biol. Cell 26, 4075–4086 (2015). DOI: 10.1091/mbc.E15-07-0461
Kolb, T., Maass, K., Hergt, M., Aebi, U. & Herrmann, H. Lamin A and lamin C form homodimers and coexist in higher complex forms both in the nucleoplasmic fraction and in the lamina of cultured human cells. Nucleus 2, 425–433 (2011). DOI: 10.4161/nucl.2.5.17765
Nmezi, B. et al. Concentric organization of A- and B-type lamins predicts their distinct roles in the spatial organization and stability of the nuclear lamina. Proc. Natl Acad. Sci. USA 116, 4307–4315 (2019). DOI: 10.1073/pnas.1810070116
Naetar, N. et al. LAP2alpha maintains a mobile and low assembly state of A-type lamins in the nuclear interior. eLife 10, e63476 (2021). DOI: 10.7554/eLife.63476
Pascual-Reguant, L. et al. Lamin B1 mapping reveals the existence of dynamic and functional euchromatin lamin B1 domains. Nat. Commun. 9, 3420 (2018). DOI: 10.1038/s41467-018-05912-z
Cho, S. et al. Mechanosensing by the lamina protects against nuclear rupture, DNA damage, and cell-cycle arrest. Dev. Cell 49, 920–935 (2019). DOI: 10.1016/j.devcel.2019.04.020
Koushki, N. et al. Lamin A redistribution mediated by nuclear deformation determines dynamic localization of YAP. bioRxiv 10.1101/2020.03.19.998708 (2020). DOI: 10.1101/2020.03.19.998708
Zhu, L. & Brangwynne, C. P. Nuclear bodies: the emerging biophysics of nucleoplasmic phases. Curr. Opin. Cell Biol. 34, 23–30 (2015). DOI: 10.1016/j.ceb.2015.04.003
Buchwalter, A., Kaneshiro, J. M. & Hetzer, M. W. Coaching from the sidelines: the nuclear periphery in genome regulation. Nat. Rev. Genet. 20, 39–50 (2019). DOI: 10.1038/s41576-018-0063-5
Bizhanova, A. & Kaufman, P. Close to the edge: heterochromatin at the nucleolar and nuclear peripheries. Biochim. Biophys. Acta 1864, 194666 (2021). DOI: 10.1016/j.bbagrm.2020.194666
Miron, E. et al. Chromatin arranges in chains of mesoscale domains with nanoscale functional topography independent of cohesin. Sci. Adv. 6, eaba8811 (2020). DOI: 10.1126/sciadv.aba8811
Cho, S., Irianto, J. & Discher, D. E. Mechanosensing by the nucleus: from pathways to scaling relationships. J. Cell Biol. 216, 305–315 (2017). DOI: 10.1083/jcb.201610042
Maeshima, K., Tamura, S., Hansen, J. C. & Itoh, Y. Fluid-like chromatin: toward understanding the real chromatin organization present in the cell. Curr. Opin. Cell Biol. 64, 77–89 (2020). DOI: 10.1016/j.ceb.2020.02.016
Hansen, J. C., Maeshima, K. & Hendzel, M. J. The solid and liquid states of chromatin. Epigenetics Chromatin 14, 50 (2021). DOI: 10.1186/s13072-021-00424-5
Jahed, Z., Domkam, N., Ornowski, J., Yerima, G. & Mofrad, M. R. K. Molecular models of LINC complex assembly at the nuclear envelope. J. Cell Sci. 134, jcs258194 (2021). DOI: 10.1242/jcs.258194
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). First function demonstration of force transmission from the cytoskeleton to the nucleus via the LINC complex. DOI: 10.1074/jbc.M111.233700
Denis, K. B. et al. The LINC complex is required for endothelial cell adhesion and adaptation to shear stress and cyclic stretch. Mol. Biol. Cell 32, 1654–1663 (2021). DOI: 10.1091/mbc.E20-11-0698
Splinter, D. et al. Bicaudal D2, Tynein, and kinesin-1 associate with nuclear pore complexes and regulate centrosome and nuclear positioning during mitotic entry. PLoS Biol. 8, e1000350 (2010). DOI: 10.1371/journal.pbio.1000350
Salpingidou, G., Smertenko, A., Hausmanowa-Petrucewicz, I., Hussey, P. J. & Hutchison, C. J. A novel role for the nuclear membrane protein emerin in association of the centrosome to the outer nuclear membrane. J. Cell Biol. 178, 897–904 (2007). DOI: 10.1083/jcb.200702026
Sosa, B. A., Rothballer, A., Kutay, U. & Schwartz, T. U. LINC complexes form by binding of three KASH peptides to domain interfaces of trimeric SUN proteins. Cell 149, 1035–1047 (2012). First detailed structural characterization of the LINC complex, elucidating how forces can be transmitted across the nesprin–SUN protein interface. DOI: 10.1016/j.cell.2012.03.046
Cruz, V. E., Demircioglu, F. E. & Schwartz, T. U. Structural analysis of different LINC complexes reveals distinct binding modes. J. Mol. Biol. 432, 6028–6041 (2020). DOI: 10.1016/j.jmb.2020.09.019
Rajgor, D. & Shanahan, C. M. Nesprins: from the nuclear envelope and beyond. Expert Rev. Mol. Med. 15, e5 (2013). DOI: 10.1017/erm.2013.6
Wilson, M. H. & Holzbaur, E. L. F. Nesprins anchor kinesin-1 motors to the nucleus to drive nuclear distribution in muscle cells. Development 142, 218–228 (2014). DOI: 10.1242/dev.114769
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). DOI: 10.1016/j.ydbio.2009.12.004
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). DOI: 10.1083/jcb.200506083
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). DOI: 10.1073/pnas.0808602106
Horn, H. F. et al. A mammalian KASH domain protein coupling meiotic chromosomes to the cytoskeleton. J. Cell Biol. 202, 1023–1039 (2013). DOI: 10.1083/jcb.201304004
Agrawal, A. & Lele, T. P. Mechanics of nuclear membranes. J. Cell Sci. 132, jcs229245 (2019). DOI: 10.1242/jcs.229245
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). DOI: 10.1091/mbc.E19-01-0027
Versaevel, M. et al. Super-resolution microscopy reveals LINC complex recruitment at nuclear indentation sites. Sci. Rep. 4, 7362 (2015). DOI: 10.1038/srep07362
Davidson, P. M. et al. Nesprin-2 accumulates at the front of the nucleus during confined cell migration. EMBO Rep. 21, e49910 (2020). DOI: 10.15252/embr.201949910
Lim, S. M., Cruz, V. E., Antoku, S., Gundersen, G. G. & Schwartz, T. U. Structures of FHOD1-nesprin1/2 complexes reveal alternate binding modes for the FH3 domain of formins. Structure 29, 540–552 (2021). DOI: 10.1016/j.str.2020.12.013
Saunders, C. A. et al. TorsinA controls TAN line assembly and the retrograde flow of dorsal perinuclear actin cables during rearward nuclear movement. J. Cell Biol. 216, 657–674 (2017). DOI: 10.1083/jcb.201507113
Gudise, S., Figueroa, R. A., Lindberg, R., Larsson, V. & Hallberg, E. Samp1 is functionally associated with the LINC complex and A-type lamina networks. J. Cell Sci. 124, 2077–2085 (2011). DOI: 10.1242/jcs.078923
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). DOI: 10.1093/hmg/ddt079
Hao, H. et al. The nesprin-1/-2 ortholog ANC-1 regulates organelle positioning in C. elegans independently from its KASH or actin-binding domains. eLife 10, e61069 (2021). DOI: 10.7554/eLife.61069
Versaevel, M., Riaz, M., Grevesse, T. & Gabriele, S. Cell confinement: putting the squeeze on the nucleus. Soft Matter 9, 6665–6676 (2013). DOI: 10.1039/c3sm00147d
Guilak, F., Tedrow, J. R. & Burgkart, R. Viscoelastic properties of the cell nucleus. Biochem. Biophys. Res. Commun. 269, 781–786 (2000). DOI: 10.1006/bbrc.2000.2360
Hobson, C. M., Falvo, M. R. & Superfine, R. A survey of physical methods for studying nuclear mechanics and mechanobiology. Appl. Bioeng. 5, 041508 (2021). DOI: 10.1063/5.0068126
Pajerowski, J. D., Dahl, K. N., Zhong, F. L., Sammak, P. J. & Discher, D. E. Physical plasticity of the nucleus in stem cell differentiation. Proc. Natl Acad. Sci. USA 104, 15619–15624 (2007). DOI: 10.1073/pnas.0702576104
Davidson, P. M. et al. High-throughput microfluidic micropipette aspiration device to probe time-scale dependent nuclear mechanics in intact cells. Lab. Chip 19, 3652–3663 (2019). DOI: 10.1039/C9LC00444K
Rowat, A. C., Foster, L. J., Nielsen, M. M., Weiss, M. & Ipsen, J. H. Characterization of the elastic properties of the nuclear envelope. J. R. Soc. Interface 2, 63–69 (2005). DOI: 10.1098/rsif.2004.0022
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). DOI: 10.1529/biophysj.105.062554
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). DOI: 10.1091/mbc.e16-09-0653
Grevesse, T., Dabiri, B. E., Parker, K. K. & Gabriele, S. Opposite rheological properties of neuronal microcompartments predict axonal vulnerability in brain injury. Sci. Rep. 5, 9475 (2015). DOI: 10.1038/srep09475
Lammerding, J. et al. Lamins A and C but not lamin B1 regulate nuclear mechanics. J. Biol. Chem. 281, 25768–25780 (2006). This report establishes the important role of lamins A/C in determining nuclear deformability. DOI: 10.1074/jbc.M513511200
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). DOI: 10.1073/pnas.1502111112
Lammerding, J. et al. Lamin A/C deficiency causes defective nuclear mechanics and mechanotransduction. J. Clin. Invest. 113, 370–378 (2004). First report of a role of lamins A/C in mediating nuclear mechanotransduction and providing nuclear stability to allow cells to withstand mechanical stress. DOI: 10.1172/JCI200419670
Lammerding, J. et al. Abnormal nuclear shape and impaired mechanotransduction in emerin-deficient cells. J. Cell Biol. 170, 781–791 (2005). DOI: 10.1083/jcb.200502148
Cao, X. et al. A chemomechanical model for nuclear morphology and stresses during cell transendothelial migration. Biophys. J. 111, 1541–1552 (2016). DOI: 10.1016/j.bpj.2016.08.011
Versaevel, M. et al. Probing cytoskeletal pre-stress and nuclear mechanics in endothelial cells with spatiotemporally controlled (de-)adhesion kinetics on micropatterned substrates. Cell Adhes. Migr. 11, 98–109 (2017). DOI: 10.1080/19336918.2016.1182290
Ferrera, D. et al. Lamin B1 overexpression increases nuclear rigidity in autosomal dominant leukodystrophy fibroblasts. FASEB J. 28, 3906–3918 (2014). DOI: 10.1096/fj.13-247635
Chen, N. Y. et al. An absence of lamin B1 in migrating neurons causes nuclear membrane ruptures and cell death. Proc. Natl Acad. Sci. USA 116, 25870–25879 (2019). DOI: 10.1073/pnas.1917225116
Hatch, E. M. & Hetzer, M. W. Nuclear envelope rupture is induced by actin-based nucleus confinement. J. Cell Biol. 215, 27–36 (2016). DOI: 10.1083/jcb.201603053
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). DOI: 10.1126/science.aad7611
Denais, C. M. et al. Nuclear envelope rupture and repair during cancer cell migration. Science 352, 353–358 (2016). Together with the work by Raab et al. (ref. 75), this is the first report of migration-induced nuclear envelope rupture and DNA damage, while also identifying a role of ESCRT proteins in interphase nuclear envelope resealing. DOI: 10.1126/science.aad7297
Zhang, Q. et al. Local, transient tensile stress on the nuclear membrane causes membrane rupture. Mol. Biol. Cell 30, 899–906 (2019). DOI: 10.1091/mbc.E18-09-0604
Furusawa, T. et al. Chromatin decompaction by the nucleosomal binding protein HMGN5 impairs nuclear sturdiness. Nat. Commun. 6, 6138 (2015). DOI: 10.1038/ncomms7138
Samwer, M. et al. DNA cross-bridging shapes a single nucleus from a set of mitotic chromosomes. Cell 170, 956–972 (2017). DOI: 10.1016/j.cell.2017.07.038
Wang, P. et al. WDR5 modulates cell motility and morphology and controls nuclear changes induced by a 3D environment. Proc. Natl Acad. Sci. USA 115, 8581–8586 (2018). DOI: 10.1073/pnas.1719405115
Serra-Marques, A. et al. The mitotic protein NuMA plays a spindle-independent role in nuclear formation and mechanics. J. Cell Biol. 219, e202004202 (2020). DOI: 10.1083/jcb.202004202
Tamashunas, A. C. et al. High-throughput gene screen reveals modulators of nuclear shape. Mol. Biol. Cell 31, 1392–1402 (2020). DOI: 10.1091/mbc.E19-09-0520
Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017). DOI: 10.1038/nature22822
Welsh, T. J., Shen, Y., Levin, A. & Knowles, T. P. J. Mechanobiology of protein droplets: force arises from disorder. Cell 175, 1457–1459 (2018). DOI: 10.1016/j.cell.2018.11.020
Gibson, B. A. et al. Organization of chromatin by intrinsic and regulated phase separation. Cell 179, 470–484 (2019). DOI: 10.1016/j.cell.2019.08.037
Zidovska, A. The rich inner life of the cell nucleus: dynamic organization, active flows, and emergent rheology. Biophys. Rev. 12, 1093–1106 (2020). DOI: 10.1007/s12551-020-00761-x
Jord, A. A. et al. Cytoplasmic forces functionally reorganize nuclear condensates in oocytes. bioRxiv 10.1101/2021.03.15.434387 (2021). DOI: 10.1101/2021.03.15.434387
Bracha, D. et al. Mapping local and global liquid phase behavior in living cells using photo-oligomerizable seeds. Cell 175, 1467–1480 (2018). DOI: 10.1016/j.cell.2018.10.048
Shin, Y. et al. Spatiotemporal control of intracellular phase transitions using light-activated optoDroplets. Cell 168, 159–171 (2017). DOI: 10.1016/j.cell.2016.11.054
Shin, Y. et al. Liquid nuclear condensates mechanically sense and restructure the genome. Cell 175, 1481–1491 (2018). DOI: 10.1016/j.cell.2018.10.057
Stephens, A. D. et al. Physicochemical mechanotransduction alters nuclear shape and mechanics via heterochromatin formation. Mol. Biol. Cell 30, 2320–2330 (2019). DOI: 10.1091/mbc.E19-05-0286
Cantwell, H. & Nurse, P. Unravelling nuclear size control. Curr. Genet. 65, 1281–1285 (2019). DOI: 10.1007/s00294-019-00999-3
Versaevel, M., Grevesse, T. & Gabriele, S. Spatial coordination between cell and nuclear shape within micropatterned endothelial cells. Nat. Commun. 3, 671 (2012). This work demonstrates that actomyosin stress fibres regulate nuclear deformations in response to cell shape changes and report a drastic condensation of chromatin in deformed nuclei. DOI: 10.1038/ncomms1668
Seirin-Lee, S. et al. Role of dynamic nuclear deformation on genomic architecture reorganization. PLoS Comput. Biol. 15, e1007289 (2019). DOI: 10.1371/journal.pcbi.1007289
Alisafaei, F., Jokhun, D. S., Shivashankar, G. V. & Shenoy, V. B. Regulation of nuclear architecture, mechanics, and nucleocytoplasmic shuttling of epigenetic factors by cell geometric constraints. Proc. Natl Acad. Sci. USA 116, 13200–13209 (2019). DOI: 10.1073/pnas.1902035116
Petrie, R. J., Koo, H. & Yamada, K. M. Generation of compartmentalized pressure by a nuclear piston governs cell motility in a 3D matrix. Science 345, 1062–1065 (2014). DOI: 10.1126/science.1256965
Mistriotis, P. et al. Confinement hinders motility by inducing RhoA-mediated nuclear influx, volume expansion, and blebbing. J. Cell Biol. 218, 4093–4111 (2019). DOI: 10.1083/jcb.201902057
Mitchison, T. J. Colloid osmotic parameterization and measurement of subcellular crowding. Mol. Biol. Cell 30, 173–180 (2019). DOI: 10.1091/mbc.E18-09-0549
Deviri, D. & Safran, S. A. Balance of osmotic pressures determines the volume of the cell nucleus. bioRxiv 10.1101/2021.10.01.462771v1 (2021). DOI: 10.1101/2021.10.01.462771v1
Lemière, J., Real-Calderon, P., Holt, L. J., Fai, T. G. & Chang, F. Control of nuclear size by osmotic forces in Schizosaccharomyces pombe. bioRxiv 10.1101/2021.12.05.471221 (2021). DOI: 10.1101/2021.12.05.471221
Takata, H. et al. Chromatin compaction protects genomic DNA from radiation damage. PLoS ONE 8, e75622 (2013). DOI: 10.1371/journal.pone.0075622
Holaska, J. M., Kowalski, A. K. & Wilson, K. L. Emerin caps the pointed end of actin filaments: evidence for an actin cortical network at the nuclear inner membrane. PLoS Biol. 2, e231 (2004). DOI: 10.1371/journal.pbio.0020231
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 study identifies how mechanical stretch can result in emerin translocation to the ONM, where it facilitates perinuclear actin polymerization that results in depletion of intranuclear actin and changes in chromatin organization. DOI: 10.1038/ncb3387
Davidson, P. M. & Cadot, B. Actin on and around the nucleus. Trends Cell Biol. 31, 211–223 (2020). DOI: 10.1016/j.tcb.2020.11.009
Buxboim, A. et al. Matrix elasticity regulates lamin-a,c phosphorylation and turnover with feedback to actomyosin. Curr. Biol. 24, 1909–1917 (2014). DOI: 10.1016/j.cub.2014.07.001
Mattout, A. et al. An EDMD mutation in C. elegans lamin blocks muscle-specific gene relocation and compromises muscle integrity. Curr. Biol. 21, 1603–1614 (2011). DOI: 10.1016/j.cub.2011.08.030
Solovei, I. et al. LBR and lamin A/C sequentially tether peripheral heterochromatin and inversely regulate differentiation. Cell 152, 584–598 (2013). DOI: 10.1016/j.cell.2013.01.009
Shin, J.-W. et al. Lamins regulate cell trafficking and lineage maturation of adult human hematopoietic cells. Proc. Natl Acad. Sci. USA 110, 18892–18897 (2013). DOI: 10.1073/pnas.1304996110
Swift, J. et al. Nuclear lamin-a scales with tissue stiffness and enhances matrix-directed differentiation. Science 341, 1240104 (2013). Detailed proteomic analysis linking higher levels of lamins A/C (and collagen) to cells residing in stiff tissues, suggesting a mechano-adaptive role of lamin A/C expression. DOI: 10.1126/science.1240104
Zuela, N., Dorfman, J. & Gruenbaum, Y. Global transcriptional changes caused by an EDMD mutation correlate to tissue specific disease phenotypes in C. elegans. Nucleus 8, 60–69 (2017). DOI: 10.1080/19491034.2016.1238999
Iyer, K. V. et al. Apico-basal cell compression regulates lamin A/C levels in epithelial tissues. Nat. Commun. 12, 1756 (2021). DOI: 10.1038/s41467-021-22010-9
Olins, D. E. & Olins, A. L. Granulocyte heterochromatin: defining the epigenome. BMC Cell Biol. 6, 39 (2005). DOI: 10.1186/1471-2121-6-39
Rowat, A. C. et al. Nuclear envelope composition determines the ability of neutrophil-type cells to passage through micron-scale constrictions. J. Biol. Chem. 288, 8610–8618 (2013). One of the first reports providing functional evidence that nuclear envelope composition and deformability determine the ability of cells to transit through tight constrictions. DOI: 10.1074/jbc.M112.441535
Roberts, A. B. et al. Tumor cell nuclei soften during transendothelial migration. J. Biomech. 121, 110400 (2021). DOI: 10.1016/j.jbiomech.2021.110400
Bell, E. S. et al. Low lamin A levels enhance confined cell migration and metastatic capacity in breast cancer. bioRxiv 10.1101/2021.07.12.451842v1 (2021). DOI: 10.1101/2021.07.12.451842v1
Infante, E. et al. LINC complex–Lis1 interplay controls MT1–MMP matrix digest-on-demand response for confined tumor cell migration. Nat. Commun. 9, 2443 (2018). DOI: 10.1038/s41467-018-04865-7
Krause, M. et al. Cell migration through three-dimensional confining pores: speed accelerations by deformation and recoil of the nucleus. Philos. Trans. R. Soc. Lond. B Biol. Sci. 374, 20180225 (2019). DOI: 10.1098/rstb.2018.0225
Gerlitz, G. The emerging roles of heterochromatin in cell migration. Front. Cell Dev. Biol. 8, 394 (2020). DOI: 10.3389/fcell.2020.00394
Earle, A. J. et al. Mutant lamins cause nuclear envelope rupture and DNA damage in skeletal muscle cells. Nat. Mater. 19, 464–473 (2020). DOI: 10.1038/s41563-019-0563-5
Mitchell, M. J. et al. Lamin A/C deficiency reduces circulating tumor cell resistance to fluid shear stress. Am. J. Physiol. Cell Physiol. 309, C736–C746 (2015). DOI: 10.1152/ajpcell.00050.2015
Gundersen, G. G. & Worman, H. J. Nuclear positioning. Cell 152, 1376–1389 (2013). DOI: 10.1016/j.cell.2013.02.031
Roman, W. & Gomes, E. R. Nuclear positioning in skeletal muscle. Semin. Cell Dev. Biol. 82, 51–56 (2018). DOI: 10.1016/j.semcdb.2017.11.005
Collins, M. A. et al. Ensconsin-dependent changes in microtubule organization and LINC complex-dependent changes in nucleus-nucleus interactions result in quantitatively distinct myonuclear positioning defects. Mol. Biol. Cell 32, ar27 (2021). DOI: 10.1091/mbc.E21-06-0324
Lorber, D., Rotkopf, R. & Volk, T. A minimal constraint device for imaging nuclei in live Drosophila contractile larval muscles reveals novel nuclear mechanical dynamics. Lab. Chip 20, 2100–2112 (2020). DOI: 10.1039/D0LC00214C
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). This report, along with work by Harada et al. (ref. 175) and Wolf et al. (ref. 126), presents some of the first evidence that increased nuclear deformability caused by reduced lamin A/C expression enhances cell migration through confined environments. DOI: 10.1007/s12195-014-0342-y
Wolf, K. et al. Physical limits of cell migration: control by ECM space and nuclear deformation and tuning by proteolysis and traction force. J. Cell Biol. 201, 1069–1084 (2013). This study demonstrates that nuclear deformability presents a rate-limiting factor in the ability of cells to migrate through constrictions smaller than ~10% of the undeformed nuclear cross-section. DOI: 10.1083/jcb.201210152
Venturini, V. et al. The nucleus measures shape changes for cellular proprioception to control dynamic cell behavior. Science 370, eaba2644 (2020). As presented in work by Lomakin et al. (ref. 17), it was shown that the nuclear envelope provides a gauge of cell deformation and activates a mechanotransduction pathway that controls actomyosin contractility via mechanically induced recruitment of cPLA2 to the INM. DOI: 10.1126/science.aba2644
Smith, E. R. et al. Nuclear envelope structural proteins facilitate nuclear shape changes accompanying embryonic differentiation and fidelity of gene expression. BMC Cell Biol. 18, 8 (2017). DOI: 10.1186/s12860-017-0125-0
Spear, P. C. & Erickson, C. A. Interkinetic nuclear migration: a mysterious process in search of a function. Dev. Growth Differ. 54, 306–316 (2012). DOI: 10.1111/j.1440-169X.2012.01342.x
Tsukamoto, S. et al. Compressive forces driven by lateral actin fibers are a key to the nuclear deformation under uniaxial cell-substrate stretching. Biochem. Biophys. Res. Commun. 597, 37–43 (2022). DOI: 10.1016/j.bbrc.2022.01.107
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). DOI: 10.1242/jcs.161703
Aureille, J. et al. Nuclear envelope deformation controls cell cycle progression in response to mechanical force. EMBO Rep. 20, e48084 (2019). DOI: 10.15252/embr.201948084
Lammerding, J. & Wolf, K. Nuclear envelope rupture: actin fibers are putting the squeeze on the nucleus. J. Cell Biol. 215, 5–8 (2016). DOI: 10.1083/jcb.201609102
Hatch, E. M. Nuclear envelope rupture: little holes, big openings. Curr. Opin. Cell Biol. 52, 66–72 (2018). DOI: 10.1016/j.ceb.2018.02.001
Khatau, S. B. et al. A perinuclear actin cap regulates nuclear shape. Proc. Natl Acad. Sci. USA 106, 19017–19022 (2009). DOI: 10.1073/pnas.0908686106
Lovett, D. B., Shekhar, N., Nickerson, J. A., Roux, K. J. & Lele, T. P. Modulation of nuclear shape by substrate rigidity. Cell. Mol. Bioeng. 6, 230–238 (2013). DOI: 10.1007/s12195-013-0270-2
Gupta, M. et al. Adaptive rheology and ordering of cell cytoskeleton govern matrix rigidity sensing. Nat. Commun. 6, 7525 (2015). DOI: 10.1038/ncomms8525
Nagayama, K., Yahiro, Y. & Matsumoto, T. Apical and basal stress fibers have different roles in mechanical regulation of the nucleus in smooth muscle cells cultured on a substrate. Cell. Mol. Bioeng. 6, 473–481 (2013). DOI: 10.1007/s12195-013-0294-7
Bruyère, C. et al. Actomyosin contractility scales with myoblast elongation and enhances differentiation through YAP nuclear export. Sci. Rep. 9, 15565 (2019). DOI: 10.1038/s41598-019-52129-1
Azevedo, M. & Baylies, M. K. Getting into position: nuclear movement in muscle cells. Trends Cell Biol. 30, 303–316 (2020). DOI: 10.1016/j.tcb.2020.01.002
Roman, W. et al. Muscle repair after physiological damage relies on nuclear migration for cellular reconstruction. Science 374, 355–359 (2021). DOI: 10.1126/science.abe5620
Gimpel, P. et al. Nesprin-1α-dependent microtubule nucleation from the nuclear envelope via Akap450 is necessary for nuclear positioning in muscle cells. Curr. Biol. 27, 2999–3009.e9 (2017). DOI: 10.1016/j.cub.2017.08.031
Roman, W. et al. Myofibril contraction and crosslinking drive nuclear movement to the periphery of skeletal muscle. Nat. Cell Biol. 19, 1189–1201 (2017). DOI: 10.1038/ncb3605
Wu, Y. K., Umeshima, H., Kurisu, J. & Kengaku, M. Nesprins and opposing microtubule motors generate a point force that drives directional nuclear motion in migrating neurons. Development 145, dev158782 (2018). DOI: 10.1242/dev.158782
Picariello, H. S. et al. Myosin IIA suppresses glioblastoma development in a mechanically sensitive manner. Proc. Natl Acad. Sci. USA 116, 15550–15559 (2019). DOI: 10.1073/pnas.1902847116
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). DOI: 10.4161/nucl.18954
Chai, R. J. et al. Disrupting the LINC complex by AAV mediated gene transduction prevents progression of lamin induced cardiomyopathy. Nat. Commun. 12, 4722 (2021). First report showing that LINC complex disruption can improve disease progression in a laminopathy mouse model of dilated cardiomyopathy. DOI: 10.1038/s41467-021-24849-4
Piccus, R. & Brayson, D. The nuclear envelope: LINCing tissue mechanics to genome regulation in cardiac and skeletal muscle. Biol. Lett. 16, 20200302 (2020). DOI: 10.1098/rsbl.2020.0302
Razafsky, D., Potter, C. & Hodzic, D. Validation of a mouse model to disrupt LINC complexes in a cell-specific manner. J. Vis. Exp. 106, e53318 (2015).
Hampoelz, B. et al. Microtubule-induced nuclear envelope fluctuations control chromatin dynamics in Drosophila embryos. Development 138, 3377–3386 (2011). DOI: 10.1242/dev.065706
Schulze, S. R. et al. A comparative study of Drosophila and human A-type lamins. PLoS ONE 4, e7564 (2009). DOI: 10.1371/journal.pone.0007564
Bone, C. R., Chang, Y.-T., Cain, N. E., Murphy, S. P. & Starr, D. A. Nuclei migrate through constricted spaces using microtubule motors and actin networks in C. elegans hypodermal cells. Development 143, 4193–4202 (2016).
Driver, E. C., Northrop, A. & Kelley, M. W. Cell migration, intercalation and growth regulate mammalian cochlear extension. Development 144, 3766–3776 (2017).
Mohammed, D. et al. Substrate area confinement is a key determinant of cell velocity in collective migration. Nat. Phys. 15, 858–866 (2019). DOI: 10.1038/s41567-019-0543-3
Yanakieva, I., Erzberger, A., Matejčić, M., Modes, C. D. & Norden, C. Cell and tissue morphology determine actin-dependent nuclear migration mechanisms in neuroepithelia. J. Cell Biol. 218, 3272–3289 (2019). DOI: 10.1083/jcb.201901077
Norden, C., Young, S., Link, B. A. & Harris, W. A. Actomyosin is the main driver of interkinetic nuclear migration in the retina. Cell 138, 1195–1208 (2009). DOI: 10.1016/j.cell.2009.06.032
Tsai, L.-H. & Gleeson, J. G. Nucleokinesis in neuronal migration. Neuron 46, 383–388 (2005). DOI: 10.1016/j.neuron.2005.04.013
Cooper, J. A. Mechanisms of cell migration in the nervous system. J. Cell Biol. 202, 725–734 (2013). DOI: 10.1083/jcb.201305021
Young, S. G., Jung, H.-J., Lee, J. M. & Fong, L. G. Nuclear lamins and neurobiology. Mol. Cell Biol. 34, 2776–2785 (2014). DOI: 10.1128/MCB.00486-14
Wolf, K. et al. Collagen-based cell migration models in vitro and in vivo. Semin. Cell Dev. Biol. 20, 931–941 (2009). DOI: 10.1016/j.semcdb.2009.08.005
Yamada, K. M. & Sixt, M. Mechanisms of 3D cell migration. Nat. Rev. Mol. Cell Biol. 20, 738–752 (2019). DOI: 10.1038/s41580-019-0172-9
Renkawitz, J. et al. Nuclear positioning facilitates amoeboid migration along the path of least resistance. Nature 568, 546–550 (2019). DOI: 10.1038/s41586-019-1087-5
Maciejowski, J. & Hatch, E. M. Nuclear membrane rupture and its consequences. Annu. Rev. Cell Dev. Biol. 36, 85–114 (2020). DOI: 10.1146/annurev-cellbio-020520-120627
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). DOI: 10.1038/ncomms10997
Fridolfsson, H. N. & Starr, D. A. Kinesin-1 and dynein at the nuclear envelope mediate the bidirectional migrations of nuclei. J. Cell Biol. 191, 115–128 (2010). DOI: 10.1083/jcb.201004118
Marks, P. C. & Petrie, R. J. Push or pull: how cytoskeletal crosstalk facilitates nuclear movement through 3D environments. Phys. Biol. 19, 021003 (2022). DOI: 10.1088/1478-3975/ac45e3
de Noronha, C. M. C. et al. Dynamic disruptions in nuclear envelope architecture and integrity induced by HIV-1 Vpr. Science 294, 1105–1108 (2001). DOI: 10.1126/science.1063957
Vos, W. H. D. et al. Repetitive disruptions of the nuclear envelope invoke temporary loss of cellular compartmentalization in laminopathies. Hum. Mol. Genet. 20, 4175–4186 (2011). First report of spontaneous nuclear envelope rupture in laminopathy cells. DOI: 10.1093/hmg/ddr344
Srivastava, N. et al. Nuclear fragility, blaming the blebs. Curr. Opin. Cell Biol. 70, 100–108 (2021). DOI: 10.1016/j.ceb.2021.01.007
Pfeifer, C. R. et al. Gaussian curvature dilutes the nuclear lamina, favoring nuclear rupture, especially at high strain rate. Nucleus 13, 129–143 (2022). DOI: 10.1080/19491034.2022.2045726
Pfeifer, C. R., Vashisth, M., Xia, Y. & Discher, D. E. Nuclear failure, DNA damage, and cell cycle disruption after migration through small pores: a brief review. Essays Biochem. 63, 569–577 (2019). DOI: 10.1042/EBC20190007
Goldman, R. D. et al. Accumulation of mutant lamin A causes progressive changes in nuclear architecture in Hutchinson-Gilford progeria syndrome. Proc. Natl Acad. Sci. USA 101, 8963–8968 (2004). DOI: 10.1073/pnas.0402943101
Muchir, A. et al. Nuclear envelope alterations in fibroblasts from patients with muscular dystrophy, cardiomyopathy, and partial lipodystrophy carrying lamin A/C gene mutations. Muscle Nerve 30, 444–450 (2004). DOI: 10.1002/mus.20122
Karoutas, A. et al. The NSL complex maintains nuclear architecture stability via lamin A/C acetylation. Nat. Cell Biol. 21, 1248–1260 (2019). DOI: 10.1038/s41556-019-0397-z
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). Study demonstrating that increased nuclear deformability caused by reduced lamin A/C expression enhances cell migration through confined environments but renders cells more susceptible to mechanically induced damage and cell death. DOI: 10.1083/jcb.201308029
Rowat, A. C., Lammerding, J. & Ipsen, J. H. Mechanical properties of the cell nucleus and the effect of emerin deficiency. Biophys. J. 91, 4649–4664 (2006). DOI: 10.1529/biophysj.106.086454
Berre, M. L., 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). DOI: 10.1039/c2ib20056b
Takaki, T. et al. Actomyosin drives cancer cell nuclear dysmorphia and threatens genome stability. Nat. Commun. 8, 16013 (2017). DOI: 10.1038/ncomms16013
Jain, N. & Vogel, V. Spatial confinement downsizes the inflammatory response of macrophages. Nat. Mater. 17, 1134–1144 (2018). DOI: 10.1038/s41563-018-0190-6
Isermann, P. & Lammerding, J. Consequences of a tight squeeze: nuclear envelope rupture and repair. Nucleus 8, 268–274 (2017). DOI: 10.1080/19491034.2017.1292191
Deviri, D. et al. Scaling laws indicate distinct nucleation mechanisms of holes in the nuclear lamina. Nat. Phys. 15, 823–829 (2019). DOI: 10.1038/s41567-019-0506-8
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). This study reports increasing DNA damage and chromosomal abnormalities in tumour cells after repeated migration through small constrictions. DOI: 10.1016/j.cub.2016.11.049
Halfmann, C. T. et al. Repair of nuclear ruptures requires barrier-to-autointegration factor. J. Cell Biol. 218, 2136–2149 (2019). DOI: 10.1083/jcb.201901116
Penfield, L. et al. Dynein pulling forces counteract lamin-mediated nuclear stability during nuclear envelope repair. Mol. Biol. Cell 29, 852–868 (2018). DOI: 10.1091/mbc.E17-06-0374
Young, A. M., Gunn, A. L. & Hatch, E. M. BAF facilitates interphase nuclear membrane repair through recruitment of nuclear transmembrane proteins. Mol. Biol. Cell 31, 1551–1560 (2020). DOI: 10.1091/mbc.E20-01-0009
Nader, G. P. et al. Compromised nuclear envelope integrity drives TREX1-dependent DNA damage and tumor cell invasion. Cell 184, 5230–5246 (2021). DOI: 10.1016/j.cell.2021.08.035
Shah, P. et al. Nuclear deformation causes DNA damage by increasing replication stress. Curr. Biol. 31, 753–765 (2021). DOI: 10.1016/j.cub.2020.11.037
Kidiyoor, G. R. et al. ATR is essential for preservation of cell mechanics and nuclear integrity during interstitial migration. Nat. Commun. 11, 4828 (2020). DOI: 10.1038/s41467-020-18580-9
Jiang, Y. N. et al. Interleukin 6-triggered ataxia-telangiectasia mutated kinase activation facilitates epithelial-to-mesenchymal transition in lung cancer by upregulating vimentin expression. Exp. Cell Res. 381, 165–171 (2019). DOI: 10.1016/j.yexcr.2019.05.011
Peng, B., Ortega, J., Gu, L., Chang, Z. & Li, G.-M. Phosphorylation of proliferating cell nuclear antigen promotes cancer progression by activating the ATM/Akt/GSK3β/Snail signaling pathway. J. Biol. Chem. 294, 7037–7045 (2019). DOI: 10.1074/jbc.RA119.007897
Bakhoum, S. F. et al. Chromosomal instability drives metastasis through a cytosolic DNA response. Nature 553, 467–472 (2018). DOI: 10.1038/nature25432
Enyedi, B., Jelcic, M. & Niethammer, P. The cell nucleus serves as a mechanotransducer of tissue damage-induced inflammation. Cell 165, 1160–1170 (2016). First demonstration that increased nuclear membrane tension can trigger recruitment of cPLA2 to the, where it can induce further downstream signalling. DOI: 10.1016/j.cell.2016.04.016
Shen, Z. et al. A synergy between mechanosensitive calcium- and membrane-binding mediates tension-sensing by C2-like domains. Proc. Natl Acad. Sci. USA 119, e2112390119 (2022).
Niethammer, P. Components and mechanisms of nuclear mechanotransduction. Annu. Rev. Cell Dev. Biol. 37, 233–256 (2021). DOI: 10.1146/annurev-cellbio-120319-030049
Shen, Z. & Niethammer, P. A cellular sense of space and pressure. Science 370, 295–296 (2020). DOI: 10.1126/science.abe3881
Katayama, T. et al. Stimulatory effects of arachidonic acid on myosin ATPase activity and contraction of smooth muscle via myosin motor domain. Am. J. Physiol. Heart Circ. Physiol. 298, H505–H514 (2010). DOI: 10.1152/ajpheart.00577.2009
Brown, M., Roulson, J.-A., Hart, C. A., Tawadros, T. & Clarke, N. W. Arachidonic acid induction of Rho-mediated transendothelial migration in prostate cancer. Br. J. Cancer 110, 2099–2108 (2014). DOI: 10.1038/bjc.2014.99
Elosegui-Artola, A. et al. Force triggers YAP nuclear entry by regulating transport across nuclear pores. Cell 171, 1397–1410 (2017). First report of mechanically induced opening of NPCs, mediating import of the mechanoresponsive transcription factor YAP. DOI: 10.1016/j.cell.2017.10.008
Zimmerli, C. E. et al. Nuclear pores dilate and constrict in cellulo. Science 374, eabd9776 (2021). DOI: 10.1126/science.abd9776
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). DOI: 10.1016/j.bpj.2015.05.010
Moya, I. M. & Halder, G. Hippo-YAP/TAZ signalling in organ regeneration and regenerative medicine. Nat. Rev. Mol. Cell Biol. 20, 211–226 (2019). DOI: 10.1038/s41580-018-0086-y
Luciano, M. et al. Cell monolayers sense curvature by exploiting active mechanics and nuclear mechanoadaptation. Nat. Phys. 17, 1382–1390 (2021). DOI: 10.1038/s41567-021-01374-1
Aragona, M. et al. A mechanical checkpoint controls multicellular growth through YAP/TAZ regulation by actin-processing factors. Cell 154, 1047–1059 (2013). DOI: 10.1016/j.cell.2013.07.042
Tajik, A. et al. Transcription upregulation via force-induced direct stretching of chromatin. Nat. Mater. 15, 1287–1296 (2016). First report of mechanically induced chromosome stretching and increased gene expression. DOI: 10.1038/nmat4729
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). DOI: 10.1126/sciadv.aay9095
Almonacid, M. et al. Active fluctuations of the nuclear envelope shape the transcriptional dynamics in oocytes. Dev. Cell 51, 145–157.e10 (2019). DOI: 10.1016/j.devcel.2019.09.010
Hsia, C.-R. et al. Confined migration induces heterochromatin formation and alters chromatin accessibility. bioRxiv 10.1101/2021.09.22.461293 (2021). DOI: 10.1101/2021.09.22.461293
Jacobson, E. C. et al. Migration through a small pore disrupts inactive chromatin organization in neutrophil-like cells. BMC Biol. 16, 142 (2018). DOI: 10.1186/s12915-018-0608-2
Golloshi, R. et al. Constricted migration is associated with stable 3D genome structure differences in melanoma cells. bioRxiv 10.1101/856583 (2020). DOI: 10.1101/856583
Damodaran, K. et al. Compressive force induces reversible chromatin condensation and cell geometry-dependent transcriptional response. Mol. Biol. Cell 29, 3039–3051 (2018). DOI: 10.1091/mbc.E18-04-0256
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). DOI: 10.1038/nature12105
Killaars, A. R., Walker, C. J. & Anseth, K. S. Nuclear mechanosensing controls MSC osteogenic potential through HDAC epigenetic remodeling. Proc. Natl Acad. Sci. USA 117, 21258–21266 (2020). DOI: 10.1073/pnas.2006765117
Walker, C. J. et al. Nuclear mechanosensing drives chromatin remodelling in persistently activated fibroblasts. Nat. Biomed. Eng. 5, 1485–1499 (2021). DOI: 10.1038/s41551-021-00709-w
Seelbinder, B. et al. Nuclear deformation guides chromatin reorganization in cardiac development and disease. Nat. Biomed. Eng. 5, 1500–1516 (2021). DOI: 10.1038/s41551-021-00823-9
Heo, S.-J. et al. Mechanically induced chromatin condensation requires cellular contractility in mesenchymal stem cells. Biophys. J. 111, 864–874 (2016). DOI: 10.1016/j.bpj.2016.07.006
Heo, S.-J. et al. Biophysical regulation of chromatin architecture instills a mechanical memory in mesenchymal stem cells. Sci. Rep. 5, 16895 (2015). DOI: 10.1038/srep16895
Hannezo, E. & Heisenberg, C.-P. Mechanochemical feedback loops in development and disease. Cell 178, 12–25 (2019). DOI: 10.1016/j.cell.2019.05.052
Kirby, T. J. & Lammerding, J. Emerging views of the nucleus as a cellular mechanosensor. Nat. Cell Biol. 20, 373–381 (2018). DOI: 10.1038/s41556-018-0038-y
Miroshnikova, Y. A. & Wickström, S. A. Mechanical forces in nuclear organization. Cold Spring Harb. Perspect. Biol. 14, a039685 (2022). DOI: 10.1101/cshperspect.a039685
Swift, J. & Discher, D. E. The nuclear lamina is mechano-responsive to ECM elasticity in mature tissue. J. Cell Sci. 127, 3005–3015 (2014).
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). DOI: 10.1038/nmat4389
Sapra, K. T. et al. Nonlinear mechanics of lamin filaments and the meshwork topology build an emergent nuclear lamina. Nat. Commun. 11, 6205 (2020). DOI: 10.1038/s41467-020-20049-8
Guilluy, C. et al. Isolated nuclei adapt to force and reveal a mechanotransduction pathway in the nucleus. Nat. Cell Biol. 16, 376–381 (2014). First report of mechano-adaptive changes in isolated nuclei, indicating a nucleus-intrinsic ability to respond to mechanical forces. DOI: 10.1038/ncb2927
Zwerger, M., Ho, C. Y. & Lammerding, J. Nuclear mechanics in disease. Annu. Rev. Biomed. Eng. 13, 397–428 (2011). DOI: 10.1146/annurev-bioeng-071910-124736
Nyirenda, N., Farkas, D. L. & Ramanujan, V. K. Preclinical evaluation of nuclear morphometry and tissue topology for breast carcinoma detection and margin assessment. Breast Cancer Res. Treat. 126, 345–354 (2011). DOI: 10.1007/s10549-010-0914-z
Mueller, J. L. et al. Rapid staining and imaging of subnuclear features to differentiate between malignant and benign breast tissues at a point-of-care setting. J. Cancer Res. Clin. Oncol. 142, 1475–1486 (2016). DOI: 10.1007/s00432-016-2165-9
Somech, R., Shaklai, S., Amariglio, N., Rechavi, G. & Simon, A. J. Nuclear envelopathies — raising the nuclear veil. Pediatr. Res. 57, 8–15 (2005). DOI: 10.1203/01.PDR.0000159566.54287.6C
Hershberger, R. E., Hedges, D. J. & Morales, A. Dilated cardiomyopathy: the complexity of a diverse genetic architecture. Nat. Rev. Cardiol. 10, 531–547 (2013). DOI: 10.1038/nrcardio.2013.105
Wong, X. & Stewart, C. L. The laminopathies and the insights they provide into the structural and functional organization of the nucleus. Annu. Rev. Genomics Hum. Genet. 21, 263–288 (2020). DOI: 10.1146/annurev-genom-121219-083616
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). DOI: 10.1038/6799
Sandre-Giovannoli, A. D. et al. Lamin a truncation in Hutchinson-Gilford progeria. Science 300, 2055 (2003). DOI: 10.1126/science.1084125
Folker, E. S., Ostlund, C., Luxton, G. W. G., 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). DOI: 10.1073/pnas.1000824108
Méjat, A. & Misteli, T. LINC complexes in health and disease. Nucleus 1, 40–52 (2010). DOI: 10.4161/nucl.1.1.10530
Fischer, M., Rikeit, P., Knaus, P. & Coirault, C. YAP-mediated mechanotransduction in skeletal muscle. Front. Physiol. 7, 41 (2016). DOI: 10.3389/fphys.2016.00041
Owens, D. J. et al. Lamin mutations cause increased YAP nuclear entry in muscle stem cells. Cells 9, 816 (2020). DOI: 10.3390/cells9040816
Eriksson, M. et al. Recurrent de novo point mutations in lamin A cause Hutchinson–Gilford progeria syndrome. Nature 423, 293–298 (2003). DOI: 10.1038/nature01629
Verstraeten, V. L. R. M., Ji, J. Y., Cummings, K. S., Lee, R. T. & Lammerding, J. Increased mechanosensitivity and nuclear stiffness in Hutchinson-Gilford progeria cells: effects of farnesyltransferase inhibitors. Aging Cell 7, 383–393 (2008). DOI: 10.1111/j.1474-9726.2008.00382.x
Booth, E. A., Spagnol, S. T., Alcoser, T. A. & Dahl, K. N. Nuclear stiffening and chromatin softening with progerin expression leads to an attenuated nuclear response to force. Soft Matter 11, 6412–6418 (2015). DOI: 10.1039/C5SM00521C
Kim, P. H. et al. Disrupting the LINC complex in smooth muscle cells reduces aortic disease in a mouse model of Hutchinson-Gilford progeria syndrome. Sci. Transl. Med. 10, eaat7163 (2018). DOI: 10.1126/scitranslmed.aat7163
Dahl, K. N. et al. Distinct structural and mechanical properties of the nuclear lamina in Hutchinson-Gilford progeria syndrome. Proc. Natl Acad. Sci. USA 103, 10271–10276 (2006). DOI: 10.1073/pnas.0601058103
Columbaro, M. et al. Rescue of heterochromatin organization in Hutchinson-Gilford progeria by drug treatment. Cell. Mol. Life Sci. 62, 2669–2678 (2005). DOI: 10.1007/s00018-005-5318-6
Coffinier, C. et al. Deficiencies in lamin B1 and lamin B2 cause neurodevelopmental defects and distinct nuclear shape abnormalities in neurons. Mol. Biol. Cell 22, 4683–4693 (2011). DOI: 10.1091/mbc.e11-06-0504
Young, S. G., Jung, H.-J., Coffinier, C. & Fong, L. G. Understanding the roles of nuclear A- and B-type lamins in brain development. J. Biol. Chem. 287, 16103–16110 (2012). DOI: 10.1074/jbc.R112.354407
Coffinier, C., Fong, L. G. & Young, S. G. LINCing lamin B2 to neuronal migration: growing evidence for cell-specific roles of B-type lamins. Nucleus 1, 407–411 (2010). DOI: 10.4161/nucl.1.5.12830
Vortmeyer-Krause, M. et al. Lamin B2 follows lamin A/C- mediated nuclear mechanics and cancer cell invasion efficacy. bioRxiv 10.1101/2020.04.07.028969 (2020). DOI: 10.1101/2020.04.07.028969
Padiath, Q. S. et al. Lamin B1 duplications cause autosomal dominant leukodystrophy. Nat. Genet. 38, 1114–1123 (2006). DOI: 10.1038/ng1872
Ballatore, C., Lee, V. M.-Y. & Trojanowski, J. Q. Tau-mediated neurodegeneration in Alzheimer’s disease and related disorders. Nat. Rev. Neurosci. 8, 663–672 (2007). DOI: 10.1038/nrn2194
Sergent, C., Baillet, S. & Dehaene, S. Timing of the brain events underlying access to consciousness during the attentional blink. Nat. Neurosci. 8, 1391–1400 (2005). DOI: 10.1038/nn1549
Fernández-Nogales, M. et al. Huntington’s disease is a four-repeat tauopathy with tau nuclear rods. Nat. Med. 20, 881–885 (2014). DOI: 10.1038/nm.3617
Crisp, M. et al. Coupling of the nucleus and cytoplasm: role of the LINC complex. J. Cell Biol. 172, 41–53 (2006). First description of the LINC complex and its role in connecting the cytoskeleton and nuclear interior. DOI: 10.1083/jcb.200509124
Paonessa, F. et al. Microtubules deform the nuclear membrane and disrupt nucleocytoplasmic transport in tau-mediated frontotemporal dementia. Cell Rep. 26, 582–593.e5 (2019). DOI: 10.1016/j.celrep.2018.12.085
Fernández-Nogales, M. & Lucas, J. J. Altered levels and isoforms of tau and nuclear membrane invaginations in Huntington’s disease. Front Cell. Neurosci. 13, 574 (2020). DOI: 10.3389/fncel.2019.00574
von Appen, A. et al. LEM2 phase separation promotes ESCRT-mediated nuclear envelope reformation. Nature 582, 115–118 (2020). DOI: 10.1038/s41586-020-2232-x
Stephens, A. D., Banigan, E. J. & Marko, J. F. Separate roles for chromatin and lamins in nuclear mechanics. Nucleus 9, 119–124 (2018). DOI: 10.1080/19491034.2017.1414118
Bronshtein, I. et al. Loss of lamin A function increases chromatin dynamics in the nuclear interior. Nat. Commun. 6, 8044 (2015). DOI: 10.1038/ncomms9044
Corne, T. D. J. et al. Deregulation of focal adhesion formation and cytoskeletal tension due to loss of A-type lamins. Cell Adhes. Migr. 11, 447–463 (2017). DOI: 10.1080/19336918.2016.1247144
Nikolova, V. et al. Defects in nuclear structure and function promote dilated cardiomyopathy in lamin A/C-deficient mice. J. Clin. Invest. 113, 357–369 (2004). DOI: 10.1172/JCI200419448
Puckelwartz, M. J. et al. Nesprin-1 mutations in human and murine cardiomyopathy. J. Mol. Cell. Cardiol. 48, 600–608 (2010). DOI: 10.1016/j.yjmcc.2009.11.006