Adherens junctions as molecular regulators of emergent tissue mechanics.
Journal
Nature reviews. Molecular cell biology
ISSN: 1471-0080
Titre abrégé: Nat Rev Mol Cell Biol
Pays: England
ID NLM: 100962782
Informations de publication
Date de publication:
13 Dec 2023
13 Dec 2023
Historique:
accepted:
08
11
2023
medline:
14
12
2023
pubmed:
14
12
2023
entrez:
13
12
2023
Statut:
aheadofprint
Résumé
Tissue and organ development during embryogenesis relies on the collective and coordinated action of many cells. Recent studies have revealed that tissue material properties, including transitions between fluid and solid tissue states, are controlled in space and time to shape embryonic structures and regulate cell behaviours. Although the collective cellular flows that sculpt tissues are guided by tissue-level physical changes, these ultimately emerge from cellular-level and subcellular-level molecular mechanisms. Adherens junctions are key subcellular structures, built from clusters of classical cadherin receptors. They mediate physical interactions between cells and connect biochemical signalling to the physical characteristics of cell contacts, hence playing a fundamental role in tissue morphogenesis. In this Review, we take advantage of the results of recent, quantitative measurements of tissue mechanics to relate the molecular and cellular characteristics of adherens junctions, including adhesion strength, tension and dynamics, to the emergent physical state of embryonic tissues. We focus on systems in which cell-cell interactions are the primary contributor to morphogenesis, without significant contribution from cell-matrix interactions. We suggest that emergent tissue mechanics is an important direction for future research, bridging cell biology, developmental biology and mechanobiology to provide a holistic understanding of morphogenesis in health and disease.
Identifiants
pubmed: 38093099
doi: 10.1038/s41580-023-00688-7
pii: 10.1038/s41580-023-00688-7
doi:
Types de publication
Journal Article
Review
Langues
eng
Sous-ensembles de citation
IM
Informations de copyright
© 2023. Springer Nature Limited.
Références
Tepass, U., Troung, K., Godt, D., Ikura, M. & Peifer, M. Cadherins in embyronic and neural morphogenesis. Nat. Rev. Mol. Cell Biol. 1, 91–100 (2000).
pubmed: 11253370
doi: 10.1038/35040042
Mongera, A. et al. A fluid-to-solid jamming transition underlies vertebrate body axis elongation. Nature 561, 401–405 (2018).
pubmed: 30185907
doi: 10.1038/s41586-018-0479-2
pmcid: 6148385
Lenne, P. F. & Trivedi, V. Sculpting tissues by phase transitions. Nat. Commun. 13, 664 (2022).
pubmed: 35115507
doi: 10.1038/s41467-022-28151-9
pmcid: 8814027
Jain, A. et al. Regionalized tissue fluidization is required for epithelial gap closure during insect gastrulation. Nat. Commun. 11, 5604 (2020).
pubmed: 33154375
doi: 10.1038/s41467-020-19356-x
pmcid: 7645651
Tetley, R. J. et al. Tissue fluidity promotes epithelial wound healing. Nat. Phys. 15, 1195–1203 (2019).
pubmed: 31700525
doi: 10.1038/s41567-019-0618-1
pmcid: 6837871
Petridou, N. I., Grigolon, S., Salbreux, G., Hannezo, E. & Heisenberg, C. P. Fluidization-mediated tissue spreading by mitotic cell rounding and non-canonical Wnt signalling. Nat. Cell Biol. 21, 169–178 (2019).
pubmed: 30559456
doi: 10.1038/s41556-018-0247-4
Wang, X. et al. Anisotropy links cell shapes to tissue flow during convergent extension. Proc. Natl Acad. Sci. USA 117, 13541–13551 (2020).
pubmed: 32467168
doi: 10.1073/pnas.1916418117
pmcid: 7306759
Atia, L. et al. Geometric constraints during epithelial jamming. Nat. Phys. 14, 629–629 (2018).
Founounou, N. et al. Tissue fluidity mediated by adherens junction dynamics promotes planar cell polarity-driven ommatidial rotation. Nat. Commun. 12, 6974 (2021).
pubmed: 34848713
doi: 10.1038/s41467-021-27253-0
pmcid: 8632910
Palamidessi, A. et al. Unjamming overcomes kinetic and proliferation arrest in terminally differentiated cells and promotes collective motility of carcinoma. Nat. Mater. 18, 1252–1263 (2019).
pubmed: 31332337
doi: 10.1038/s41563-019-0425-1
Frittoli, E. et al. Tissue fluidification promotes a cGAS-STING cytosolic DNA response in invasive breast cancer. Nat. Mater. 22, 644–655 (2023).
pubmed: 36581770
doi: 10.1038/s41563-022-01431-x
Kang, W. et al. A novel jamming phase diagram links tumor invasion to non-equilibrium phase separation. iScience 24, 103252 (2021).
pubmed: 34755092
doi: 10.1016/j.isci.2021.103252
pmcid: 8564056
Grosser, S. et al. Cell and nucleus shape as an indicator of tissue fluidity in carcinoma. Phys. Rev. X 11, 011033 (2021).
Campas, O. A toolbox to explore the mechanics of living embryonic tissues. Semin. Cell Dev. Biol. 55, 119–130 (2016).
pubmed: 27061360
doi: 10.1016/j.semcdb.2016.03.011
pmcid: 4903887
Gomez-Gonzalez, M., Latorre, E., Arroyo, M. & Trepat, X. Measuring mechanical stress in living tissues. Nat. Rev. Phys. 2, 300–317 (2020).
doi: 10.1038/s42254-020-0184-6
Sugimura, K., Lenne, P. F. & Graner, F. Measuring forces and stresses in situ in living tissues. Development 143, 186–196 (2016).
pubmed: 26786209
doi: 10.1242/dev.119776
Acharya, B. R. et al. A mechanosensitive RhoA pathway that protects epithelia against acute tensile stress. Dev. Cell 47, 439–452.e6 (2018).
pubmed: 30318244
doi: 10.1016/j.devcel.2018.09.016
Prakash, V., Bull, M. S. & Prakash, M. Motility-induced fracture reveals a ductile-to-brittle crossover in a simple animal’s epithelia. Nat. Phys. 17, 504–511 (2021).
doi: 10.1038/s41567-020-01134-7
Sherwood, D. R. Basement membrane remodeling guides cell migration and cell morphogenesis during development. Curr. Opin. Cell Biol. 72, 19–27 (2021).
pubmed: 34015751
doi: 10.1016/j.ceb.2021.04.003
pmcid: 8530833
Walma, D. A. C. & Yamada, K. M. The extracellular matrix in development. Development 147, dev175596 (2020).
pubmed: 32467294
doi: 10.1242/dev.175596
pmcid: 7272360
Wu, D., Yamada, K. M. & Wang, S. Tissue morphogenesis through dynamic cell and matrix interactions. Annu. Rev. Cell Dev. Biol. 39, 123–144 (2023).
pubmed: 37315160
doi: 10.1146/annurev-cellbio-020223-031019
Bonnans, C., Chou, J. & Werb, Z. Remodelling the extracellular matrix in development and disease. Nat. Rev. Mol. Cell Biol. 15, 786–801 (2014).
pubmed: 25415508
doi: 10.1038/nrm3904
pmcid: 4316204
Nelson, C. M. & Bissell, M. J. Of extracellular matrix, scaffolds, and signaling: tissue architecture regulates development, homeostasis, and cancer. Annu. Rev. Cell Dev. Biol. 22, 287–309 (2006).
pubmed: 16824016
doi: 10.1146/annurev.cellbio.22.010305.104315
pmcid: 2933192
Rozario, T. & DeSimone, D. W. The extracellular matrix in development and morphogenesis: a dynamic view. Dev. Biol. 341, 126–140 (2010).
pubmed: 19854168
doi: 10.1016/j.ydbio.2009.10.026
Farquhar, M. G. & Palade, G. E. Junctional complexes in various epithelia. J. Cell Biol. 17, 375–412 (1963).
pubmed: 13944428
doi: 10.1083/jcb.17.2.375
pmcid: 2106201
Boller, K., Vestweber, D. & Kemler, R. Cell-adhesion molecule uvomorulin is localized in the intermediate junctions of adult intestinal epithelial cells. J. Cell Biol. 100, 327–332 (1985).
pubmed: 3880756
doi: 10.1083/jcb.100.1.327
Strale, P. O. et al. The formation of ordered nanoclusters controls cadherin anchoring to actin and cell-cell contact fluidity. J. Cell Biol. 210, 333–346 (2015).
pubmed: 26195669
doi: 10.1083/jcb.201410111
pmcid: 4508897
Tepass, U. & Hartenstein, V. The development of cellular junctions in the Drosophila embryo. Dev. Biol. 161, 563–596 (1994).
pubmed: 8314002
doi: 10.1006/dbio.1994.1054
Huebner, R. J. et al. Mechanical heterogeneity along single cell-cell junctions is driven by lateral clustering of cadherins during vertebrate axis elongation. eLife 10, e65390 (2021).
pubmed: 34032216
doi: 10.7554/eLife.65390
pmcid: 8205493
Mege, R. M. & Ishiyama, N. Integration of cadherin adhesion and cytoskeleton at adherens junctions. Cold Spring Harb. Perspect. Biol. 9, a028738 (2017).
pubmed: 28096263
doi: 10.1101/cshperspect.a028738
pmcid: 5411698
Ozawa, M. & Kemler, R. Molecular organization of the uvomorulin-catenin complex. J. Cell Biol. 116, 989–996 (1992).
pubmed: 1734027
doi: 10.1083/jcb.116.4.989
Priest, A. V., Shafraz, O. & Sivasankar, S. Biophysical basis of cadherin mediated cell-cell adhesion. Exp. Cell Res. 358, 10–13 (2017).
pubmed: 28300566
doi: 10.1016/j.yexcr.2017.03.015
Rubsam, M. et al. Adherens junctions and desmosomes coordinate mechanics and signaling to orchestrate tissue morphogenesis and function: an evolutionary perspective. Cold Spring Harb. Perspect. Biol. 10, a029207 (2018).
pubmed: 28893859
doi: 10.1101/cshperspect.a029207
pmcid: 6211388
Noordstra, I., Morris, R. G. & Yap, A. S. Cadherins and the cortex: a matter of time? Curr. Opin. Cell Biol. 80, 102154 (2023).
pubmed: 36822056
doi: 10.1016/j.ceb.2023.102154
Charras, G. & Yap, A. S. Tensile forces and mechanotransduction at cell-cell junctions. Curr. Biol. 28, R445–R457 (2018).
pubmed: 29689229
doi: 10.1016/j.cub.2018.02.003
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).
pubmed: 17643125
doi: 10.1038/nrm2222
Ratheesh, A. et al. Centralspindlin and alpha-catenin regulate Rho signalling at the epithelial zonula adherens. Nat. Cell Biol. 14, 818–828 (2012).
pubmed: 22750944
doi: 10.1038/ncb2532
pmcid: 3939354
Noren, N. K., Liu, B. P., Burridge, K. & Kreft, B. p120 catenin regulates the actin cytoskeleton via rho family GTPases. J. Cell Biol. 150, 567–579 (2000).
pubmed: 10931868
doi: 10.1083/jcb.150.3.567
pmcid: 2175185
Noren, N. K., Niessen, C. M., Gumbiner, B. M. & Burridge, K. Cadherin engagement regulates Rho family GTPases. J. Biol. Chem. 276, 33305–33308 (2001).
pubmed: 11457821
doi: 10.1074/jbc.C100306200
Leckband, D. E. & de Rooij, J. Cadherin adhesion and mechanotransduction. Annu. Rev. Cell Dev. Biol. 30, 291–315 (2014).
pubmed: 25062360
doi: 10.1146/annurev-cellbio-100913-013212
Yap, A. S., Duszyc, K. & Viasnoff, V. Mechanosensing and mechanotransduction at cell-cell junctions. Cold Spring Harb. Perspect. Biol. 10, a028761 (2017).
doi: 10.1101/cshperspect.a028761
Manibog, K., Li, H., Rakshit, S. & Sivasankar, S. Resolving the molecular mechanism of cadherin catch bond formation. Nat. Commun. 5, 3941 (2014).
pubmed: 24887573
doi: 10.1038/ncomms4941
Rakshit, S., Zhang, Y., Manibog, K., Shafraz, O. & Sivasankar, S. Ideal, catch, and slip bonds in cadherin adhesion. Proc. Natl Acad. Sci. USA 109, 18815–18820 (2012).
pubmed: 23112161
doi: 10.1073/pnas.1208349109
pmcid: 3503169
Buckley, C. D. et al. The minimal cadherin-catenin complex binds to actin filaments under force. Science 346, 1254211 (2014).
pubmed: 25359979
doi: 10.1126/science.1254211
pmcid: 4364042
Ishiyama, N. et al. Force-dependent allostery of the alpha-catenin actin-binding domain controls adherens junction dynamics and functions. Nat. Commun. 9, 5121 (2018).
pubmed: 30504777
doi: 10.1038/s41467-018-07481-7
pmcid: 6269467
Huang, D. L., Bax, N. A., Buckley, C. D., Weis, W. I. & Dunn, A. R. Vinculin forms a directionally asymmetric catch bond with F-actin. Science 357, 703–706 (2017).
pubmed: 28818948
doi: 10.1126/science.aan2556
pmcid: 5821505
Rauskolb, C., Sun, S., Sun, G., Pan, Y. & Irvine, K. D. Cytoskeletal tension inhibits Hippo signaling through an Ajuba-Warts complex. Cell 158, 143–156 (2014).
pubmed: 24995985
doi: 10.1016/j.cell.2014.05.035
pmcid: 4082802
Maitre, J. L. & Heisenberg, C. P. The role of adhesion energy in controlling cell-cell contacts. Curr. Opin. Cell Biol. 23, 508–514 (2011).
pubmed: 21807491
doi: 10.1016/j.ceb.2011.07.004
pmcid: 3188705
Lenne, P. F., Rupprecht, J. F. & Viasnoff, V. Cell junction mechanics beyond the bounds of adhesion and tension. Dev. Cell 56, 202–212 (2021).
pubmed: 33453154
doi: 10.1016/j.devcel.2020.12.018
Harrison, O. J. et al. Two-step adhesive binding by classical cadherins. Nat. Struct. Mol. Biol. 17, 348–357 (2010).
pubmed: 20190754
doi: 10.1038/nsmb.1784
pmcid: 2872554
Pinheiro, D. & Bellaiche, Y. Mechanical force-driven adherens junction remodeling and epithelial dynamics. Dev. Cell 47, 3–19 (2018).
pubmed: 30300588
doi: 10.1016/j.devcel.2018.09.014
Yap, A. S., Brieher, W. M., Pruschy, M. & Gumbiner, B. M. Lateral clustering of the adhesive ectodomain: a fundamental determinant of cadherin function. Curr. Biol. 7, 308–315 (1997).
pubmed: 9133345
doi: 10.1016/S0960-9822(06)00154-0
Angres, B., Barth, A. & Nelson, W. J. Mechanism for transition from initial to stable cell-cell adhesion: kinetic analysis of E-cadherin-mediated adhesion using a quantitative adhesion assay. J. Cell Biol. 134, 549–557 (1996).
pubmed: 8707837
doi: 10.1083/jcb.134.2.549
Ward, M. D., Dembo, M. & Hammer, D. A. Kinetics of cell detachment: peeling of discrete receptor clusters. Biophys. J. 67, 2522–2534 (1994).
pubmed: 7696491
doi: 10.1016/S0006-3495(94)80742-8
pmcid: 1225638
Mulla, Y. et al. Weak catch bonds make strong networks. Nat. Mater. 21, 1019–1023 (2022).
pubmed: 36008604
doi: 10.1038/s41563-022-01288-0
pmcid: 7613626
Maitre, J. L. et al. Adhesion functions in cell sorting by mechanically coupling the cortices of adhering cells. Science 338, 253–256 (2012).
pubmed: 22923438
doi: 10.1126/science.1225399
David, R. et al. Tissue cohesion and the mechanics of cell rearrangement. Development 141, 3672–3682 (2014).
pubmed: 25249459
doi: 10.1242/dev.104315
Farhadifar, R., Roper, J. C., Aigouy, B., Eaton, S. & Julicher, F. The influence of cell mechanics, cell-cell interactions, and proliferation on epithelial packing. Curr. Biol. 17, 2095–2104 (2007).
pubmed: 18082406
doi: 10.1016/j.cub.2007.11.049
Clement, R., Dehapiot, B., Collinet, C., Lecuit, T. & Lenne, P. F. Viscoelastic dissipation stabilizes cell shape changes during tissue morphogenesis. Curr. Biol. 27, 3132–3142.e4 (2017).
pubmed: 28988857
doi: 10.1016/j.cub.2017.09.005
Iyer, K. V., Piscitello-Gomez, R., Paijmans, J., Julicher, F. & Eaton, S. Epithelial viscoelasticity is regulated by mechanosensitive E-cadherin turnover. Curr. Biol. 29, 578–591.e5 (2019).
pubmed: 30744966
doi: 10.1016/j.cub.2019.01.021
Borghi, N. et al. E-cadherin is under constitutive actomyosin-generated tension that is increased at cell-cell contacts upon externally applied stretch. Proc. Natl Acad. Sci. USA 109, 12568–12573 (2012).
pubmed: 22802638
doi: 10.1073/pnas.1204390109
pmcid: 3411997
Lagendijk, A. K. et al. Live imaging molecular changes in junctional tension upon VE-cadherin in zebrafish. Nat. Commun. 8, 1402 (2017).
pubmed: 29123087
doi: 10.1038/s41467-017-01325-6
pmcid: 5680264
Acharya, B. R. et al. Mammalian diaphanous 1 mediates a pathway for E-cadherin to stabilize epithelial barriers through junctional contractility. Cell Rep. 18, 2854–2867 (2017).
pubmed: 28329679
doi: 10.1016/j.celrep.2017.02.078
Bambardekar, K., Clement, R., Blanc, O., Chardes, C. & Lenne, P. F. Direct laser manipulation reveals the mechanics of cell contacts in vivo. Proc. Natl Acad. Sci. USA 112, 1416–1421 (2015).
pubmed: 25605934
doi: 10.1073/pnas.1418732112
pmcid: 4321260
Salbreux, G., Charras, G. & Paluch, E. Actin cortex mechanics and cellular morphogenesis. Trends Cell Biol. 22, 536–545 (2012).
pubmed: 22871642
doi: 10.1016/j.tcb.2012.07.001
Roh-Johnson, M. et al. Triggering a cell shape change by exploiting preexisting actomyosin contractions. Science 335, 1232–1235 (2012).
pubmed: 22323741
doi: 10.1126/science.1217869
pmcid: 3298882
Martin, A. C., Kaschube, M. & Wieschaus, E. F. Pulsed contractions of an actin-myosin network drive apical constriction. Nature 457, 495–499 (2009).
pubmed: 19029882
doi: 10.1038/nature07522
Bertet, C., Sulak, L. & Lecuit, T. Myosin-dependent junction remodelling controls planar cell intercalation and axis elongation. Nature 429, 667–671 (2004).
pubmed: 15190355
doi: 10.1038/nature02590
Rauzi, M., Lenne, P. F. & Lecuit, T. Planar polarized actomyosin contractile flows control epithelial junction remodelling. Nature 468, 1110–1114 (2010).
pubmed: 21068726
doi: 10.1038/nature09566
Curran, S. et al. Myosin II controls junction fluctuations to guide epithelial tissue ordering. Dev. Cell 43, 480–492.e6 (2017).
pubmed: 29107560
doi: 10.1016/j.devcel.2017.09.018
pmcid: 5703647
Fernandez-Gonzalez, R., Simoes Sde, M., Roper, J. C., Eaton, S. & Zallen, J. A. Myosin II dynamics are regulated by tension in intercalating cells. Dev. Cell 17, 736–743 (2009).
pubmed: 19879198
doi: 10.1016/j.devcel.2009.09.003
pmcid: 2854079
Michaux, J. B., Robin, F. B., McFadden, W. M. & Munro, E. M. Excitable RhoA dynamics drive pulsed contractions in the early C. elegans embryo. J. Cell Biol. 217, 4230–4252 (2018).
pubmed: 30275107
doi: 10.1083/jcb.201806161
pmcid: 6279378
Priya, R. et al. Feedback regulation through myosin II confers robustness on RhoA signalling at E-cadherin junctions. Nat. Cell Biol. 17, 1282–1293 (2015).
pubmed: 26368311
doi: 10.1038/ncb3239
Munjal, A., Philippe, J. M., Munro, E. & Lecuit, T. A self-organized biomechanical network drives shape changes during tissue morphogenesis. Nature 524, 351–355 (2015).
pubmed: 26214737
doi: 10.1038/nature14603
Brieher, W. M. & Yap, A. S. Cadherin junctions and their cytoskeleton(s). Curr. Opin. Cell Biol. 25, 39–46 (2013).
pubmed: 23127608
doi: 10.1016/j.ceb.2012.10.010
Reymann, A. C. et al. Actin network architecture can determine myosin motor activity. Science 336, 1310–1314 (2012).
pubmed: 22679097
doi: 10.1126/science.1221708
pmcid: 3649007
Haviv, L., Gillo, D., Backouche, F. & Bernheim-Groswasser, A. A cytoskeletal demolition worker: myosin II acts as an actin depolymerization agent. J. Mol. Biol. 375, 325–330 (2008).
pubmed: 18021803
doi: 10.1016/j.jmb.2007.09.066
Creton, C. & Ciccotti, M. Fracture and adhesion of soft materials: a review. Rep. Prog. Phys. 79, 046601 (2016).
pubmed: 27007412
doi: 10.1088/0034-4885/79/4/046601
Khalilgharibi, N., Fouchard, J., Recho, P., Charras, G. & Kabla, A. The dynamic mechanical properties of cellularised aggregates. Curr. Opin. Cell Biol. 42, 113–120 (2016).
pubmed: 27371889
doi: 10.1016/j.ceb.2016.06.003
Serwane, F. et al. In vivo quantification of spatially varying mechanical properties in developing tissues. Nat. Methods 14, 181–186 (2017).
pubmed: 27918540
doi: 10.1038/nmeth.4101
Mongera, A. et al. Mechanics of the cellular microenvironment as probed by cells in vivo during zebrafish presomitic mesoderm differentiation. Nat. Mater. 22, 135–143 (2023).
pubmed: 36577855
doi: 10.1038/s41563-022-01433-9
Le, T. L., Yap, A. S. & Stow, J. L. Recycling of E-cadherin: a potential mechanism for regulating cadherin dynamics. J. Cell Biol. 146, 219–232 (1999).
pubmed: 10402472
pmcid: 2199726
Lock, J. G. & Stow, J. L. Rab11 in recycling endosomes regulates the sorting and basolateral transport of E-cadherin. Mol. Biol. Cell 16, 1744–1755 (2005).
pubmed: 15689490
doi: 10.1091/mbc.e04-10-0867
pmcid: 1073657
Kozlov, M. M. & Bershadsky, A. D. Processive capping by formin suggests a force-driven mechanism of actin polymerization. J. Cell Biol. 167, 1011–1017 (2004).
pubmed: 15596547
doi: 10.1083/jcb.200410017
pmcid: 2172604
Carramusa, L., Ballestrem, C., Zilberman, Y. & Bershadsky, A. D. Mammalian diaphanous-related formin dia1 controls the organization of E-cadherin-mediated cell-cell junctions. J. Cell Sci. 120, 3870–3882 (2007).
pubmed: 17940061
doi: 10.1242/jcs.014365
Forgacs, G., Foty, R. A., Shafrir, Y. & Steinberg, M. S. Viscoelastic properties of living embryonic tissues: a quantitative study. Biophys. J. 74, 2227–2234 (1998).
pubmed: 9591650
doi: 10.1016/S0006-3495(98)77932-9
pmcid: 1299566
Guevorkian, K., Colbert, M. J., Durth, M., Dufour, S. & Brochard-Wyart, F. Aspiration of biological viscoelastic drops. Phys. Rev. Lett. 104, 218101 (2010).
pubmed: 20867138
doi: 10.1103/PhysRevLett.104.218101
Gonzalez-Rodriguez, D., Guevorkian, K., Douezan, S. & Brochard-Wyart, F. Soft matter models of developing tissues and tumors. Science 338, 910–917 (2012).
pubmed: 23161991
doi: 10.1126/science.1226418
Marmottant, P. et al. The role of fluctuations and stress on the effective viscosity of cell aggregates. Proc. Natl Acad. Sci. USA 106, 17271–17275 (2009).
pubmed: 19805170
doi: 10.1073/pnas.0902085106
pmcid: 2765233
Zhou, J., Kim, H. Y. & Davidson, L. A. Actomyosin stiffens the vertebrate embryo during crucial stages of elongation and neural tube closure. Development 136, 677–688 (2009).
pubmed: 19168681
doi: 10.1242/dev.026211
pmcid: 2685957
Zhou, J., Kim, H. Y., Wang, J. H. & Davidson, L. A. Macroscopic stiffening of embryonic tissues via microtubules, RhoGEF and the assembly of contractile bundles of actomyosin. Development 137, 2785–2794 (2010).
pubmed: 20630946
doi: 10.1242/dev.045997
pmcid: 2910388
Chu, C. W., Masak, G., Yang, J. & Davidson, L. A. From biomechanics to mechanobiology: Xenopus provides direct access to the physical principles that shape the embryo. Curr. Opin. Genet. Dev. 63, 71–77 (2020).
pubmed: 32563783
doi: 10.1016/j.gde.2020.05.011
pmcid: 9972463
von Dassow, M. & Davidson, L. A. Natural variation in embryo mechanics: gastrulation in Xenopus laevis is highly robust to variation in tissue stiffness. Dev. Dyn. 238, 2–18 (2009).
doi: 10.1002/dvdy.21809
Franze, K. Atomic force microscopy and its contribution to understanding the development of the nervous system. Curr. Opin. Genet. Dev. 21, 530–537 (2011).
pubmed: 21840706
doi: 10.1016/j.gde.2011.07.001
von Dassow, M., Strother, J. A. & Davidson, L. A. Surprisingly simple mechanical behavior of a complex embryonic tissue. PLoS ONE 5, e15359 (2010).
doi: 10.1371/journal.pone.0015359
von Dassow, M., Miller, C. J. & Davidson, L. A. Biomechanics and the thermotolerance of development. PLoS ONE 9, e95670 (2014).
doi: 10.1371/journal.pone.0095670
Jackson, T. R., Kim, H. Y., Balakrishnan, U. L., Stuckenholz, C. & Davidson, L. A. Spatiotemporally controlled mechanical cues drive progenitor mesenchymal-to-epithelial transition enabling proper heart formation and function. Curr. Biol. 27, 1326–1335 (2017).
pubmed: 28434863
doi: 10.1016/j.cub.2017.03.065
pmcid: 5497766
Barriga, E. H., Franze, K., Charras, G. & Mayor, R. Tissue stiffening coordinates morphogenesis by triggering collective cell migration in vivo. Nature 554, 523–527 (2018).
pubmed: 29443958
doi: 10.1038/nature25742
pmcid: 6013044
Marchant, C. L., Malmi-Kakkada, A. N., Espina, J. A. & Barriga, E. H. Cell clusters softening triggers collective cell migration in vivo. Nat. Mater. 21, 1314–1323 (2022).
pubmed: 35970965
doi: 10.1038/s41563-022-01323-0
pmcid: 9622418
Koser, D. E. et al. Mechanosensing is critical for axon growth in the developing brain. Nat. Neurosci. 19, 1592–1598 (2016).
pubmed: 27643431
doi: 10.1038/nn.4394
pmcid: 5531257
Nishizawa, K., Lin, S. Z., Chardes, C., Rupprecht, J. F. & Lenne, P. F. Two-point optical manipulation reveals mechanosensitive remodeling of cell-cell contacts in vivo. Proc. Natl Acad. Sci. USA 120, e2212389120 (2023).
pubmed: 36947511
doi: 10.1073/pnas.2212389120
pmcid: 10068846
Parada, C. et al. Mechanical feedback defines organizing centers to drive digit emergence. Dev. Cell 57, 854–866 e856 (2022).
pubmed: 35413235
doi: 10.1016/j.devcel.2022.03.004
Campas, O. et al. Quantifying cell-generated mechanical forces within living embryonic tissues. Nat. Methods 11, 183–189 (2014).
pubmed: 24317254
doi: 10.1038/nmeth.2761
Mohagheghian, E. et al. Quantifying compressive forces between living cell layers and within tissues using elastic round microgels. Nat. Commun. 9, 1878 (2018).
pubmed: 29760452
doi: 10.1038/s41467-018-04245-1
pmcid: 5951850
Traber, N. et al. Polyacrylamide bead sensors for in vivo quantification of cell-scale stress in zebrafish development. Sci. Rep. 9, 17031 (2019).
pubmed: 31745109
doi: 10.1038/s41598-019-53425-6
pmcid: 6864055
D’Angelo, A., Dierkes, K., Carolis, C., Salbreux, G. & Solon, J. In vivo force application reveals a fast tissue softening and external friction increase during early embryogenesis. Curr. Biol. 29, 1564–1571 e1566 (2019).
pubmed: 31031116
doi: 10.1016/j.cub.2019.04.010
pmcid: 6509404
Lau, K. et al. Anisotropic stress orients remodelling of mammalian limb bud ectoderm. Nat. Cell Biol. 17, 569–579 (2015).
pubmed: 25893915
doi: 10.1038/ncb3156
pmcid: 4955842
Kim, S., Pochitaloff, M., Stooke-Vaughan, G. A. & Campas, O. Embryonic tissues as active foams. Nat. Phys. 17, 859–866 (2021).
pubmed: 34367313
doi: 10.1038/s41567-021-01215-1
pmcid: 8336761
Roca-Cusachs, P., Conte, V. & Trepat, X. Quantifying forces in cell biology. Nat. Cell Biol. 19, 742–751 (2017).
pubmed: 28628082
doi: 10.1038/ncb3564
Benazeraf, B. & Pourquie, O. Formation and segmentation of the vertebrate body axis. Annu. Rev. Cell Dev. Biol. 29, 1–26 (2013).
pubmed: 23808844
doi: 10.1146/annurev-cellbio-101011-155703
Mongera, A., Michaut, A., Guillot, C., Xiong, F. & Pourquie, O. Mechanics of anteroposterior axis formation in vertebrates. Annu. Rev. Cell Dev. Biol. 35, 259–283 (2019).
pubmed: 31412208
doi: 10.1146/annurev-cellbio-100818-125436
pmcid: 7394480
McMillen, P. & Holley, S. A. The tissue mechanics of vertebrate body elongation and segmentation. Curr. Opin. Genet. Dev. 32, 106–111 (2015).
pubmed: 25796079
doi: 10.1016/j.gde.2015.02.005
pmcid: 4470730
Benazeraf, B. et al. A random cell motility gradient downstream of FGF controls elongation of an amniote embryo. Nature 466, 248–252 (2010).
pubmed: 20613841
doi: 10.1038/nature09151
pmcid: 3118990
Lawton, A. K. et al. Regulated tissue fluidity steers zebrafish body elongation. Development 140, 573–582 (2013).
pubmed: 23293289
doi: 10.1242/dev.090381
pmcid: 3561786
Liu, A. J. & Nagel, S. R. Nonlinear dynamics — jamming is not just cool any more. Nature 396, 21–22 (1998).
doi: 10.1038/23819
Sadati, M., Taheri Qazvini, N., Krishnan, R., Park, C. Y. & Fredberg, J. J. Collective migration and cell jamming. Differentiation 86, 121–125 (2013).
pubmed: 23791490
doi: 10.1016/j.diff.2013.02.005
pmcid: 3795803
Bonn, D., Denn, M. M., Berthier, L., Divoux, T. & Manneville, S. Yield stress materials in soft condensed matter.Rev. Mod. Phys. 89, 035005 (2017).
doi: 10.1103/RevModPhys.89.035005
Banavar, S. P. et al. Mechanical control of tissue shape and morphogenetic flows during vertebrate body axis elongation. Sci. Rep. 11, 8591 (2021).
pubmed: 33883563
doi: 10.1038/s41598-021-87672-3
pmcid: 8060277
Cohen-Addad, S., Hohler, R. & Pitois, O. Flow in foams and flowing foams. Annu. Rev. Fluid Mech. 45, 241–267 (2013).
doi: 10.1146/annurev-fluid-011212-140634
Sumi, A. et al. Adherens junction length during tissue contraction is controlled by the mechanosensitive activity of actomyosin and junctional recycling. Dev. Cell 47, 453–463.e453 (2018).
pubmed: 30458138
doi: 10.1016/j.devcel.2018.10.025
pmcid: 6291457
Wu, S. K. et al. Cortical F-actin stabilization generates apical-lateral patterns of junctional contractility that integrate cells into epithelia. Nat. Cell Biol. 16, 167–178 (2014).
pubmed: 24413434
doi: 10.1038/ncb2900
Khalilgharibi, N. et al. Stress relaxation in epithelial monolayers is controlled by the actomyosin cortex. Nat. Phys. 15, 839 (2019).
pubmed: 33569083
doi: 10.1038/s41567-019-0516-6
pmcid: 7116713
Cavanaugh, K. E., Staddon, M. F., Munro, E., Banerjee, S. & Gardel, M. L. RhoA mediates epithelial cell shape changes via mechanosensitive endocytosis. Dev. Cell 52, 152–166.e5 (2020).
pubmed: 31883774
doi: 10.1016/j.devcel.2019.12.002
Park, J. A. et al. Unjamming and cell shape in the asthmatic airway epithelium. Nat. Mater. 14, 1040–1048 (2015).
pubmed: 26237129
doi: 10.1038/nmat4357
pmcid: 4666305
Morita, H. et al. The physical basis of coordinated tissue spreading in zebrafish gastrulation. Dev. Cell 40, 354–366 e354 (2017).
pubmed: 28216382
doi: 10.1016/j.devcel.2017.01.010
pmcid: 5364273
Petridou, N. I., Corominas-Murtra, B., Heisenberg, C. P. & Hannezo, E. Rigidity percolation uncovers a structural basis for embryonic tissue phase transitions. Cell 184, 1914–1928.e19 (2021).
pubmed: 33730596
doi: 10.1016/j.cell.2021.02.017
pmcid: 8055543
Van Hecke, M. Jamming of soft particles: geometry, mechanics, scaling and isostaticity. J. Phys. Condens Mat. 22, 033101 (2010).
doi: 10.1088/0953-8984/22/3/033101
Engl, W., Arasi, B., Yap, L. L., Thiery, J. P. & Viasnoff, V. Actin dynamics modulate mechanosensitive immobilization of E-cadherin at adherens junctions. Nat. Cell Biol. 16, 587–594 (2014).
pubmed: 24859003
doi: 10.1038/ncb2973
Smutny, M. et al. Myosin II isoforms identify distinct functional modules that support integrity of the epithelial zonula adherens. Nat. Cell Biol. 12, 696–702 (2010).
pubmed: 20543839
doi: 10.1038/ncb2072
pmcid: 3428211
Dumortier, J. G. et al. Hydraulic fracturing and active coarsening position the lumen of the mouse blastocyst. Science 365, 465–468 (2019).
pubmed: 31371608
doi: 10.1126/science.aaw7709
Yang, Q. et al. Cell fate coordinates mechano-osmotic forces in intestinal crypt formation. Nat. Cell Biol. 23, 733–744 (2021).
pubmed: 34155381
doi: 10.1038/s41556-021-00700-2
pmcid: 7611267
Schliffka, M. F. & Maitre, J. L. Stay hydrated: basolateral fluids shaping tissues. Curr. Opin. Genet. Dev. 57, 70–77 (2019).
pubmed: 31445440
doi: 10.1016/j.gde.2019.06.015
Barua, D., Nagel, M. & Winklbauer, R. Cell-cell contact landscapes in Xenopus gastrula tissues. Proc. Natl Acad. Sci. USA 118, e2017953118 (2021).
doi: 10.1073/pnas.2107953118
Ventura, G. et al. Multiciliated cells use filopodia to probe tissue mechanics during epithelial integration in vivo. Nat. Commun. 13, 6423 (2022).
pubmed: 36307428
doi: 10.1038/s41467-022-34165-0
pmcid: 9616887
Guignard, L. et al. Contact area-dependent cell communication and the morphological invariance of ascidian embryogenesis. Science 369, eaar5663 (2020).
pubmed: 32646972
doi: 10.1126/science.aar5663
Bi, D. P., Lopez, J. H., Schwarz, J. M. & Manning, M. L. A density-independent rigidity transition in biological tissues. Nat. Phys. 11, 1074 (2015).
doi: 10.1038/nphys3471
Heisenberg, C. P. & Bellaiche, Y. Forces in tissue morphogenesis and patterning. Cell 153, 948–962 (2013).
pubmed: 23706734
doi: 10.1016/j.cell.2013.05.008
Johnson, J. L., Najor, N. A. & Green, K. J. Desmosomes: regulators of cellular signaling and adhesion in epidermal health and disease. Cold Spring Harb. Perspect. Med. 4, a015297 (2014).
pubmed: 25368015
doi: 10.1101/cshperspect.a015297
pmcid: 4208714
Broussard, J. A. et al. The desmoplakin-intermediate filament linkage regulates cell mechanics. Mol. Biol. Cell 28, 3156–3164 (2017).
pubmed: 28495795
doi: 10.1091/mbc.e16-07-0520
pmcid: 5687018
Harris, A. R. et al. Characterizing the mechanics of cultured cell monolayers. Proc. Natl Acad. Sci. USA 109, 16449–16454 (2012).
pubmed: 22991459
doi: 10.1073/pnas.1213301109
pmcid: 3478631
Latorre, E. et al. Active superelasticity in three-dimensional epithelia of controlled shape. Nature 563, 203–208 (2018).
pubmed: 30401836
doi: 10.1038/s41586-018-0671-4
pmcid: 6520229
Bonfanti, A., Duque, J., Kabla, A. & Charras, G. Fracture in living tissues. Trends Cell Biol. 32, 537–551 (2022).
pubmed: 35190218
doi: 10.1016/j.tcb.2022.01.005
Casares, L. et al. Hydraulic fracture during epithelial stretching. Nat. Mater. 14, 343–351 (2015).
pubmed: 25664452
doi: 10.1038/nmat4206
pmcid: 4374166
Blanchard, G. B. et al. Tissue tectonics: morphogenetic strain rates, cell shape change and intercalation. Nat. Methods 6, 458–464 (2009).
pubmed: 19412170
doi: 10.1038/nmeth.1327
pmcid: 4894466
Teomy, E., Kessler, D. A. & Levine, H. Confluent and nonconfluent phases in a model of cell tissue. Phys. Rev. E 98, 042418 (2018).
doi: 10.1103/PhysRevE.98.042418
Higashi, T. & Miller, A. L. Tricellular junctions: how to build junctions at the TRICkiest points of epithelial cells. Mol. Biol. Cell 28, 2023–2034 (2017).
pubmed: 28705832
doi: 10.1091/mbc.e16-10-0697
pmcid: 5509417
Uechi, H. & Kuranaga, E. The tricellular junction protein sidekick regulates vertex dynamics to promote bicellular junction extension. Dev. Cell 50, 327–338.e5 (2019).
pubmed: 31353316
doi: 10.1016/j.devcel.2019.06.017
Letizia, A. et al. Sidekick is a key component of tricellular adherens junctions that acts to resolve cell rearrangements. Dev. Cell 50, 313–326.e5 (2019).
pubmed: 31353315
doi: 10.1016/j.devcel.2019.07.007
pmcid: 6748646
Tang, H. et al. Architecture of cell-cell adhesion mediated by sidekicks. Proc. Natl Acad. Sci. USA 115, 9246–9251 (2018).
pubmed: 30150416
doi: 10.1073/pnas.1801810115
pmcid: 6140505
Cho, Y. et al. Tricellulin secures the epithelial barrier at tricellular junctions by interacting with actomyosin. J. Cell Biol. 221, e202009037 (2022).
pubmed: 35148372
doi: 10.1083/jcb.202009037
pmcid: 8847807
Choi, W. et al. Remodeling the zonula adherens in response to tension and the role of afadin in this response. J. Cell Biol. 213, 243–260 (2016).
pubmed: 27114502
doi: 10.1083/jcb.201506115
pmcid: 5084271
Krndija, D. et al. Active cell migration is critical for steady-state epithelial turnover in the gut. Science 365, 705–710 (2019).
pubmed: 31416964
doi: 10.1126/science.aau3429
Krajnc, M. Solid-fluid transition and cell sorting in epithelia with junctional tension fluctuations. Soft Matter 16, 3209–3215 (2020).
pubmed: 32159536
doi: 10.1039/C9SM02310K
Levayer, R. & Lecuit, T. Biomechanical regulation of contractility: spatial control and dynamics. Trends Cell Biol. 22, 61–81 (2012).
pubmed: 22119497
doi: 10.1016/j.tcb.2011.10.001
Tetley, R. J. & Mao, Y. The same but different: cell intercalation as a driver of tissue deformation and fluidity. Philos. Trans. R Soc. Lond. 373, 20170328 (2018).
doi: 10.1098/rstb.2017.0328
Malinverno, C. et al. Endocytic reawakening of motility in jammed epithelia. Nat. Mater. 16, 587–596 (2017).
pubmed: 28135264
doi: 10.1038/nmat4848
pmcid: 5407454
Ilina, O. et al. Cell-cell adhesion and 3D matrix confinement determine jamming transitions in breast cancer invasion. Nat. Cell Biol. 22, 1103–1115 (2020).
pubmed: 32839548
doi: 10.1038/s41556-020-0552-6
pmcid: 7502685
Tachibana, K. et al. Two cell adhesion molecules, nectin and cadherin, interact through their cytoplasmic domain-associated proteins. J. Cell Biol. 150, 1161–1175 (2000).
pubmed: 10974003
doi: 10.1083/jcb.150.5.1161
pmcid: 2175253
Takahashi, K. et al. Nectin/PRR: an immunoglobulin-like cell adhesion molecule recruited to cadherin-based adherens junctions through interaction with afadin, a PDZ domain-containing protein. J. Cell Biol. 145, 539–549 (1999).
pubmed: 10225955
doi: 10.1083/jcb.145.3.539
pmcid: 2185068
Lin, H. P. et al. Cell adhesion molecule Echinoid associates with unconventional myosin VI/Jaguar motor to regulate cell morphology during dorsal closure in Drosophila. Dev. Biol. 311, 423–433 (2007).
pubmed: 17936269
doi: 10.1016/j.ydbio.2007.08.043
McLachlan, R. W. & Yap, A. S. Not so simple: the complexity of phosphotyrosine signaling at cadherin adhesive contacts. J. Mol. Med. 85, 545–554 (2007).
pubmed: 17429596
doi: 10.1007/s00109-007-0198-x
Braga, V. M. & Yap, A. S. The challenges of abundance: epithelial junctions and small GTPase signalling. Curr. Opin. Cell Biol. 17, 466–474 (2005).
pubmed: 16112561
doi: 10.1016/j.ceb.2005.08.012
Ireton, R. C. et al. A novel role for p120 catenin in E-cadherin function. J. Cell Biol. 159, 465–476 (2002).
pubmed: 12427869
doi: 10.1083/jcb.200205115
pmcid: 2173073
Nanes, B. A. et al. p120-catenin binding masks an endocytic signal conserved in classical cadherins. J. Cell Biol. 199, 365–380 (2012).
pubmed: 23071156
doi: 10.1083/jcb.201205029
pmcid: 3471230
Oakes, P. W. et al. Optogenetic control of RhoA reveals zyxin-mediated elasticity of stress fibres. Nat. Commun. 8, 15817 (2017).
pubmed: 28604737
doi: 10.1038/ncomms15817
pmcid: 5477492
Toh, P. J. Y. et al. Optogenetic control of YAP cellular localisation and function. EMBO Rep. 23, e54401 (2022).
pubmed: 35876586
doi: 10.15252/embr.202154401
pmcid: 9442306
Ollech, D. et al. An optochemical tool for light-induced dissociation of adherens junctions to control mechanical coupling between cells. Nat. Commun. 11, 472 (2020).
pubmed: 31980653
doi: 10.1038/s41467-020-14390-1
pmcid: 6981158
Izquierdo, E., Quinkler, T. & De Renzis, S. Guided morphogenesis through optogenetic activation of Rho signalling during early Drosophila embryogenesis. Nat. Commun. 9, 2366 (2018).
pubmed: 29915285
doi: 10.1038/s41467-018-04754-z
pmcid: 6006163
Wu, S. K., Budnar, S., Yap, A. S. & Gomez, G. A. Pulsatile contractility of actomyosin networks organizes the cellular cortex at lateral cadherin junctions. Eur. J. Cell Biol. 93, 396–404 (2014).
pubmed: 25269995
doi: 10.1016/j.ejcb.2014.09.001
Chugh, P. & Paluch, E. K. The actin cortex at a glance. J. Cell Sci. 131, jcs186254 (2018).
pubmed: 30026344
doi: 10.1242/jcs.186254
pmcid: 6080608
Bovellan, M. et al. Cellular control of cortical actin nucleation. Curr. Biol. 24, 1628–1635 (2014).
pubmed: 25017211
doi: 10.1016/j.cub.2014.05.069
pmcid: 4110400
Cao, L. et al. SPIN90 associates with mDia1 and the Arp2/3 complex to regulate cortical actin organization. Nat. Cell Biol. 22, 803–814 (2020).
pubmed: 32572169
doi: 10.1038/s41556-020-0531-y
Noordstra, I. et al. An E-cadherin-actin clutch translates the mechanical force of cortical flow for cell-cell contact to inhibit epithelial cell locomotion. Dev. Cell 58, 1748–1763.e6 (2023).
pubmed: 37480844
doi: 10.1016/j.devcel.2023.06.011
Padmanabhan, A., Ong, H. T. & Zaidel-Bar, R. Non-junctional E-cadherin clusters regulate the actomyosin cortex in the c. elegans zygote. Curr. Biol. 27, 103–112 (2017).
pubmed: 27989674
doi: 10.1016/j.cub.2016.10.032
Kovacs, E. M. et al. N-WASP regulates the epithelial junctional actin cytoskeleton through a non-canonical post-nucleation pathway. Nat. Cell Biol. 13, 934–943 (2011).
pubmed: 21785420
doi: 10.1038/ncb2290
Tang, V. W. & Brieher, W. α-Actinin-4/FSGS1 is required for Arp2/3-dependent actin assembly at the adherens junction. J. Cell Biol. 196, 115–130 (2012).
pubmed: 22232703
doi: 10.1083/jcb.201103116
pmcid: 3255975
Harrison, O. J. et al. The extracellular architecture of adherens junctions revealed by crystal structures of type I cadherins. Structure 19, 244–256 (2011).
pubmed: 21300292
doi: 10.1016/j.str.2010.11.016
pmcid: 3070544
Brasch, J., Harrison, O. J., Honig, B. & Shapiro, L. Thinking outside the cell: how cadherins drive adhesion. Trends Cell Biol. 22, 299–310 (2012).
pubmed: 22555008
doi: 10.1016/j.tcb.2012.03.004
pmcid: 3385655
Wu, Y., Kanchanawong, P. & Zaidel-Bar, R. Actin-delimited adhesion-independent clustering of e-cadherin forms the nanoscale building blocks of adherens junctions. Dev. Cell 32, 139–154 (2015).
pubmed: 25600236
doi: 10.1016/j.devcel.2014.12.003
Truong Quang, B. A., Mani, M., Markova, O., Lecuit, T. & Lenne, P. F. Principles of E-cadherin supramolecular organization in vivo. Curr. Biol. 23, 2197–2207 (2013).
pubmed: 24184100
doi: 10.1016/j.cub.2013.09.015
Duong, C. N. et al. Force-induced changes of alpha-catenin conformation stabilize vascular junctions independently of vinculin. J. Cell Sci. 134, jcs259012 (2021).
pubmed: 34851405
doi: 10.1242/jcs.259012
pmcid: 8729784
Gowrishankar, K. et al. Active remodeling of cortical actin regulates spatiotemporal organization of cell surface molecules. Cell 149, 1353–1367 (2012).
pubmed: 22682254
doi: 10.1016/j.cell.2012.05.008
Chandran, R., Kale, G., Philippe, J. M., Lecuit, T. & Mayor, S. Distinct actin-dependent nanoscale assemblies underlie the dynamic and hierarchical organization of E-cadherin. Curr. Biol. 31, 1726–1736 e1724 (2021).
pubmed: 33607036
doi: 10.1016/j.cub.2021.01.059
pmcid: 7611338
Yap, A. S., Gomez, G. A. & Parton, R. G. Adherens junctions revisualized: organizing cadherins as nanoassemblies. Dev. Cell 35, 12–20 (2015).
pubmed: 26460944
doi: 10.1016/j.devcel.2015.09.012
Beutel, O., Maraspini, R., Pombo-Garcia, K., Martin-Lemaitre, C. & Honigmann, A. Phase separation of zonula occludens proteins drives formation of tight junctions. Cell 179, 923–936 e911 (2019).
pubmed: 31675499
doi: 10.1016/j.cell.2019.10.011
Schwayer, C. et al. Mechanosensation of tight junctions depends on ZO-1 phase separation and flow. Cell 179, 937–952.e8 (2019).
pubmed: 31675500
doi: 10.1016/j.cell.2019.10.006
Yan, V. T., Narayanan, A., Wiegand, T., Julicher, F. & Grill, S. W. A condensate dynamic instability orchestrates actomyosin cortex activation. Nature 609, 597–604 (2022).
pubmed: 35978196
doi: 10.1038/s41586-022-05084-3
pmcid: 9477739
Kwak, M. et al. Adherens junctions organize size-selective proteolytic hotspots critical for Notch signalling. Nat. Cell Biol. 24, 1739–1753 (2022).
pubmed: 36456828
doi: 10.1038/s41556-022-01031-6
pmcid: 10665132
Guo, M. et al. Probing the stochastic, motor-driven properties of the cytoplasm using force spectrum microscopy. Cell 158, 822–832 (2014).
pubmed: 25126787
doi: 10.1016/j.cell.2014.06.051
pmcid: 4183065
Bausch, A. R., Moller, W. & Sackmann, E. Measurement of local viscoelasticity and forces in living cells by magnetic tweezers. Biophys. J. 76, 573–579 (1999).
pubmed: 9876170
doi: 10.1016/S0006-3495(99)77225-5
pmcid: 1302547
Wessel, A. D., Gumalla, M., Grosshans, J. & Schmidt, C. F. The mechanical properties of early Drosophila embryos measured by high-speed video microrheology. Biophys. J. 108, 1899–1907 (2015).
pubmed: 25902430
doi: 10.1016/j.bpj.2015.02.032
pmcid: 4407248
Stehbens, S. J. et al. Dynamic microtubules regulate the local concentration of E-cadherin at cell-cell contacts. J. Cell Sci. 119, 1801–1811 (2006).
pubmed: 16608875
doi: 10.1242/jcs.02903
Bakir, B., Chiarella, A. M., Pitarresi, J. R. & Rustgi, A. K. EMT, MET, plasticity, and tumor metastasis. Trends Cell Biol. 30, 764–776 (2020).
pubmed: 32800658
doi: 10.1016/j.tcb.2020.07.003
pmcid: 7647095
Dongre, A. & Weinberg, R. A. New insights into the mechanisms of epithelial-mesenchymal transition and implications for cancer. Nat. Rev. Mol. Cell Biol. 20, 69–84 (2019).
pubmed: 30459476
doi: 10.1038/s41580-018-0080-4
Lamouille, S., Xu, J. & Derynck, R. Molecular mechanisms of epithelial-mesenchymal transition. Nat. Rev. Mol. Cell Biol. 15, 178–196 (2014).
pubmed: 24556840
doi: 10.1038/nrm3758
pmcid: 4240281
La Porta, C. A. M. & Zapperi, S. Phase transitions in cell migration. Nat. Rev. Phys. 2, 516–517 (2020).
doi: 10.1038/s42254-020-0213-5
Mitchel, J. A. et al. In primary airway epithelial cells, the unjamming transition is distinct from the epithelial-to-mesenchymal transition. Nat. Commun. 11, 5053 (2020).
pubmed: 33028821
doi: 10.1038/s41467-020-18841-7
pmcid: 7542457
Togashi, H. et al. Nectins establish a checkerboard-like cellular pattern in the auditory epithelium. Science 333, 1144–1147 (2011).
pubmed: 21798896
doi: 10.1126/science.1208467
Wei, S. Y. et al. Echinoid is a component of adherens junctions that cooperates with DE-Cadherin to mediate cell adhesion. Dev. Cell 8, 493–504 (2005).
pubmed: 15809032
doi: 10.1016/j.devcel.2005.03.015
Chang, L. H. et al. Differential adhesion and actomyosin cable collaborate to drive Echinoid-mediated cell sorting. Development 138, 3803–3812 (2011).
pubmed: 21795280
doi: 10.1242/dev.062257
Michael, M. & Yap, A. S. The regulation and functional impact of actin assembly at cadherin cell-cell adhesions. Semin. Cell Dev. Biol. 24, 298–307 (2013).
pubmed: 23333496
doi: 10.1016/j.semcdb.2012.12.004