Development, regeneration and cancer involve drastic transitions in tissue morphology. In analogy with the behaviour of inert fluids, some of these transitions have been interpreted as wetting transitions. The validity and scope of this analogy are unclear, however, because the active cellular forces that drive tissue wetting have been neither measured nor theoretically accounted for. Here we show that the transition between two-dimensional epithelial monolayers and three-dimensional spheroidal aggregates can be understood as an active wetting transition whose physics differs fundamentally from that of passive wetting phenomena. By combining an active polar fluid model with measurements of physical forces as a function of tissue size, contractility, cell–cell and cell–substrate adhesion, and substrate stiffness, we show that the wetting transition results from the competition between traction forces and contractile intercellular stresses. This competition defines a new intrinsic length scale that gives rise to a critical size for the wetting transition in tissues, a striking feature that has no counterpart in classical wetting. Finally, we show that active shape fluctuations are dynamically amplified during tissue dewetting. Overall, we conclude that tissue spreading constitutes a prominent example of active wetting—a novel physical scenario that may explain morphological transitions during tissue morphogenesis and tumour progression.
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Gonzalez-Rodriguez, D., Guevorkian, K., Douezan, S. & Brochard-Wyart, F. Soft matter models of developing tissues and tumors. Science 338, 910–917 (2012).
Friedl, P. & Gilmour, D. Collective cell migration in morphogenesis, regeneration and cancer. Nat. Rev. Mol. Cell Biol. 10, 445–457 (2009).
Jülicher, F. & Eaton, S. Emergence of tissue shape changes from collective cell behaviours. Semin. Cell. Dev. Biol. 67, 103–112 (2017).
Ryan, P. L., Foty, R. A., Kohn, J. & Steinberg, M. S. Tissue spreading on implantable substrates is a competitive outcome of cell–cell vs. cell–substratum adhesivity. Proc. Natl Acad. Sci. USA 98, 4323–4327 (2001).
Douezan, S. et al. Spreading dynamics and wetting transition of cellular aggregates. Proc. Natl Acad. Sci. USA 108, 7315–7320 (2011).
Douezan, S., Dumond, J. & Brochard-Wyart, F. Wetting transitions of cellular aggregates induced by substrate rigidity. Soft Matter 8, 4578–4583 (2012).
Beaune, G. et al. How cells flow in the spreading of cellular aggregates. Proc. Natl Acad. Sci. USA 111, 8055–8060 (2014).
Beaune, G. et al. Reentrant wetting transition in the spreading of cellular aggregates. Soft Matter 13, 8474–8482 (2017).
Behrndt, M. et al. Forces driving epithelial spreading in zebrafish gastrulation. Science 338, 257–260 (2012).
Campinho, P. et al. Tension-oriented cell divisions limit anisotropic tissue tension in epithelial spreading during zebrafish epiboly. Nat. Cell Biol. 15, 1405–1414 (2013).
Morita, H. et al. The physical basis of coordinated tissue spreading in zebrafish gastrulation. Dev. Cell. 40, 354–366.e4 (2017).
Wallmeyer, B., Trinschek, S., Yigit, S., Thiele, U. & Betz, T. Collective cell migration in embryogenesis follows the laws of wetting. Biophys. J. 114, 213–222 (2018).
Douezan, S. & Brochard-Wyart, F. Dewetting of cellular monolayers. Eur. Phys. J. E 35, 34 (2012).
Smeets, B. et al. Emergent structures and dynamics of cell colonies by contact inhibition of locomotion. Proc. Natl Acad. Sci. USA 113, 14621–14626 (2016).
Ravasio, A. et al. Regulation of epithelial cell organization by tuning cell–substrate adhesion. Integr. Biol. 7, 1228–1241 (2015).
Sarrió, D. et al. Functional characterization of E- and P-cadherin in invasive breast cancer cells. BMC. Cancer 9, 74 (2009).
Trepat, X. et al. Physical forces during collective cell migration. Nat. Phys. 5, 426–430 (2009).
Serra-Picamal, X. et al. Mechanical waves during tissue expansion. Nat. Phys. 8, 628–634 (2012).
Tambe, D. T. et al. Collective cell guidance by cooperative intercellular forces. Nat. Mater. 10, 469–475 (2011).
Mertz, A. F. et al. Cadherin-based intercellular adhesions organize epithelial cell–matrix traction forces. Proc. Natl Acad. Sci. USA 110, 842–847 (2013).
Mertz, A. F. et al. Scaling of traction forces with the size of cohesive cell colonies. Phys. Rev. Lett. 108, 198101 (2012).
Harris, A. R., Daeden, A. & Charras, G. T. Formation of adherens junctions leads to the emergence of a tissue-level tension in epithelial monolayers. J. Cell. Sci. 127, 2507–2517 (2014).
Bazellières, E. et al. Control of cell–cell forces and collective cell dynamics by the intercellular adhesome. Nat. Cell Biol. 17, 409–420 (2015).
Lecuit, T. & Yap, A. S. E-cadherin junctions as active mechanical integrators in tissue dynamics. Nat. Cell Biol. 17, 533–539 (2015).
Muhamed, I. et al. E-cadherin-mediated force transduction signals regulate global cell mechanics. J. Cell. Sci. 129, 1843–1854 (2016).
Blanch-Mercader, C. et al. Effective viscosity and dynamics of spreading epithelia: a solvable model. Soft Matter 13, 1235–1243 (2017).
Köpf, M. H. & Pismen, L. M. A continuum model of epithelial spreading. Soft Matter 9, 3727–3734 (2013).
Banerjee, S., Utuje, K. J. C. & Marchetti, M. C. Propagating stress waves during epithelial expansion. Phys. Rev. Lett. 114, 228101 (2015).
Notbohm, J. et al. Cellular contraction and polarization drive collective cellular motion. Biophys. J. 110, 2729–2738 (2016).
Lee, P. & Wolgemuth, C. W. Crawling cells can close wounds without purse strings or signaling. PLoS Comput. Biol. 7, e1002007 (2011).
Lee, P. & Wolgemuth, C. Advent of complex flows in epithelial tissues. Phys. Rev. E 83, 061920 (2011).
Vig, D. K., Hamby, A. E. & Wolgemuth, C. W. Cellular contraction can drive rapid epithelial flows. Biophys. J. 113, 1613–1622 (2017).
Saw, T. B. et al. Topological defects in epithelia govern cell death and extrusion. Nature 544, 212–216 (2017).
Blanch-Mercader, C. & Casademunt, J. Hydrodynamic instabilities, waves and turbulence in spreading epithelia. Soft Matter 13, 6913–6928 (2017).
Yabunaka, S. & Marcq, P. Emergence of epithelial cell density waves. Soft Matter 13, 7046–7052 (2017).
Kruse, K., Joanny, J. F., Jülicher, F., Prost, J. & Sekimoto, K. Generic theory of active polar gels: a paradigm for cytoskeletal dynamics. Eur. Phys. J. E 16, 5–16 (2005).
Jülicher, F. in New Trends in the Physics and Mechanics of Biological Systems (eds. Ben Amar, M. et al) Ch. 4 (Oxford Univ. Press, Oxford, 2011).
Marchetti, M. C. et al. Hydrodynamics of soft active matter. Rev. Mod. Phys. 85, 1143–1189 (2013).
Prost, J., Jülicher, F. & Joanny, J.-F. Active gel physics. Nat. Phys. 11, 111–117 (2015).
de Gennes, P.-G. & Prost, J. The Physics of Liquid Crystals (Oxford Univ. Press, Oxford, 1993).
Oriola, D., Alert, R. & Casademunt, J. Fluidization and active thinning by molecular kinetics in active gels. Phys. Rev. Lett. 118, 088002 (2017).
Deforet, M., Hakim, V., Yevick, H. G., Duclos, G. & Silberzan, P. Emergence of collective modes and tri-dimensional structures from epithelial confinement. Nat. Commun. 5, 3747 (2014).
Kaliman, S., Jayachandran, C., Rehfeldt, F. & Smith, A.-S. Novel growth regime of MDCK II model tissues on soft substrates. Biophys. J. 106, L25–L28 (2014).
Schwarz, U. S. & Safran, S. A. Physics of adherent cells. Rev. Mod. Phys. 85, 1327–1381 (2013).
Edwards, A. M. J., Ledesma-Aguilar, R., Newton, M. I., Brown, C. V. & McHale, G. Not spreading in reverse: The dewetting of a liquid film into a single drop. Sci. Adv. 2, e1600183 (2016).
de Gennes, P. Wetting: statics and dynamics. Rev. Mod. Phys. 57, 827–863 (1985).
Bonn, D., Eggers, J., Indekeu, J., Meunier, J. & Rolley, E. Wetting and spreading. Rev. Mod. Phys. 81, 739–805 (2009).
Chepizhko, O. et al. Bursts of activity in collective cell migration. Proc. Natl Acad. Sci. USA 113, 11408–11413 (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).
Stirbat, T. V. et al. Fine tuning of tissues’ viscosity and surface tension through contractility suggests a new role for α-Catenin. PLoS One 8, e52554 (2013).
Rodriguez-Hernandez, I., Cantelli, G., Bruce, F. & Sanz-Moreno, V. Rho, ROCK and actomyosin contractility in metastasis as drug targets. F1000Res. 5, 783 (2016).
Ouderkirk, J. L. & Krendel, M. Non-muscle myosins in tumor progression, cancer cell invasion, and metastasis. Cytoskeleton. 71, 447–463 (2014).
Paredes, J. et al. Epithelial E- and P-cadherins: Role and clinical significance in cancer. Biochim. Biophys. Acta 1826, 297–311 (2012).
Paschos, K. A., Canovas, D. & Bird, N. C. The role of cell adhesion molecules in the progression of colorectal cancer and the development of liver metastasis. Cell. Signal. 21, 665–674 (2009).
Clark, A. G. & Vignjevic, D. M. Modes of cancer cell invasion and the role of the microenvironment. Curr. Opin. Cell Biol. 36, 13–22 (2015).
Lu, P., Weaver, V. M. & Werb, Z. The extracellular matrix: A dynamic niche in cancer progression. J. Cell. Biol. 196, 395–406 (2012).
Cortina, C. et al. EphB–ephrin-B interactions suppress colorectal cancer progression by compartmentalizing tumor cells. Nat. Genet. 39, 1376–1383 (2007).
Casares, L. et al. Hydraulic fracture during epithelial stretching. Nat. Mater. 14, 343–351 (2015).
Tambe, D. T. et al. Monolayer stress microscopy: Limitations, artifacts, and accuracy of recovered intercellular stresses. PLoS One 8, e55172 (2013).
We thank D. Sarrió and G. Moreno-Bueno for providing the E-cadherin inducible cells; N. Castro for technical assistance; A. Elosegui, V. González, E. Latorre, L. Valon and R. Vincent for stimulating discussions. R.A. thanks G. Torrents for assistance with mathematical details. C.P-G. and R.A. were funded by Fundació ‘La Caixa’. R.A. thanks J. Prost and acknowledges EMBO (Short Term Fellowship ASTF 365-2015), The Company of Biologists (Development Travelling Fellowship DEVTF-151206), and Fundació Universitària Agustí Pedro i Pons for supporting visits to Institut Curie. This work was supported by the Spanish Ministry of Economy and Competitiveness/FEDER (BFU2015-65074-P to X.T., FIS2016-78507-C2-2-P to J.C.), the Generalitat de Catalunya (2014-SGR-927 and CERCA Program to X.T., 2014-SGR-878 to J.C.), the European Research Council (CoG-616480 to X.T.), European Commission (H2020-FETPROACT-01-2016-731957 to X.T.) and Obra Social ‘La Caixa’. IBEC is recipient of a Severo Ochoa Award of Excellence from the MINECO.
The authors declare no competing interests.
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Supplementary Note, Supplementary Figures 1–16, Supplementary References 1–45
Unconfined monolayer exhibiting a transition from wetting to dewetting.Representative example of a spreading monolayer (shown in Fig. 1f) undergoing a wetting transition.The release of confinement at t = 0 h allows the monolayer to freely spread. At ~25 h, the monolayerspontaneously starts retracting until it collapses into a spheroidal aggregate.
Another example of a wetting transition in a spreading monolayer. Another spreading monolayer (shown in Supplementary Fig. 2) undergoing a wetting transition.
Evolution of traction and tension fields during wetting and dewetting. Videos of phase contrast images (left), maps of traction (centre) and monolayer tension (right) in a monolayer with increasing concentration of E-cadherin. A wetting transition is observed at time t = 22 h.
Orthogonal views of monolayer dewetting. Timelapse of MDA-MB-231 cells stably expressing a cell membrane marker (CAAX-iRFP). The tissue-substrate contact area decreases pronouncedly during dewetting, while the tissue evolves from a monolayer to a spheroidal cell aggregate, resembling a droplet.
Calcium chelation hinders the increase of tissue forces and prevents dewetting. Phase contrast, and maps of traction forces and monolayer tension of control (left) and EGTA-treated (right)cell islands. Cells treated with EGTA move individually rather than forming a cohesive monolayer, suggesting that cell–cell junctions are efficiently abrogated. In the presence of EGTA, both tractions and monolayer tension increase much more slowly than in control islands, and the wetting transition does not occur.
Dewetting is inhibited and reversed when tissue contractility is externally decreased. Dewetting (left), dewetting inhibition (centre) and reversibility (right) assays. Partial inhibition of contractility with blebbistatin clearly delays the wetting transition. A sudden inhibition of contractility with Y27632 (t = 46 h) is enough to revert dewetting, inducing a rewetting of the substrate. The name of the drug indicates its presence in the cell medium.
Cell rearrangements in the monolayer. Phase contrast (left) and cell nuclei (right) in a 200 µm radius island during the wetting phase of the experiment. Cells incessantly exchange neighbours, a fact that provides support to the fluid behaviour of the monolayer. Moreover, cells progressively accumulate at the edge of the monolayer, which develops a gentle cell density gradient.
Evolution of traction and monolayer tension fields in islands of different radii. For all sizes, the magnitude of tractions and monolayer tension increase in time as E-cadherin is progressively expressed. Tractions accumulate at the edges of the monolayers, while monolayer tension has a maximum at the centre. Red frames indicate monolayer dewetting.
Evolution of traction and monolayer tension fields in islands on substrates of different stiffnesses. For monolayer on substrates of Young’s modulus 3 and 12 kPa, tissue forces increase in time, eventually triggering monolayer dewetting. This transition occurs earlier for the softest substrate. For the stiffest substrate (30 kPa), tissue forces keep increasing until the end of the experiment, suggesting that the critical contractility to induce dewetting is not reached. Red frames indicate monolayer dewetting.
The wetting transition time depends on tissue radius and substrate ligand density.Cell islands of different radii seeded on substrates with different substrate ligand densities exhibit the wetting transition at different times. Red frames indicate monolayer dewetting.
Symmetry breaking of monolayer shape during dewetting. A 200 µm radius cell island divided in 24 sectors. Blue = wetting, red = dewetting. Dewetting starts in diametrically opposed regions of the monolayer edge. Hence, the monolayer loses its initial circular shape and acquires an elliptic-like shape during the early stages of dewetting.
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Pérez-González, C., Alert, R., Blanch-Mercader, C. et al. Active wetting of epithelial tissues. Nature Phys 15, 79–88 (2019). https://doi.org/10.1038/s41567-018-0279-5
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