Key Points
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In many biological situations in vivo, including tissue shaping during morphogenesis, tissue repair and cancer invasion, cells do not move as single bodies but as a collective.
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Two main mechanisms support collective dynamics: polarized collective cell migration and coordinated contractile processes of cell groups that involve multicellular actomyosin-based structures.
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In vitro wound-healing assays exploiting microfabricated devices have been models of choice to study collective cell behaviours. Such in vitro approaches are the most important methods to achieve multiscale analysis from the molecular to the multicellular level.
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In contrast to a single cell, collective cell migration relies not only on the interactions with the extracellular matrix but also with neighbouring cells.
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Coordinated movements strongly depend on intercellular interactions via mechanosensitive cadherin-based adhesions.
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Cellular coordination is a mechanoregulated multiscale process integrating events at the molecular, cellular and multicellular scales, and it occurs at a wide range of timescales, from milliseconds to minutes to days.
Abstract
The way in which cells coordinate their behaviours during various biological processes, including morphogenesis, cancer progression and tissue remodelling, largely depends on the mechanical properties of the external environment. In contrast to single cells, collective cell behaviours rely on the cellular interactions not only with the surrounding extracellular matrix but also with neighbouring cells. Collective dynamics is not simply the result of many individually moving blocks. Instead, cells coordinate their movements by actively interacting with each other. These mechanisms are governed by mechanosensitive adhesion complexes at the cell–substrate interface and cell–cell junctions, which respond to but also further transmit physical signals. The mechanosensitivity and mechanotransduction at adhesion complexes are important for regulating tissue cohesiveness and thus are important for collective cell behaviours. Recent studies have shown that the physical properties of the cellular environment, which include matrix stiffness, topography, geometry and the application of external forces, can alter collective cell behaviours, tissue organization and cell-generated forces. On the basis of these findings, we can now start building our understanding of the mechanobiology of collective cell movements that span over multiple length scales from the molecular to the tissue level.
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References
Friedl, P. & Gilmour, D. Collective cell migration in morphogenesis, regeneration and cancer. Nat. Rev. Mol. Cell Biol. 10, 445–457 (2009).
Bianco, A. et al. Two distinct modes of guidance signalling during collective migration of border cells. Nature 448, 362–365 (2007).
Lecaudey, V., Cakan-Akdogan, G., Norton, W. H. & Gilmour, D. Dynamic Fgf signaling couples morphogenesis and migration in the zebrafish lateral line primordium. Development 135, 2695–2705 (2008).
Nabeshima, K., Inoue, T., Shimao, Y., Kataoka, H. & Koono, M. Cohort migration of carcinoma cells: differentiated colorectal carcinoma cells move as coherent cell clusters or sheets. Histol. Histopathol. 14, 1183–1197 (1999).
Einenkel, J., Braumann, U. D., Horn, L. C., Kuska, J. P. & Hockel, M. 3D analysis of the invasion front in squamous cell carcinoma of the uterine cervix: histopathologic evidence for collective invasion per continuitatem. Anal. Quantitative Cytol. Histol. 29, 279–290 (2007).
Giampieri, S. et al. Localized and reversible TGFβ signalling switches breast cancer cells from cohesive to single cell motility. Nat. Cell Biol. 11, 1287–1296 (2009).
Behrndt, M. et al. Forces driving epithelial spreading in zebrafish gastrulation. Science 338, 257–260 (2012).
Abreu-Blanco, M. T., Verboon, J. M., Liu, R., Watts, J. J. & Parkhurst, S. M. Drosophila embryos close epithelial wounds using a combination of cellular protrusions and an actomyosin purse string. J. Cell Sci. 125, 5984–5997 (2012).
Campas, O. et al. Quantifying cell-generated mechanical forces within living embryonic tissues. Nat. Methods 11, 183–189 (2014). In this paper, the authors develop a method to measure the mechanical forces that cells exert while building tissues. They insert calibrated oil droplets in living embryos in contact with other cells. When cells push and pull on an oil droplet, they deform it, and this deformation provides a direct readout of the pressure they exert.
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).
Hakim, V. & Silberzan, P. Collective cell migration: a physics perspective. Reports on progress in physics. Phys. Soc. 80, 076601 (2017).
Marchetti, M. C. et al. Hydrodynamics of soft active matter. Rev. Modern Phys. 85, 1143 (2013).
Parsons, J. T., Horwitz, A. R. & Schwartz, M. A. Cell adhesion: integrating cytoskeletal dynamics and cellular tension. Nat. Rev. Mol. Cell Biol. 11, 633–643 (2010).
Kanchanawong, P. et al. Nanoscale architecture of integrin-based cell adhesions. Nature 468, 580–584 (2010).
Sarangi, B. R. et al. Coordination between intra- and extracellular forces regulates focal adhesion dynamics. Nano Lett. 17, 399–406 (2017).
Lo, C. M., Wang, H. B., Dembo, M. & Wang, Y. L. Cell movement is guided by the rigidity of the substrate. Biophys. J. 79, 144–152 (2000).
Sniadecki, N. J. et al. Magnetic microposts as an approach to apply forces to living cells. Proc. Natl Acad. Sci. USA 104, 14553–14558 (2007).
Saxena, M. et al. EGFR and HER2 activate rigidity sensing only on rigid matrices. Nat. Mater. 16, 775–781 (2017).
Ladoux, B., Mège, R. M. & Trepat, X. Front-rear polarization by mechanical cues: from single cells to tissues. Trends Cell Biol. 26, 420–433 (2016).
Cetera, M. et al. Epithelial rotation promotes the global alignment of contractile actin bundles during Drosophila egg chamber elongation. Nature Commun. 5, 5511 (2014). During the elongation of the D. melanogaster egg chamber into an ellipsoidal shape, the follicular epithelial cells undergo a collective migration that causes the egg chamber to rotate within its surrounding basement membrane. This study demonstrates that collective rotation plays a critical role in building up the actin-based component of a corset that forms during the elongation, in which actin bundles in the epithelium and fibrils in the basement membrane are all aligned perpendicular to the elongation axis.
Bertet, C., Sulak, L. & Lecuit, T. Myosin-dependent junction remodelling controls planar cell intercalation and axis elongation. Nature 429, 667–671 (2004).
Dona, E. et al. Directional tissue migration through a self-generated chemokine gradient. Nature 503, 285–289 (2013).
Szabo, A. et al. In vivo confinement promotes collective migration of neural crest cells. J. Cell Biol. 213, 543–555 (2016). Combining computational and experimental approaches, the authors show that the in vivo collective migration of neural crest cells depends on confinement and geometrical constraints. They demonstrate that such confinement promotes directional migration of cell groups in vivo.
Haeger, A., Krause, M., Wolf, K. & Friedl, P. Cell jamming: collective invasion of mesenchymal tumor cells imposed by tissue confinement. Biochim. Biophys. Acta 1840, 2386–2395 (2014).
Abercrombie, M. & Ambrose, E. J. The surface properties of cancer cells: a review. Cancer Res. 22, 525–548 (1962).
Poujade, M. et al. Collective migration of an epithelial monolayer in response to a model wound. Proc. Natl Acad. Sci. USA 104, 15988–15993 (2007).
Deforet, M. et al. Automated velocity mapping of migrating cell populations (AVeMap). Nat. Methods 9, 1081–1083 (2012).
Mayor, R. & Etienne-Manneville, S. The front and rear of collective cell migration. Nat. Rev. Mol. Cell Biol. 17, 97–109 (2016).
Reffay, M. et al. Orientation and polarity in collectively migrating cell structures: statics and dynamics. Biophys. J. 100, 2566–2575 (2011).
Purcell, E. M. Life at low Reynolds number. Am. J. Phys. 45, 3–11 (1977).
Vedula, S. R. K. et al. Emerging modes of collective cell migration induced by geometrical constraints. Proc. Natl Acad. Sci. USA 109, 12974–12979 (2012). In this study, the authors show that collective cell migration can be regulated by confinement. Cells migrate more quickly on narrow lines of the size of a single cell than on larger stripes of several cell diameters, and the mode of migration varies depending on the width of the micropatterned lines, consistent with different patterns of force transmission through cell–cell junctions.
Foty, R. A. & Steinberg, M. S. The differential adhesion hypothesis: a direct evaluation. Dev. Biol. 278, 255–263 (2005).
Ranft, J. et al. Fluidization of tissues by cell division and apoptosis. Proc. Natl Acad. Sci. USA 107, 20863–20868 (2010).
Lecuit, T. & Yap, A. S. E-Cadherin junctions as active mechanical integrators in tissue dynamics. Nat. Cell Biol. 17, 533–539 (2015).
Rossen, N. S., Tarp, J. M., Mathiesen, J., Jensen, M. H. & Oddershede, L. B. Long-range ordered vorticity patterns in living tissue induced by cell division. Nature Commun. 5, 5720 (2014). In this study, the authors analyse the dynamic movements of endothelial cells induced by cell division. Cell divisions induce long-range reorganization of cells together with the formation of vortex patterns that extend far away from the division site.
Saw, T. B. et al. Topological defects in epithelia govern cell death and extrusion. Nature 544, 212–216 (2017). This paper shows that epithelial cells are elongated and closely packed, which means that they can spontaneously align in a similar way to the molecules in nematic liquid crystals. Accordingly, compressive stresses induced by oriented ordering and defects in the epithelium provide a physical trigger for cell extrusion and death.
Petitjean, L. et al. Velocity fields in a collectively migrating epithelium. Biophys. J. 98, 1790–1800 (2010).
Vedula, S. R. K. et al. Epithelial bridges maintain tissue integrity during collective cell migration. Nat. Mater. 13, 87–96 (2014).
Das, T. et al. A molecular mechanotransduction pathway regulates collective migration of epithelial cells. Nat. Cell Biol. 17, 276–287 (2015). In this study, the authors show that the protein Merlin is involved in coordinating motility forces from the edge into the bulk of the monolayer.
Bazellieres, E. et al. Control of cell-cell forces and collective cell dynamics by the intercellular adhesome. Nat. Cell Biol. 17, 409–420 (2015). The paper analyses the role of molecules at desmosomes, tight junctions and adherens junctions involved in cell–cell interactions in the establishment of forces and stresses during collective cell migration. It also shows that E-cadherins and P-cadherins respond to the level of force or to the rate at which the intercellular stress builds up, respectively, which allows for a particularly efficient control of mechanosensation at cadherin-based junctions.
Basan, M., Elgeti, J., Hannezo, E., Rappel, W. J. & Levine, H. Alignment of cellular motility forces with tissue flow as a mechanism for efficient wound healing. Proc. Natl Acad. Sci. USA 110, 2452–2459 (2013).
Farooqui, R. & Fenteany, G. Multiple rows of cells behind an epithelial wound edge extend cryptic lamellipodia to collectively drive cell-sheet movement. J. Cell Sci. 118, 51–63 (2005).
Takeichi, M. Dynamic contacts: rearranging adherens junctions to drive epithelial remodelling. Nat. Rev. Mol. Cell Biol. 15, 397–410 (2014).
Brugues, A. et al. Forces driving epithelial wound healing. Nat. Phys. 10, 684–691 (2014).
Vedula, S. R. K. et al. Mechanics of epithelial closure over non-adherent environments. Nat. Commun. 6, 6111 (2015).
Clark, A. G. et al. Integration of single and multicellular wound responses. Curr. Biol. 19, 1389–1395 (2009).
Wood, W. et al. Wound healing recapitulates morphogenesis in Drosophila embryos. Nat. Cell Biol. 4, 907–912 (2002).
Tamada, M., Perez, T. D., Nelson, W. J. & Sheetz, M. P. Two distinct modes of myosin assembly and dynamics during epithelial wound closure. J. Cell Biol. 176, 27–33 (2007).
Anon, E. et al. Cell crawling mediates collective cell migration to close undamaged epithelial gaps. Proc. Natl Acad. Sci. USA 109, 10891–10896 (2012).
Cochet-Escartin, O., Ranft, J., Silberzan, P. & Marcq, P. Border forces and friction control epithelial closure dynamics. Biophys. J. 106, 65–73 (2014).
Vitorino, P. & Meyer, T. Modular control of endothelial sheet migration. Genes Dev. 22, 3268–3281 (2008).
Ng, M. R., Besser, A., Danuser, G. & Brugge, J. S. Substrate stiffness regulates cadherin-dependent collective migration through myosin-II contractility. J. Cell Biol. 199, 545–563 (2012).
Gumbiner, B. Generation and maintenance of epithelial cell polarity. Curr. Opin. Cell Biol. 2, 881–887 (1990).
Sunyer, R. et al. Collective cell durotaxis emerges from long-range intercellular force transmission. Science 353, 1157–1161 (2016). In this study, the authors show that collective cell migration can be regulated by substrate stiffness, with cells moving towards stiff substrates — cells on a stiffer substrate have more resistance, enabling them to generate higher traction forces, so they win the tug of war with cells on the softer part. They further show that even cells that do not show durotaxis as single cells can durotax as a collective.
Abercrombie, M. & Heaysman, J. E. Observations on the social behaviour of cells in tissue culture: I. Speed of movement of chick heart fibroblasts in relation to their mutual contacts. Exp. Cell Res. 5, 111–131 (1953).
Carmona-Fontaine, C. et al. Contact inhibition of locomotion in vivo controls neural crest directional migration. Nature 456, 957–961 (2008).
Theveneau, E. et al. Chase-and-run between adjacent cell populations promotes directional collective migration. Nat. Cell Biol. 15, 763–772 (2013). In this paper, the authors show the importance of multiple cellular interactions of different cell types for collective cell migration. They study the interactions between neural crest cells and placode cells, an epithelial tissue, identifying a mechanism based on a coupling between chemotaxis and mechanical forces that promotes the correct migration of neural crest cells.
Bahm, I. et al. PDGF controls contact inhibition of locomotion by regulating N-cadherin during neural crest migration. Development 144, 2456–2468 (2017).
Desai, R. A., Gao, L., Raghavan, S., Liu, W. F. & Chen, C. S. Cell polarity triggered by cell-cell adhesion via E-cadherin. J. Cell Sci. 122, 905–911 (2009).
Ouyang, M. et al. N-Cadherin regulates spatially polarized signals through distinct p120ctn and β-catenin-dependent signalling pathways. Nat. Commun. 4, 1589 (2013).
Kocgozlu, L. et al. Epithelial cell packing induces distinct modes of cell extrusions. Curr. Biol. 26, 2942–2950 (2016).
Park, J. A., Atia, L., Mitchel, J. A., Fredberg, J. J. & Butler, J. P. Collective migration and cell jamming in asthma, cancer and development. J. Cell Sci. 129, 3375–3383 (2016).
Garcia, S. et al. Physics of active jamming during collective cellular motion in a monolayer. Proc. Natl Acad. Sci. USA 112, 15314–15319 (2015).
Deforet, M., Hakim, V., Yevick, H., Duclos, G. & Silberzan, P. Emergence of collective modes and tri-dimensional structures from epithelial confinement. Nat. Commun. 5, 3747 (2014). This paper shows that cellular confinement of epithelial cells in vitro can lead to spontaneous modes of collective oscillatory movements. A simple mathematical model, in which cells are described as persistent random walkers that adapt their motion to their neighbours, with an additional term that mimics cell adhesion, captures the essential characteristics of these oscillatory modes.
Sadati, M., Qazvini, N. T., Krishnan, R., Park, C. Y. & Fredberg, J. J. Collective migration and cell jamming. Differentiation 86, 121–125 (2013).
Tambe, D. T. et al. Collective cell guidance by cooperative intercellular forces. Nat. Mater. 10, 469–475 (2011).
Bi, D. P., Lopez, J. H., Schwarz, J. M. & Manning, M. L. A density-independent rigidity transition in biological tissues. Nat. Phys. 11, 1074–1079 (2015).
Reffay, M. et al. Interplay of RhoA and mechanical forces in collective cell migration driven by leader cells. Nat. Cell Biol. 16, 217–223 (2014). In this paper, the authors analyse the mechanical role of leader cells during collective cell migration. They measure the forces exerted by finger-like structures composed of multiple cells at the migrating front and correlate the mechanical forces with RHOA activity.
Trepat, X. et al. Physical forces during collective cell migration. Nat. Phys. 5, 426–430 (2009).
du Roure, O. et al. Force mapping in epithelial cell migration. Proc. Natl Acad. Sci. USA 102, 2390–2395 (2005).
Jacinto, A., Martinez-Arias, A. & Martin, P. Mechanisms of epithelial fusion and repair. Nat. Cell Biol. 3, E117–E123 (2001).
Ravasio, A. et al. Gap geometry dictates epithelial closure efficiency. Nat. Commun. 6, 7683 (2015). To analyse epithelial gap closure, the authors experimentally implemented microscopic gaps of defined geometries and curvatures into confluent epithelial sheets. Their study clarifies the roles of two gap-closing mechanisms — cell migration and actomyosin-based contractile cable dynamics — and describes how the relative contributions of the two mechanisms are affected by gap geometry.
Nier, V. et al. Inference of internal stress in a cell monolayer. Biophys. J. 110, 1625–1635 (2016).
Hutson, M. S. et al. Forces for morphogenesis investigated with laser microsurgery and quantitative modeling. Science 300, 145–149 (2003).
Serra-Picamal, X. et al. Mechanical waves during tissue expansion. Nat. Phys. 8, 628–634 (2012).
Nelson, C. M. et al. Emergent patterns of growth controlled by multicellular form and mechanics. Proc. Natl Acad. Sci. USA 102, 11594–11599 (2005).
Warmflash, A., Sorre, B., Etoc, F., Siggia, E. D. & Brivanlou, A. H. A method to recapitulate early embryonic spatial patterning in human embryonic stem cells. Nat. Methods 11, 847–854 (2014).
Buckley, C. D. et al. The minimal cadherin-catenin complex binds to actin filaments under force. Science 346, 1254211 (2014). In this paper, the authors use single molecule assays to analyse cadherin–α-catenin complexes and their interaction with actin filaments under force. They show that tension is required to stabilize a linkage between the cadherin–catenin complex and actin filaments, and they clarify how the cadherin–catenin complex could interact directly with the actin cytoskeleton in cells.
Yonemura, S., Wada, Y., Watanabe, T., Nagafuchi, A. & Shibata, M. α-Catenin as a tension transducer that induces adherens junction development. Nat. Cell Biol. 12, 533–542 (2010).
Ladoux, B. et al. Strength dependence of cadherin-mediated adhesions. Biophys. J. 98, 534–542 (2010).
Blanchoin, L., Boujemaa-Paterski, R., Sykes, C. & Plastino, J. Actin dynamics, architecture, and mechanics in cell motility. Physiol. Rev. 94, 235–263 (2014).
Plotnikov, S. V., Pasapera, A. M., Sabass, B. & Waterman, C. M. Force fluctuations within focal adhesions mediate ECM-rigidity sensing to guide directed cell migration. Cell 151, 1513–1527 (2012).
Gupta, M. et al. Adaptive rheology and ordering of cell cytoskeleton govern matrix rigidity sensing. Nat. Commun. 6, 7525 (2015).
Gudipaty, S. A. et al. Mechanical stretch triggers rapid epithelial cell division through Piezo1. Nature 543, 118–121 (2017).
Kaliman, S., Jayachandran, C., Rehfeldt, F. & Smith, A. S. Novel growth regime of MDCK II model tissues on soft substrates. Biophys. J. 106, L25–28 (2014).
de Rooij, J., Kerstens, A., Danuser, G., Schwartz, M. A. & Waterman-Storer, C. M. Integrin-dependent actomyosin contraction regulates epithelial cell scattering. J. Cell Biol. 171, 153–164 (2005).
Mertz, A. F. et al. Cadherin-based intercellular adhesions organize epithelial cell-matrix traction forces. Proc. Natl Acad. Sci. USA 110, 842–847 (2013).
Maki, K. et al. Mechano-adaptive sensory mechanism of α-catenin under tension. Sci. Rep. 6, 24878 (2016).
Yao, M. et al. Force-dependent conformational switch of α-catenin controls vinculin binding. Nat. Commun. 5, 4525 (2014).
Barry, A. K. et al. α-Catenin cytomechanics — role in cadherin-dependent adhesion and mechanotransduction. J. Cell Sci. 127, 1779–1791 (2014).
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).
Kim, T. J. et al. Dynamic visualization of α-catenin reveals rapid, reversible conformation switching between tension states. Curr. Biol. 25, 218–224 (2015).
Mège, R. M. & Ishiyama, N. Integration of cadherin adhesion and cytoskeleton at adherens junctions. Cold Spring Harb. Perspect. Biol. 9, a028738 (2017).
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).
Bertocchi, C. et al. Nanoscale architecture of cadherin-based cell adhesions. Nat. Cell Biol. 19, 28–37 (2017).
Vasioukhin, V., Bauer, C., Yin, M. & Fuchs, E. Directed actin polymerization is the driving force for epithelial cell-cell adhesion. Cell 100, 209–219 (2000).
Twiss, F. et al. Vinculin-dependent cadherin mechanosensing regulates efficient epithelial barrier formation. Biol. Open 1, 1128–1140 (2012).
Causeret, M., Taulet, N., Comunale, F., Favard, C. & Gauthier-Rouviere, C. N-Cadherin association with lipid rafts regulates its dynamic assembly at cell-cell junctions in C2C12 myoblasts. Mol. Biol. Cell 16, 2168–2180 (2005).
le Duc, Q. et al. Vinculin potentiates E-cadherin mechanosensing and is recruited to actin-anchored sites within adherens junctions in a myosin II-dependent manner. J. Cell Biol. 189, 1107–1115 (2010).
Ladoux, B., Nelson, W. J., Yan, J. & Mège, R. M. The mechanotransduction machinery at work at adherens junctions. Integr. Biol. 1109–1119 (2015).
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).
Mason, F. M., Xie, S., Vasquez, C. G., Tworoger, M. & Martin, A. C. RhoA GTPase inhibition organizes contraction during epithelial morphogenesis. J. Cell Biol. 214, 603–617 (2016).
Martin, A. C. & Goldstein, B. Apical constriction: themes and variations on a cellular mechanism driving morphogenesis. Development 141, 1987–1998 (2014).
Acharya, B. R. & Yap, A. S. Pli selon pli: mechanochemical feedback and the morphogenetic role of contractility at cadherin cell-cell junctions. Curr. Top. Dev. Biol. 117, 631–646 (2016).
Priya, R. & Yap, A. S. Active tension: the role of cadherin adhesion and signaling in generating junctional contractility. Curr. Top. Dev. Biol. 112, 65–102 (2015).
Jodoin, J. N. et al. Stable force balance between epithelial cells arises from F-actin turnover. Dev. Cell 35, 685–697 (2015).
Begnaud, S., Chen, T., Delacour, D., Mège, R. M. & Ladoux, B. Mechanics of epithelial tissues during gap closure. Curr. Opin. Cell Biol. 42, 52–62 (2016).
Lawson, C. D. & Burridge, K. The on-off relationship of Rho and Rac during integrin-mediated adhesion and cell migration. Small GTPases 5, e27958 (2014).
Omelchenko, T. et al. β-Pix directs collective migration of anterior visceral endoderm cells in the early mouse embryo. Genes Dev. 28, 2764–2777 (2014).
Cai, D. et al. Mechanical feedback through E-cadherin promotes direction sensing during collective cell migration. Cell 157, 1146–1159 (2014). This paper illustrates the importance of E-cadherin during border cell migration in D. melanogaster , showing that E-cadherin adhesions maintain the cohesion and polarity of the group through mechanical feedback.
Hayer, A. et al. Engulfed cadherin fingers are polarized junctional structures between collectively migrating endothelial cells. Nat. Cell Biol. 18, 1311–1323 (2016). This paper shows the formation of cadherin fingers along moving endothelial cell monolayers, which are cadherin-rich protrusions that are extended from leading migrating cells and engulfed by follower cells to guide collective migration.
Dorland, Y. L. et al. The F-BAR protein pacsin2 inhibits asymmetric VE-cadherin internalization from tensile adherens junctions. Nat. Commun. 7, 12210 (2016).
Davies, P. F. Flow-mediated endothelial mechanotransduction. Physiol Rev. 75, 519–560 (1995).
Solon, J., Kaya-Copur, A., Colombelli, J. & Brunner, D. Pulsed forces timed by a ratchet-like mechanism drive directed tissue movement during dorsal closure. Cell 137, 1331–1342 (2009).
Segerer, F. J., Thuroff, F., Piera Alberola, A., Frey, E. & Radler, J. O. Emergence and persistence of collective cell migration on small circular micropatterns. Phys. Rev. Lett. 114, 228102 (2015).
Rolli, C. G. et al. Switchable adhesive substrates: revealing geometry dependence in collective cell behavior. Biomaterials 33, 2409–2418 (2012).
Friedl, P., Locker, J., Sahai, E. & Segall, J. E. Classifying collective cancer cell invasion. Nat. Cell Biol. 14, 777–783 (2012).
Doxzen, K. et al. Guidance of collective cell migration by substrate geometry. Integr. Biol. 5, 1026–1035 (2013).
Tanner, K., Mori, H., Mroue, R., Bruni-Cardoso, A. & Bissell, M. J. Coherent angular motion in the establishment of multicellular architecture of glandular tissues. Proc. Natl Acad. Sci. USA 109, 1973–1978 (2012).
Haigo, S. L. & Bilder, D. Global tissue revolutions in a morphogenetic movement controlling elongation. Science 331, 1071–1074 (2011).
Notbohm, J. et al. Cellular contraction and polarization drive collective cellular motion. Biophys. J. 110, 2729–2738 (2016).
Grasso, S., Hernández, J. A. & Chifflet, S. Roles of wound geometry, wound size, and extracellular matrix in the healing response of bovine corneal endothelial cells in culture. Am. J. Physiol. Cell Physiol. 293, C1327–C1337 (2007).
Klarlund, J. K. Dual modes of motility at the leading edge of migrating epithelial cell sheets. Proc. Natl Acad. Sci. USA 109, 15799–15804 (2012).
Rausch, S. et al. Polarizing cytoskeletal tension to induce leader cell formation during collective cell migration. Biointerphases 8, 32 (2013).
Watt, F. M. & Fujiwara, H. Cell-extracellular matrix interactions in normal and diseased skin. Cold Spring Harb. Perspect. Biol. 3, a005124 (2011).
Goodwin, K. et al. Basal cell-extracellular matrix adhesion regulates force transmission during tissue morphogenesis. Dev. Cell 39, 611–625 (2016).
Paszek, M. J. et al. Tensional homeostasis and the malignant phenotype. Cancer Cell 8, 241–254 (2005).
Pelham, R. J. & Wang, Y. L. Cell locomotion and focal adhesions are regulated by substrate flexibility. Proc. Natl Acad. Sci. USA 94, 13661–13665 (1997).
Koser, D. E. et al. Mechanosensing is critical for axon growth in the developing brain. Nat. Neurosci. 19, 1592–1598 (2016).
Reinhart-King, C. A., Dembo, M. & Hammer, D. A. Cell-cell mechanical communication through compliant substrates. Biophys. J. 95, 6044–6051 (2008).
Angelini, T. E., Hannezo, E., Trepat, X., Fredberg, J. J. & Weitz, D. A. Cell migration driven by cooperative substrate deformation patterns. Phys. Rev. Lett. (2010).
Saez, A., Buguin, A., Silberzan, P. & Ladoux, B. Is the mechanical activity of epithelial cells controlled by deformations or forces? Biophys. J. 89, L52–L54 (2005).
Saez, A., Ghibaudo, M., Buguin, A., Silberzan, P. & Ladoux, B. Rigidity-driven growth and migration of epithelial cells on microstructured anisotropic substrates. Proc. Natl Acad. Sci. USA 104, 8281–8286 (2007).
Guo, W. H., Frey, M. T., Burnham, N. A. & Wang, Y. L. Substrate rigidity regulates the formation and maintenance of tissues. Biophys. J. 90, 2213–2220 (2006).
Douezan, S., Dumond, J. & Brochard-Wyart, F. Wetting transitions of cellular aggregates induced by substrate rigidity. Soft Matter 8, 4578–4583 (2012).
Raab, M. et al. Crawling from soft to stiff matrix polarizes the cytoskeleton and phosphoregulates myosin-II heavy chain. J. Cell Biol. 199, 669–683 (2012).
Williams, C. M., Engler, A. J., Slone, R. D., Galante, L. L. & Schwarzbauer, J. E. Fibronectin expression modulates mammary epithelial cell proliferation during acinar differentiation. Cancer Res. 68, 3185–3192 (2008).
Tomasek, J. J., Gabbiani, G., Hinz, B., Chaponnier, C. & Brown, R. A. Myofibroblasts and mechano-regulation of connective tissue remodelling. Nat. Rev. Mol. Cell Biol. 3, 349–363 (2002).
Chao, C. Y. et al. In vivo and ex vivo approaches to studying the biomechanical properties of healing wounds in rat skin. J. Biomechan. Engineer. 135, 101009–101008 (2013).
Levental, K. R. et al. Matrix crosslinking forces tumor progression by enhancing integrin signaling. Cell 139, 891–906 (2009).
Odenthal, J., Takes, R. & Friedl, P. Plasticity of tumor cell invasion: governance by growth factors and cytokines. Carcinogenesis 37, 1117–1128 (2016).
Takai, Y., Miyoshi, J., Ikeda, W. & Ogita, H. Nectins and nectin-like molecules: roles in contact inhibition of cell movement and proliferation. Nat. Rev. Mol. Cell Biol. 9, 603–615 (2008).
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).
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).
Hong, S., Troyanovsky, R. B. & Troyanovsky, S. M. Binding to F-actin guides cadherin cluster assembly, stability, and movement. J. Cell Biol. 201, 131–143 (2013).
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).
Wu, Y., Vendome, J., Shapiro, L., Ben-Shaul, A. & Honig, B. Transforming binding affinities from three dimensions to two with application to cadherin clustering. Nature 475, 510–513 (2011).
Harrison, O. J. et al. The extracellular architecture of adherens junctions revealed by crystal structures of type I cadherins. Structure 19, 244–256 (2011).
Thomas, W. A. et al. α-Catenin and vinculin cooperate to promote high E-cadherin-based adhesion strength. J. Biol. Chem. 288, 4957–4969 (2013).
Niewiadomska, P., Godt, D. & Tepass, U. DE-cadherin is required for intercellular motility during Drosophila oogenesis. J. Cell Biol. 144, 533–547 (1999).
Bruce, A. E. Zebrafish epiboly: spreading thin over the yolk. Dev. Dyn. 245, 244–258 (2016).
Harrison, O. J. et al. Nectin ectodomain structures reveal a canonical adhesive interface. Nat. Struct. Mol. Biol. 19, 906–915 (2012).
Troyanovsky, R. B., Indra, I., Chen, C. S., Hong, S. & Troyanovsky, S. M. Cadherin controls nectin recruitment into adherens junctions by remodeling the actin cytoskeleton. J. Cell Sci. 128, 140–149 (2015).
Honda, T. et al. Antagonistic and agonistic effects of an extracellular fragment of nectin on formation of E-cadherin-based cell-cell adhesion. Genes Cells 8, 51–63 (2003).
Nikolic, D. L., Boettiger, A. N., Bar-Sagi, D., Carbeck, J. D. & Shvartsman, S. Y. Role of boundary conditions in an experimental model of epithelial wound healing. Am. J. Physiol. Cell Physiol. 291, C68–C75 (2006).
Murrell, M., Kamm, R. & Matsudaira, P. Tension, free space, and cell damage in a microfluidic wound healing assay. PLoS ONE 6, e24283 (2011).
van der Meer, A. D., Vermeul, K., Poot, A. A., Feijen, J. & Vermes, I. A microfluidic wound-healing assay for quantifying endothelial cell migration. Am. J. Physiol. Heart Circ. Physiol. 298, H719–H725 (2010).
Gul, I. S., Hulpiau, P., Saeys, Y. & van Roy, F. Evolution and diversity of cadherins and catenins. Exp. Cell Res. 358, 3–9 (2017).
van Roy, F. Beyond E-cadherin: roles of other cadherin superfamily members in cancer. Nat. Rev. Cancer 14, 121–134 (2014).
Plutoni, C. et al. P-Cadherin promotes collective cell migration via a Cdc42-mediated increase in mechanical forces. J. Cell Biol. 212, 199–217 (2016).
Chu, Y. S. et al. Prototypical type I E-cadherin and type II cadherin-7 mediate very distinct adhesiveness through their extracellular domains. J. Biol. Chem. 281, 2901–2910 (2006).
Labernadie, A. et al. A mechanically active heterotypic E-cadherin/N-cadherin adhesion enables fibroblasts to drive cancer cell invasion. Nat. Cell Biol. 19, 224–237 (2017). This paper shows that cancer-associated fibroblasts (CAFs) form heterotypic E-cadherin–N-cadherin contacts with tumour cells, exerting a physical force on cancer cells that enables their collective invasion.
Sawyer, J. K. et al. A contractile actomyosin network linked to adherens junctions by Canoe/afadin helps drive convergent extension. Mol. Biol. Cell 22, 2491–2508 (2011).
Sawyer, J. K., Harris, N. J., Slep, K. C., Gaul, U. & Peifer, M. The Drosophila afadin homologue Canoe regulates linkage of the actin cytoskeleton to adherens junctions during apical constriction. J. Cell Biol. 186, 57–73 (2009).
Mandai, K. et al. Afadin: a novel actin filament-binding protein with one PDZ domain localized at cadherin-based cell-to-cell adherens junction. J. Cell Biol. 139, 517–528 (1997).
Ikeda, W. et al. Afadin: a key molecule essential for structural organization of cell-cell junctions of polarized epithelia during embryogenesis. J. Cell Biol. 146, 1117–1132 (1999).
Zhadanov, A. B. et al. Absence of the tight junctional protein AF-6 disrupts epithelial cell-cell junctions and cell polarity during mouse development. Curr. Biol. 9, 880–888 (1999).
Weber, G. F., Bjerke, M. A. & DeSimone, D. W. A mechanoresponsive cadherin-keratin complex directs polarized protrusive behavior and collective cell migration. Dev. Cell 22, 104–115 (2012).
Mayer, M., Depken, M., Bois, J. S., Julicher, F. & Grill, S. W. Anisotropies in cortical tension reveal the physical basis of polarizing cortical flows. Nature 467, 617–621 (2010).
Dembo, M., Oliver, T., Ishihara, A. & Jacobson, K. Imaging the traction stresses exerted by locomoting cells with the elastic substratum method. Biophys. J. 70, 2008–2022 (1996).
Tan, J. L. et al. Cells lying on a bed of microneedles: an approach to isolate mechanical force. Proc. Natl Acad. Sci. USA 100, 1484–1489 (2003).
Yamaguchi, N., Mizutani, T., Kawabata, K. & Haga, H. Leader cells regulate collective cell migration via Rac activation in the downstream signaling of integrin β1 and PI3K. Sci. Rep. 5, 7656 (2015).
Osmani, N., Peglion, F., Chavrier, P. & Etienne-Manneville, S. Cdc42 localization and cell polarity depend on membrane traffic. J. Cell Biol. 191, 1261–1269 (2010).
Acknowledgements
The authors thank all members of the Institut Jacques Monod (IJM) and Mechanobiology Institute (MBI) teams and T. Chen, D. Delacour, P. Marcq, T. B. Saw and R. Voituriez for helpful discussions and/or critical reading of the manuscript. The authors also thank D. Pitta de Araujo, C. X. Wong and S. Wolf from MBI Science Communication Core for their substantial help with the illustrations. The authors receive financial support from the National University of Singapore/Université Sorbonne Paris Cité (NUS/USPC) and Le Projet International de Coopération Scientifique (PICS) Centre National de la Recherche Scientifique (CNRS) programmes, the European Research Council under the European Union's Seventh Framework Program (FP7/2007–2013)/European Research Council (ERC) (grant agreement number 617233), the LABEX 'Who am I?' and the Mechanobiology Institute (Singapore).
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Supplementary information
Supplementary information S1 (box)
Examples of morphogenetic processes involving collective cell migration in vivo (PDF 404 kb)
Supplementary information S2 (box)
Physical models for collective cell behaviours. (PDF 359 kb)
Supplementary information S3 (box)
Mechanocoupling by tight junctions. (PDF 170 kb)
Supplementary information S4 (box)
Mechanocoupling by desmosomes and intermediary filaments. (PDF 176 kb)
Glossary
- Border cells
-
In Drosophila melanogaster, a cluster of six to eight migratory cells migrating in the egg chamber from the follicular epithelium towards the oocyte.
- Lateral line
-
A system of sense organs found in fish, allowing the detection of movement, vibration and pressure gradients in the water.
- Traction forces
-
Forces exerted by cells on their underlying surfaces during adhesion or migration.
- Velocity fields
-
Maps of the flows of tissues by measuring the displacement field of natural tracers inside the tissues between successive images.
- Reynolds number
-
Dimensionless quantity used in hydrodynamics that represent the ratio between inertial forces and viscous forces. Laminar flows occur at low Reynolds number, whereas turbulent flows appear at high Reynolds number.
- Cell extrusion
-
An expulsion of apoptotic, non-apoptotic or transformed cells from a cell monolayer (apically or basally).
- Cryptic lamellipodia
-
Short lamellipodia forming under neighbouring cells in a migrating monolayer.
- Neural crest cells
-
In vertebrates, a group of migrating cells that arises from the border between the neural plate and the non-neural ectoderm.
- Confluence
-
In cell culture, the state in which 100% of the surface is covered by cells; also called 100% confluence.
- Cortical tension
-
The force per unit length exerted on a part of the cortex, which is a thin layer mainly composed of actin-based proteins and myosin attached to the cell membrane, by the network around it.
- Focal adhesions
-
Integrin-mediated adhesion structures formed at the cell–ECM interface, at the anchorage points of stress fibres.
- Optical tweezers
-
Highly focused laser beams that attract small objects to the centre of the beam.
- Stress
-
The force per unit area.
- Strain
-
A measure of deformation representing the length change in a body relative to a reference length.
- Shear
-
Local stresses exerted tangentially to a defined surface.
- Normal stress
-
Forces perpendicular to a defined surface, such as a cell–cell interface.
- Catch bond
-
A noncovalent bond that shows an increased lifetime with increasing amounts of tensile force applied to the bond.
- Förster resonance energy transfer
-
(FRET). When applied to optical microscopy, a method allowing the determination of the distance (or dynamic changes in the proximity) between two fluorescent molecules within several nanometres.
- Cancer-associated fibroblasts
-
(CAFs). Stromal fibroblasts closely associated with primary tumour cells and participating in the neoplastic process.
- Hippo pathway
-
Also known as the Salvador–Warts–Hippo pathway, this pathway controls organ size in animals through the regulation of cell proliferation and apoptosis.
- ERM family
-
A protein family that is named for three closely related proteins: ezrin, radixin and moesin.
- BAR domain
-
A highly conserved protein dimerization domain that occurs in many proteins involved in cellular membrane dynamics. The BAR domain is banana-shaped, binds to membranes and is capable of sensing membrane curvature by binding preferentially to curved membranes.
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Ladoux, B., Mège, RM. Mechanobiology of collective cell behaviours. Nat Rev Mol Cell Biol 18, 743–757 (2017). https://doi.org/10.1038/nrm.2017.98
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DOI: https://doi.org/10.1038/nrm.2017.98
