Abstract
How the shape of embryos and organs emerges during development is a fundamental question that has fascinated scientists for centuries. Tissue dynamics arise from a small set of cell behaviours, including shape changes, cell contact remodelling, cell migration, cell division and cell extrusion. These behaviours require control over cell mechanics, namely active stresses associated with protrusive, contractile and adhesive forces, and hydrostatic pressure, as well as material properties of cells that dictate how cells respond to active stresses. In this Review, we address how cell mechanics and the associated cell behaviours are robustly organized in space and time during tissue morphogenesis. We first outline how not only gene expression and the resulting biochemical cues, but also mechanics and geometry act as sources of morphogenetic information to ultimately define the time and length scales of the cell behaviours driving morphogenesis. Next, we present two idealized modes of how this information flows — how it is read out and translated into a biological effect — during morphogenesis. The first, akin to a programme, follows deterministic rules and is hierarchical. The second follows the principles of self-organization, which rests on statistical rules characterizing the system’s composition and configuration, local interactions and feedback. We discuss the contribution of these two modes to the mechanisms of four very general classes of tissue deformation, namely tissue folding and invagination, tissue flow and extension, tissue hollowing and, finally, tissue branching. Overall, we suggest a conceptual framework for understanding morphogenetic information that encapsulates genetics and biochemistry as well as mechanics and geometry as information modules, and the interplay of deterministic and self-organized mechanisms of their deployment, thereby diverging considerably from the traditional notion that shape is fully encoded and determined by genes.
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References
Waddington, C. H. The Strategy of the Genes: a Discussion of Some Aspects of Theoretical Biology (George Allen and Unwin, London, 1957).
Slack, J. M. Conrad Hal Waddington: the last renaissance biologist? Nat. Rev. Genet. 3, 889–895 (2002).
Roux, W. in Foundations of Experimental Embryology (eds Willier, B. H. & Oppenheimer, J. M.) 2–37 (Hafner, 1888).
De Robertis, E. M. Spemann’s organizer and self-regulation in amphibian embryos. Nat. Rev. Mol. Cell Biol. 7, 296–302 (2006).
Sulston, J. E., Schierenberg, E., White, J. G. & Thomson, J. N. The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev. Biol. 100, 64–119 (1983).
Maduro, M. F. Cell fate specification in the C. elegans embryo. Dev. Dyn. 239, 1315–1329 (2010).
Nishida, H. & Stach, T. Cell lineages and fate maps in tunicates: conservation and modification. Zool. Sci. 31, 645–652 (2014).
Garcia-Bellido, A. & Santamaria, P. Developmental analysis of the wing disc in the mutant engrailed of Drosophila melanogaster. Genetics 72, 87–104 (1972).
Pradel, J. & White, R. A. From selectors to realizators. Int. J. Dev. Biol. 42, 417–421 (1998).
Halder, G., Callaerts, P. & Gehring, W. J. Induction of ectopic eyes by targeted expression of the eyeless gene in Drosophila. Science 267, 1788–1792 (1995).
Halder, G., Callaerts, P. & Gehring, W. J. New perspectives on eye evolution. Curr. Opin. Genet. Dev. 5, 602–609 (1995).
Chanut-Delalande, H., Fernandes, I., Roch, F., Payre, F. & Plaza, S. Shavenbaby couples patterning to epidermal cell shape control. PLoS Biol. 4, e290 (2006).
Driesch, H. The potency of the first two cleavage cells in echinoderm development: experimental production of double and partial formation. Reprinted in Foundations of Experimental Embryology (eds Willier, B. H. & Oppenheimer, J. M.) (Hafner, 1892).
Morgan, T. H. Half-embryos and whole-embryos from one of the first two blastomeres of the frog’s egg. Anat. Anz. 10, 623–628 (1895).
Spemann, H. Embryonic Development and Induction, Vol. 10 (Taylor & Francis, 1988).
Browne, E. N. The production of new hydranths in Hydra by the insertion of small grafts. J. Exp. Zool. 7, 1–23 (1909).
Spemann, H. & Mangold, H. Induction of embryonic primordia by implantation of organizers from a different species. Roux Arch. Entwickl. Mech. 100, 599–638 (1924).
Gilmour, D., Rembold, M. & Leptin, M. From morphogen to morphogenesis and back. Nature 541, 311–320 (2017).
Hannezo, E. & Heisenberg, C. P. Mechanochemical feedback loops in development and disease. Cell 178, 12–25 (2019).
Schweisguth, F. & Corson, F. Self-organization in pattern formation. Dev. Cell 49, 659–677 (2019).
Turing, A. M. The chemical basis of morphogenesis. Philos. Trans. R. Soc. London. Ser. B, Biol. Sci. 237, 37–72 (1952).
Prusinkiewicz, P., Meinhardt, H. & Fowler, D. R. The Algorithmic Beauty of Sea Shells (Springer, 2003).
Gelens, L., Anderson, G. A. & Ferrell, J. E. Jr. Spatial trigger waves: positive feedback gets you a long way. Mol. Biol. Cell 25, 3486–3493 (2014).
Wartlick, O. et al. Dynamics of Dpp signaling and proliferation control. Science 331, 1154–1159 (2011).
Rogers, K. W. & Schier, A. F. Morphogen gradients: from generation to interpretation. Annu. Rev. Cell Dev. Biol. 27, 377–407 (2011).
Sagner, A. & Briscoe, J. Morphogen interpretation: concentration, time, competence, and signaling dynamics. Wiley Interdiscip. Rev. Dev. Biol. 6, e271 (2017).
Economou, A. D. et al. Periodic stripe formation by a Turing mechanism operating at growth zones in the mammalian palate. Nat. Genet. 44, 348–351 (2012).
Sheth, R. et al. Hox genes regulate digit patterning by controlling the wavelength of a Turing-type mechanism. Science 338, 1476–1480 (2012).
Pourquie, O. The segmentation clock: converting embryonic time into spatial pattern. Science 301, 328–330 (2003).
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).
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 e3134 (2017).
Dasbiswas, K., Hannezo, E. & Gov, N. S. Theory of epithelial cell shape transitions induced by mechanoactive chemical gradients. Biophys. J. 114, 968–977 (2018).
Dicko, M. et al. Geometry can provide long-range mechanical guidance for embryogenesis. PLoS Comput. Biol. 13, e1005443 (2017).
Oster, G. F., Murray, J. D. & Harris, A. K. Mechanical aspects of mesenchymal morphogenesis. J. Embryol. Exp. Morphol. 78, 83–125 (1983).
Shyer, A. E. et al. Emergent cellular self-organization and mechanosensation initiate follicle pattern in the avian skin. Science 357, 811–815 (2017).
Hannezo, E., Dong, B., Recho, P., Joanny, J. F. & Hayashi, S. Cortical instability drives periodic supracellular actin pattern formation in epithelial tubes. Proc. Natl Acad. Sci. USA 112, 8620–8625 (2015).
Ladoux, B., Nelson, W. J., Yan, J. & Mège, R. M. The mechanotransduction machinery at work at adherens junctions. Integr. Biol. 7, 1109–1119 (2015).
Collinet, C. & Lecuit, T. Stability and dynamics of cell-cell junctions. Prog. Mol. Biol. Transl Sci. 116, 25–47 (2013).
Papusheva, E. & Heisenberg, C.-P. Spatial organization of adhesion: force-dependent regulation and function in tissue morphogenesis. EMBO J. 29, 2753–2768 (2010).
Schwayer, C. et al. Mechanosensation of tight junctions depends on ZO-1 phase separation and flow. Cell 179, 937–952 e918 (2019).
Lecuit, T., Lenne, P. F. & Munro, E. Force generation, transmission, and integration during cell and tissue morphogenesis. Annu. Rev. Cell Dev. Biol. 27, 157–184 (2011).
Geiger, B., Spatz, J. P. & Bershadsky, A. D. Environmental sensing through focal adhesions. Nat. Rev. Mol. Cell Biol. 10, 21–33 (2009).
Bailles, A. et al. Genetic induction and mechanochemical propagation of a morphogenetic wave. Nature 572, 467–473 (2019). Shows that a mechano-chemical wave of Rho1–MyoII activation drives invagination and anterior movement of the posterior endoderm during D. melanogaster gastrulation. It shows the relevance of self-organized contractile waves in driving tissue invagination and movements.
Kindberg, A., Hu, J. K. & Bush, J. O. Forced to communicate: integration of mechanical and biochemical signaling in morphogenesis. Curr. Opin. Cell Biol. 66, 59–68 (2020).
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). Shows how pulsed contractility during D. melanogaster germband extension depends on self-organizational properties of actomyosin networks, that is, an advection-mediated positive feedback on MyoII activation due to the association of Rho1GTP, Rok to actomyosin cortex.
Goehring, N. W. et al. Polarization of PAR proteins by advective triggering of a pattern-forming system. Science 334, 1137–1141 (2011).
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).
Durdu, S. et al. Luminal signalling links cell communication to tissue architecture during organogenesis. Nature 515, 120–124 (2014).
Shyer, A. E., Huycke, T. R., Lee, C., Mahadevan, L. & Tabin, C. J. Bending gradients: how the intestinal stem cell gets its home. Cell 161, 569–580 (2015). Shows that mechanical buckling of intestinal villi in chick and mouse embryos distorts a chemical gradient of SHH and restricts stem cell proliferation at the villus base. It shows the relevance of tissue geometry in shaping the morphogenetic information.
Martin, A. C. & Goldstein, B. Apical constriction: themes and variations on a cellular mechanism driving morphogenesis. Development 141, 1987–1998 (2014).
Munjal, A. & Lecuit, T. Actomyosin networks and tissue morphogenesis. Development 141, 1789–1793 (2014).
Leptin, M. & Grunewald, B. Cell shape changes during gastrulation in Drosophila. Development 110, 73–84 (1990).
Sweeton, D., Parks, S., Costa, M. & Wieschaus, E. Gastrulation in Drosophila: the formation of the ventral furrow and posterior midgut invaginations. Development 112, 775–789 (1991).
Martin, A. C., Kaschube, M. & Wieschaus, E. F. Pulsed contractions of an actin–myosin network drive apical constriction. Nature 457, 495–499 (2009). Shows that cell apical constriction during invagination of the mesoderm in D. melanogaster is driven by medio-apical actomyosin pulsed contractions instead of purse-string contractility at adherens junctions as previously thought.
Gelbart, M. A. et al. Volume conservation principle involved in cell lengthening and nucleus movement during tissue morphogenesis. Proc. Natl Acad. Sci. USA 109, 19298–19303 (2012).
Krueger, D., Tardivo, P., Nguyen, C. & De Renzis, S. Downregulation of basal myosin-II is required for cell shape changes and tissue invagination. EMBO J. 37, e100170, https://doi.org/10.15252/embj.2018100170 (2018).
Gracia, M. et al. Mechanical impact of epithelial-mesenchymal transition on epithelial morphogenesis in Drosophila. Nat. Commun. 10, 2951 (2019).
Sherrard, K., Robin, F., Lemaire, P. & Munro, E. Sequential activation of apical and basolateral contractility drives ascidian endoderm invagination. Curr. Biol. 20, 1499–1510 (2010). Identifies a two-step mechanism for endoderm invagination in ascidian embryos: first, a Rho1–ROCK–MyoII-mediated phase of cell apical constriction and second, a phase of apico-basal shortening dependent on MyoII but not on Rho1–ROCK. It shows the relevance of polarized actomyosin contractility in driving 3D cell shape changes during invagination.
Mason, F. M., Tworoger, M. & Martin, A. C. Apical ___domain polarization localizes actin-myosin activity to drive ratchet-like apical constriction. Nat. Cell Biol. 15, 926–936 (2013).
Vasquez, C. G., Tworoger, M. & Martin, A. C. Dynamic myosin phosphorylation regulates contractile pulses and tissue integrity during epithelial morphogenesis. J. Cell Biol. 206, 435–450 (2014).
Izquierdo, E., Quinkler, T. & De Renzis, S. Guided morphogenesis through optogenetic activation of Rho signalling during early Drosophila embryogenesis. Nat. Commun. 9, 2366 (2018).
Reeves, G. T. & Stathopoulos, A. Graded dorsal and differential gene regulation in the Drosophila embryo. Cold Spring Harb. Perspect. Biol. 1, a000836 (2009).
Manning, A. J., Peters, K. A., Peifer, M. & Rogers, S. L. Regulation of epithelial morphogenesis by the G protein-coupled receptor mist and its ligand fog. Sci. Signal. 6, ra98 (2013).
Costa, M., Wilson, E. T. & Wieschaus, E. A putative cell signal encoded by the folded gastrulation gene coordinates cell shape changes during Drosophila gastrulation. Cell 76, 1075–1089 (1994).
Kolsch, V., Seher, T., Fernandez-Ballester, G. J., Serrano, L. & Leptin, M. Control of Drosophila gastrulation by apical localization of adherens junctions and RhoGEF2. Science 315, 384–386 (2007).
Lee, J. Y. et al. Wnt/Frizzled signaling controls C. elegans gastrulation by activating actomyosin contractility. Curr. Biol. 16, 1986–1997 (2006).
Roh-Johnson, M. et al. Triggering a cell shape change by exploiting preexisting actomyosin contractions. Science 335, 1232–1235 (2012).
Marston, D. J. et al. MRCK-1 drives apical constriction in C. elegans by linking developmental patterning to force generation. Curr. Biol. 26, 2079–2089 (2016).
Nishimura, T. & Takeichi, M. Shroom3-mediated recruitment of Rho kinases to the apical cell junctions regulates epithelial and neuroepithelial planar remodeling. Development 135, 1493–1502 (2008).
Haigo, S. L., Hildebrand, J. D., Harland, R. M. & Wallingford, J. B. Shroom induces apical constriction and is required for hingepoint formation during neural tube closure. Curr. Biol. 13, 2125–2137 (2003).
Chung, M. I., Nascone-Yoder, N. M., Grover, S. A., Drysdale, T. A. & Wallingford, J. B. Direct activation of Shroom3 transcription by Pitx proteins drives epithelial morphogenesis in the developing gut. Development 137, 1339–1349 (2010).
Plageman, T. F. Jr. et al. Pax6-dependent Shroom3 expression regulates apical constriction during lens placode invagination. Development 137, 405–415 (2010).
Plageman, T. F. Jr. et al. A Trio-RhoA-Shroom3 pathway is required for apical constriction and epithelial invagination. Development 138, 5177–5188 (2011).
Ernst, S. et al. Shroom3 is required downstream of FGF signalling to mediate proneuromast assembly in zebrafish. Development 139, 4571–4581 (2012).
Hildebrand, J. D. Shroom regulates epithelial cell shape via the apical positioning of an actomyosin network. J. Cell Sci. 118, 5191–5203 (2005).
Lang, R. A., Herman, K., Reynolds, A. B., Hildebrand, J. D. & Plageman, T. F. Jr. p120-catenin-dependent junctional recruitment of Shroom3 is required for apical constriction during lens pit morphogenesis. Development 141, 3177–3187 (2014).
Rauzi, M., Lenne, P. F. & Lecuit, T. Planar polarized actomyosin contractile flows control epithelial junction remodelling. Nature 468, 1110–1114 (2010).
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).
Munro, E., Nance, J. & Priess, J. R. Cortical flows powered by asymmetrical contraction transport PAR proteins to establish and maintain anterior-posterior polarity in the early C. elegans embryo. Dev. Cell 7, 413–424 (2004).
Kim, H. Y. & Davidson, L. A. Punctuated actin contractions during convergent extension and their permissive regulation by the non-canonical Wnt-signaling pathway. J. Cell Sci. 124, 635–646 (2011).
Maitre, J. L., Niwayama, R., Turlier, H., Nedelec, F. & Hiiragi, T. Pulsatile cell-autonomous contractility drives compaction in the mouse embryo. Nat. Cell Biol. 17, 849–855 (2015).
Xie, S. & Martin, A. C. Intracellular signalling and intercellular coupling coordinate heterogeneous contractile events to facilitate tissue folding. Nat. Commun. 6, 7161 (2015).
Yevick, H. G., Miller, P. W., Dunkel, J. & Martin, A. C. Structural redundancy in supracellular actomyosin networks enables robust tissue folding. Dev. Cell 50, 586–598 e583 (2019).
Bhide, S. et al. Mechanical competition alters the cellular interpretation of an endogenous genetic programme. Preprint at bioRxiv https://doi.org/10.1101/2020.10.15.333963 (2020).
Odell, G. M., Oster, G., Alberch, P. & Burnside, B. The mechanical basis of morphogenesis. I. Epithelial folding and invagination. Dev. Biol. 85, 446–462 (1981).
Llinares-Benadero, C. & Borrell, V. Deconstructing cortical folding: genetic, cellular and mechanical determinants. Nat. Rev. Neurosci. 20, 161–176 (2019).
Garcia, K. E., Kroenke, C. D. & Bayly, P. V. Mechanics of cortical folding: stress, growth and stability. Philos. Trans. R. Soc. Lond. B Biol. Sci. 373, 20170321 (2018).
Huycke, T. R. & Tabin, C. J. Chick midgut morphogenesis. Int. J. Dev. Biol. 62, 109–119 (2018).
Walton, K. D., Mishkind, D., Riddle, M. R., Tabin, C. J. & Gumucio, D. L. Blueprint for an intestinal villus: species-specific assembly required. Wiley Interdiscip. Rev. Dev. Biol. 7, e317 (2018).
Vasung, L. et al. Quantitative and qualitative analysis of transient fetal compartments during prenatal human brain development. Front. Neuroanat. 10, 11 (2016).
Neal, J. et al. Insights into the gyrification of developing ferret brain by magnetic resonance imaging. J. Anat. 210, 66–77 (2007).
Lohmann, G., von Cramon, D. Y. & Colchester, A. C. Deep sulcal landmarks provide an organizing framework for human cortical folding. Cereb. Cortex 18, 1415–1420 (2008).
Ono, M., Kubik, S. & Abernathey, C. D. Atlas of the Cerebral Sulci (G. Thieme Verlag, 1990).
Mota, B. & Herculano-Houzel, S. Cortical folding scales universally with surface area and thickness, not number of neurons. Science 349, 74–77 (2015).
Van Essen, D. C. A tension-based theory of morphogenesis and compact wiring in the central nervous system. Nature 385, 313–318 (1997).
Lefevre, J. & Mangin, J. F. A reaction-diffusion model of human brain development. PLoS Comput. Biol. 6, e1000749 (2010).
Richman, D. P., Stewart, R. M., Hutchinson, J. W. & Caviness, V. S. Jr. Mechanical model of brain convolutional development. Science 189, 18–21 (1975).
Hannezo, E., Prost, J. & Joanny, J. F. Instabilities of monolayered epithelia: shape and structure of villi and crypts. Phys. Rev. Lett. 107, 078104 (2011).
Ben Amar, M. & Jia, F. Anisotropic growth shapes intestinal tissues during embryogenesis. Proc. Natl Acad. Sci. USA 110, 10525–10530 (2013).
Mahadevan, L. & Rica, S. Self-organized origami. Science 307, 1740 (2005).
Xu, G. et al. Axons pull on the brain, but tension does not drive cortical folding. J. Biomech. Eng. 132, 071013 (2010).
Tallinen, T., Chung, J. Y., Biggins, J. S. & Mahadevan, L. Gyrification from constrained cortical expansion. Proc. Natl Acad. Sci. USA 111, 12667–12672 (2014).
Tallinen, T. et al. On the growth and form of cortical convolutions. Nat. Phys. 12, 588–593 (2016). Shows that the initial size and geometry of the structure coupled with mechanical instability due to differential tangential growth accounts for the cortical folding pattern observed in fetal human brains. It shows the importance of the initial geometry in positioning folds.
Kriegstein, A., Noctor, S. & Martinez-Cerdeno, V. Patterns of neural stem and progenitor cell division may underlie evolutionary cortical expansion. Nat. Rev. Neurosci. 7, 883–890 (2006).
Smart, I. H., Dehay, C., Giroud, P., Berland, M. & Kennedy, H. Unique morphological features of the proliferative zones and postmitotic compartments of the neural epithelium giving rise to striate and extrastriate cortex in the monkey. Cereb. Cortex 12, 37–53 (2002).
Reillo, I., de Juan Romero, C., Garcia-Cabezas, M. A. & Borrell, V. A role for intermediate radial glia in the tangential expansion of the mammalian cerebral cortex. Cereb. Cortex 21, 1674–1694 (2011).
de Juan Romero, C., Bruder, C., Tomasello, U., Sanz-Anquela, J. M. & Borrell, V. Discrete domains of gene expression in germinal layers distinguish the development of gyrencephaly. EMBO J. 34, 1859–1874 (2015).
Savin, T. et al. On the growth and form of the gut. Nature 476, 57–62 (2011). Shows that gut looping arises from growth differences between the gut tube and the anchoring dorsal mesenteric sheet. It documents that equilibrium linear elasticity theory accounts for complex tissue geometry in vivo.
Lyons, K. M., Pelton, R. W. & Hogan, B. L. Organogenesis and pattern formation in the mouse: RNA distribution patterns suggest a role for bone morphogenetic protein-2A (BMP-2A). Development 109, 833–844 (1990).
Nerurkar, N. L., Mahadevan, L. & Tabin, C. J. BMP signaling controls buckling forces to modulate looping morphogenesis of the gut. Proc. Natl Acad. Sci. USA 114, 2277–2282 (2017).
Coulombre, A. J. & Coulombre, J. L. Intestinal development. I. Morphogenesis of the villi and musculature. J. Embryol. Exp. Morphol. 6, 403–411 (1958).
Shyer, A. E. et al. Villification: how the gut gets its villi. Science 342, 212–218 (2013). Shows that the different steps of avian gut villification result from a mechanical instability due to the expansion of the growing endoderm and the constraints imposed by surrounding muscle layers.
Grey, R. D. Morphogenesis of intestinal villi. I. Scanning electron microscopy of the duodenal epithelium of the developing chick embryo. J. Morphol. 137, 193–213 (1972).
Burgess, D. R. Morphogenesis of intestinal villi. II. Mechanism of formation of previllous ridges. J. Embryol. Exp. Morphol. 34, 723–740 (1975).
Walton, K. D., Freddo, A. M., Wang, S. & Gumucio, D. L. Generation of intestinal surface: an absorbing tale. Development 143, 2261–2272 (2016).
Walton, K. D. et al. Villification in the mouse: Bmp signals control intestinal villus patterning. Development 143, 427–436 (2016).
Harris, A. K., Stopak, D. & Warner, P. Generation of spatially periodic patterns by a mechanical instability: a mechanical alternative to the Turing model. J. Embryol. Exp. Morphol. 80, 1–20 (1984).
Devenport, D. Tissue morphodynamics: translating planar polarity cues into polarized cell behaviors. Semin. Cell Dev. Biol. 55, 99–110 (2016).
Shindo, A. Models of convergent extension during morphogenesis. Wiley Interdiscip. Rev. Dev. Biol. 7, e293 (2018).
Huebner, R. J. & Wallingford, J. B. Coming to consensus: a unifying model emerges for convergent extension. Dev. Cell 46, 389–396 (2018).
Keller, R. & Tibbetts, P. Mediolateral cell intercalation in the dorsal, axial mesoderm of Xenopus laevis. Dev. Biol. 131, 539–549 (1989).
Wilson, P. & Keller, R. Cell rearrangement during gastrulation of Xenopus: direct observation of cultured explants. Development 112, 289–300 (1991).
Shih, J. & Keller, R. Patterns of cell motility in the organizer and dorsal mesoderm of Xenopus laevis. Development 116, 915–930 (1992).
Shih, J. & Keller, R. Cell motility driving mediolateral intercalation in explants of Xenopus laevis. Development 116, 901–914 (1992).
Davidson, L. A., Marsden, M., Keller, R. & Desimone, D. W. Integrin α5β1 and fibronectin regulate polarized cell protrusions required for Xenopus convergence and extension. Curr. Biol. 16, 833–844 (2006).
Skoglund, P., Rolo, A., Chen, X., Gumbiner, B. M. & Keller, R. Convergence and extension at gastrulation require a myosin IIB-dependent cortical actin network. Development 135, 2435–2444 (2008).
Munro, E. M. & Odell, G. M. Polarized basolateral cell motility underlies invagination and convergent extension of the ascidian notochord. Development 129, 13–24 (2002).
Williams-Masson, E. M., Heid, P. J., Lavin, C. A. & Hardin, J. The cellular mechanism of epithelial rearrangement during morphogenesis of the Caenorhabditis elegans dorsal hypodermis. Dev. Biol. 204, 263–276 (1998).
Walck-Shannon, E., Reiner, D. & Hardin, J. Polarized Rac-dependent protrusions drive epithelial intercalation in the embryonic epidermis of C. elegans. Development 142, 3549–3560 (2015).
Sun, Z. et al. Basolateral protrusion and apical contraction cooperatively drive Drosophila germ-band extension. Nat. Cell Biol. 19, 375–383 (2017).
Williams, M., Yen, W., Lu, X. & Sutherland, A. Distinct apical and basolateral mechanisms drive planar cell polarity-dependent convergent extension of the mouse neural plate. Dev. Cell 29, 34–46 (2014).
Irvine, K. D. & Wieschaus, E. Cell intercalation during Drosophila germband extension and its regulation by pair-rule segmentation genes. Development 120, 827–841 (1994).
Bertet, C., Sulak, L. & Lecuit, T. Myosin-dependent junction remodelling controls planar cell intercalation and axis elongation. Nature 429, 667–671 (2004).
Blankenship, J. T., Backovic, S. T., Sanny, J. S., Weitz, O. & Zallen, J. A. Multicellular rosette formation links planar cell polarity to tissue morphogenesis. Dev. Cell 11, 459–470 (2006). Together with Bertet et al. (2004), shows that cell intercalation during germband extension in D. melanogaster is due to a planar polarized process of cell–cell contact remodelling in which MyoII accumulates at shrinking junctions.
Fernandez-Gonzalez, R. & Zallen, J. A. Oscillatory behaviors and hierarchical assembly of contractile structures in intercalating cells. Phys. Biol. 8, 045005 (2011).
Collinet, C., Rauzi, M., Lenne, P. F. & Lecuit, T. Local and tissue-scale forces drive oriented junction growth during tissue extension. Nat. Cell Biol. 17, 1247–1258 (2015).
Yu, J. C. & Fernandez-Gonzalez, R. Local mechanical forces promote polarized junctional assembly and axis elongation in Drosophila. eLife 5, e10757 (2016).
Rauzi, M., Verant, P., Lecuit, T. & Lenne, P. F. Nature and anisotropy of cortical forces orienting Drosophila tissue morphogenesis. Nat. Cell Biol. 10, 1401–1410 (2008).
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).
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).
Kale, G. R. et al. Distinct contributions of tensile and shear stress on E-cadherin levels during morphogenesis. Nat. Commun. 9, 5021 (2018).
Rozbicki, E. et al. Myosin-II-mediated cell shape changes and cell intercalation contribute to primitive streak formation. Nat. Cell Biol. 17, 397–408 (2015).
Nishimura, T., Honda, H. & Takeichi, M. Planar cell polarity links axes of spatial dynamics in neural-tube closure. Cell 149, 1084–1097 (2012).
Shindo, A. & Wallingford, J. B. PCP and septins compartmentalize cortical actomyosin to direct collective cell movement. Science 343, 649–652 (2014).
Shindo, A., Inoue, Y., Kinoshita, M. & Wallingford, J. B. PCP-dependent transcellular regulation of actomyosin oscillation facilitates convergent extension of vertebrate tissue. Dev. Biol. 446, 159–167 (2019).
Zallen, J. A. & Wieschaus, E. Patterned gene expression directs bipolar planar polarity in Drosophila. Dev. Cell 6, 343–355 (2004).
Pare, A. C. et al. A positional Toll receptor code directs convergent extension in Drosophila. Nature 515, 523–527 (2014). Identifies three Toll receptors as downstream targets of pair-rule genes that direct planar polarization in cells of D. melanogaster germband.
Lavalou, J. et al. Formation of mechanical interfaces by self-organized Toll-8/Cirl GPCR asymmetry. Preprint at bioRxiv https://doi.org/10.1101/2020.03.16.993758 (2020).
Wallingford, J. B. Planar cell polarity and the developmental control of cell behavior in vertebrate embryos. Annu. Rev. Cell Dev. Biol. 28, 627–653 (2012).
Yang, Y. & Mlodzik, M. Wnt-Frizzled/planar cell polarity signaling: cellular orientation by facing the wind (Wnt). Annu. Rev. Cell Dev. Biol. 31, 623–646 (2015).
Butler, M. T. & Wallingford, J. B. Planar cell polarity in development and disease. Nat. Rev. Mol. Cell Biol. 18, 375–388 (2017).
Strutt, D. I., Weber, U. & Mlodzik, M. The role of RhoA in tissue polarity and Frizzled signalling. Nature 387, 292–295 (1997).
Winter, C. G. et al. Drosophila Rho-associated kinase (Drok) links Frizzled-mediated planar cell polarity signaling to the actin cytoskeleton. Cell 105, 81–91 (2001).
Andreeva, A. et al. PTK7-Src signaling at epithelial cell contacts mediates spatial organization of actomyosin and planar cell polarity. Dev. Cell 29, 20–33 (2014).
Habas, R., Dawid, I. B. & He, X. Coactivation of Rac and Rho by Wnt/Frizzled signaling is required for vertebrate gastrulation. Genes Dev. 17, 295–309 (2003).
Heisenberg, C. P. et al. Silberblick/Wnt11 mediates convergent extension movements during zebrafish gastrulation. Nature 405, 76–81 (2000).
Wallingford, J. B. et al. Dishevelled controls cell polarity during Xenopus gastrulation. Nature 405, 81–85 (2000).
Keys, D. N., Levine, M., Harland, R. M. & Wallingford, J. B. Control of intercalation is cell-autonomous in the notochord of Ciona intestinalis. Dev. Biol. 246, 329–340 (2002).
Butler, M. T. & Wallingford, J. B. Spatial and temporal analysis of PCP protein dynamics during neural tube closure. eLife 7, e36456 (2018).
Habas, R., Kato, Y. & He, X. Wnt/Frizzled activation of Rho regulates vertebrate gastrulation and requires a novel Formin homology protein Daam1. Cell 107, 843–854 (2001).
Kim, S. K. et al. Planar cell polarity acts through septins to control collective cell movement and ciliogenesis. Science 329, 1337–1340 (2010).
Levayer, R. & Lecuit, T. Oscillation and polarity of E-cadherin asymmetries control actomyosin flow patterns during morphogenesis. Dev. Cell 26, 162–175 (2013).
Lye, C. M. et al. Mechanical coupling between endoderm invagination and axis extension in Drosophila. PLoS Biol. 13, e1002292 (2015).
Saadaoui, M., Rocancourt, D., Roussel, J., Corson, F. & Gros, J. A tensile ring drives tissue flows to shape the gastrulating amniote embryo. Science 367, 453–458 (2020).
Aigouy, B. et al. Cell flow reorients the axis of planar polarity in the wing epithelium of Drosophila. Cell 142, 773–786 (2010). Shows that contraction of the hinge region during morphogenesis of the wing in Drosophila mechanically directs cell flows and ultimately orients PCP in the blade region.
Etournay, R. et al. Interplay of cell dynamics and epithelial tension during morphogenesis of the Drosophila pupal wing. eLife 4, e07090 (2015).
Ray, R. P. et al. Patterned anchorage to the apical extracellular matrix defines tissue shape in the developing appendages of Drosophila. Dev. Cell 34, 310–322 (2015).
Munster, S. et al. Attachment of the blastoderm to the vitelline envelope affects gastrulation of insects. Nature 568, 395–399 (2019). Shows that blastoderm attachment to the enveloping vitelline membrane orients cellular flows during gastrulation of the beetle Tribolium castaneum, similar to Bailles et al. (2019). Shows the relevance of tissue boundary conditions in directing tissue flows during morphogenesis.
Sato, K. et al. Left-right asymmetric cell intercalation drives directional collective cell movement in epithelial morphogenesis. Nat. Commun. 6, 10074 (2015).
Coulombre, A. J. The role of intraocular pressure in the development of the chick eye. II. Control of corneal size. AMA Arch. Ophthalmol. 57, 250–253 (1957).
Desmond, M. E. & Jacobson, A. G. Embryonic brain enlargement requires cerebrospinal fluid pressure. Dev. Biol. 57, 188–198 (1977).
Abbas, L. & Whitfield, T. T. Nkcc1 (Slc12a2) is required for the regulation of endolymph volume in the otic vesicle and swim bladder volume in the zebrafish larva. Development 136, 2837–2848 (2009).
Navis, A., Marjoram, L. & Bagnat, M. Cftr controls lumen expansion and function of Kupffer’s vesicle in zebrafish. Development 140, 1703–1712 (2013).
Ruiz-Herrero, T., Alessandri, K., Gurchenkov, B. V., Nassoy, P. & Mahadevan, L. Organ size control via hydraulically gated oscillations. Development 144, 4422–4427 (2017).
Dasgupta, S., Gupta, K., Zhang, Y., Viasnoff, V. & Prost, J. Physics of lumen growth. Proc. Natl Acad. Sci. USA 115, E4751–E4757 (2018). Quantitatively analyses the contributions of osmotic pressure, cortical tension, cell–cell contact geometry and paracellular leak in the expansion of intercellular lumens.
Chan, C. J. et al. Hydraulic control of mammalian embryo size and cell fate. Nature 571, 112–116 (2019). Shows that feedback between luminal pressure and cortical tension controls the expansion of the mouse blastocysts and affects cell fate specification.
Bryant, D. M. & Mostov, K. E. From cells to organs: building polarized tissue. Nat. Rev. Mol. Cell Biol. 9, 887–901 (2008).
Sigurbjornsdottir, S., Mathew, R. & Leptin, M. Molecular mechanisms of de novo lumen formation. Nat. Rev. Mol. Cell Biol. 15, 665–676 (2014).
Sperber, I. Secretion of organic anions in the formation of urine and bile. Pharmacol. Rev. 11, 109–134 (1959).
Boyer, J. L. Bile formation and secretion. Compr. Physiol. 3, 1035–1078 (2013).
Navis, A. & Bagnat, M. Developing pressures: fluid forces driving morphogenesis. Curr. Opin. Genet. Dev. 32, 24–30 (2015).
Navis, A. & Nelson, C. M. Pulling together: tissue-generated forces that drive lumen morphogenesis. Semin. Cell Dev. Biol. 55, 139–147 (2016).
Mosaliganti, K. R. et al. Size control of the inner ear via hydraulic feedback. eLife 8, e39596 (2019).
Takaoka, K. & Hamada, H. Cell fate decisions and axis determination in the early mouse embryo. Development 139, 3–14 (2012).
Dumortier, J. G. et al. Hydraulic fracturing and active coarsening position the lumen of the mouse blastocyst. Science 365, 465–468 (2019). Illustrates how the formation of blastocoel and the first symmetry breaking event in the mouse embryo depend on a self-organized process of hydraulic fracturing of cell–cell contacts followed by contractility-directed coarsening of microlumens.
Maitre, J. L. et al. Asymmetric division of contractile domains couples cell positioning and fate specification. Nature 536, 344–348 (2016).
Samarage, C. R. et al. Cortical tension allocates the first inner cells of the mammalian embryo. Dev. Cell 34, 435–447 (2015).
Plusa, B. et al. Downregulation of Par3 and aPKC function directs cells towards the ICM in the preimplantation mouse embryo. J. Cell Sci. 118, 505–515 (2005).
Nishioka, N. et al. The Hippo signaling pathway components Lats and Yap pattern Tead4 activity to distinguish mouse trophectoderm from inner cell mass. Dev. Cell 16, 398–410 (2009).
Korotkevich, E. et al. The apical ___domain is required and sufficient for the first lineage segregation in the mouse embryo. Dev. Cell 40, 235–247 e237 (2017).
Ryan, A. Q., Chan, C. J., Graner, F. & Hiiragi, T. Lumen expansion facilitates epiblast-primitive endoderm fate specification during mouse blastocyst formation. Dev. Cell 51, 684–697 e684 (2019).
Metzger, R. J. & Krasnow, M. A. Genetic control of branching morphogenesis. Science 284, 1635–1639 (1999).
Affolter, M., Zeller, R. & Caussinus, E. Tissue remodelling through branching morphogenesis. Nat. Rev. Mol. Cell Biol. 10, 831–842 (2009).
Ochoa-Espinosa, A. & Affolter, M. Branching morphogenesis: from cells to organs and back. Cold Spring Harb Perspect. Biol. 4, a008243, https://doi.org/10.1101/cshperspect.a008243 (2012).
Affolter, M. & Caussinus, E. Tracheal branching morphogenesis in Drosophila: new insights into cell behaviour and organ architecture. Development 135, 2055–2064 (2008).
Sutherland, D., Samakovlis, C. & Krasnow, M. A. branchless encodes a Drosophila FGF homolog that controls tracheal cell migration and the pattern of branching. Cell 87, 1091–1101 (1996).
Metzger, R. J., Klein, O. D., Martin, G. R. & Krasnow, M. A. The branching programme of mouse lung development. Nature 453, 745–750 (2008). Reveals that the branching pattern of the mouse airways follows stereotypical patterns and sequences and suggests that it reflects the execution of a genetic programme instructing the morphogenesis of branched organs.
Hannezo, E. et al. A unifying theory of branching morphogenesis. Cell 171, 242–255 e227 (2017). Shows how the stochastic behaviour at the tips of a growing network accounts for several features of branching in the mouse mammary glands and kidney and the human prostate. It shows the relevance of self-organization in the morphogenesis of branched organs.
Halevi, S. et al. The C. elegans ric-3 gene is required for maturation of nicotinic acetylcholine receptors. EMBO J. 21, 1012–1020 (2002).
Tsalik, E. L. et al. LIM homeobox gene-dependent expression of biogenic amine receptors in restricted regions of the C. elegans nervous system. Dev. Biol. 263, 81–102 (2003).
Dong, X., Liu, O. W., Howell, A. S. & Shen, K. An extracellular adhesion molecule complex patterns dendritic branching and morphogenesis. Cell 155, 296–307 (2013).
Salzberg, Y. et al. Skin-derived cues control arborization of sensory dendrites in Caenorhabditis elegans. Cell 155, 308–320 (2013).
Zou, W. et al. A multi-protein receptor-ligand complex underlies combinatorial dendrite guidance choices in C. elegans. eLife 5, e18345 (2016).
Palavalli, A., Tizón-Escamilla, N., Rupprecht, J.-F. & Lecuit, T. Deterministic and stochastic rules of branching govern dendritic morphogenesis of sensory neurons. Curr. Biol. https://doi.org/10.1016/j.cub.2020.10.054 (2020). Shows that the morphogenesis of the dendritic arbour in a class of sensory neurons in Drosophila embryos results from a combination of deterministic events and self-organizaion.
Hashimoto, H., Robin, F. B., Sherrard, K. M. & Munro, E. M. Sequential contraction and exchange of apical junctions drives zippering and neural tube closure in a simple chordate. Dev. Cell 32, 241–255 (2015). Shows that the process of neural tube closure in the chordate Ciona robusta depends on a self-organized cycle of MyoII contraction and neighbour exchange at the front of the zipping wave. Shows the relevance of self-organized processes in moprhogenesis.
Hashimoto, H. & Munro, E. Differential expression of a classic cadherin directs tissue-level contractile asymmetry during neural tube closure. Dev. Cell 51, 158–172 e154 (2019).
Munro, E. & Bowerman, B. Cellular symmetry breaking during Caenorhabditis elegans development. Cold Spring Harb. Perspect. Biol. 1, a003400 (2009).
Green, J. B. & Sharpe, J. Positional information and reaction-diffusion: two big ideas in developmental biology combine. Development 142, 1203–1211 (2015).
Acknowledgements
The authors thank all members of the Lecuit group for stimulating discussions and useful feedback on this manuscript. This review emerged from a lecture series at the Collège de France in 2018. The lab is supported by the ERC grant SelfControl #788308 and the Ligue contre le Cancer. C.C. is supported by the CNRS and T.L. by the Collège de France.
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Glossary
- Denticles
-
Small cuticular bristles on the ventral side of Drosophila melanogaster larvae that are used for locomotion.
- Strain
-
A measure of deformation of an object with respect to a reference length upon application of a mechanical stress. This is a dimensionless parameter
- Reaction–diffusion systems
-
Mathematical models describing the change in space and time of the concentration of one or more chemical substances. They typically consider local chemical reactions producing or consuming chemical species and their diffusion.
- Turing instabilities
-
A reaction–diffusion system in which the homogeneous equilibrium of mixed chemical substances is unstable owing to random fluctuations and differential diffusion. This gives rise to stationary wave patterns.
- Excitable systems
-
Mechanochemical systems in which positive feedback and delayed negative feedback produce dynamical patterns of activity exhibiting bistability (a stable switch to an on or off state), pulses or oscillations. Spatial coupling mechanisms (for example, diffusion) lead to the emergence of waves of activity as illustrated in the classical example of the action potential.
- Morphogenetic fields
-
Groups of cells responding to discrete, localized biochemical signals leading to the development of specific morphological structures or organs.
- Mechanical stress
-
A physical quantity that expresses the mechanical forces that neighbouring particles of a continuous material exert on each other. It has the dimension of force per surface area (N m−2) or pressure (Pa).
- Viscous response
-
Deformation of a viscous element, which resists shear flow and strain linearly with time when a stress is applied.
- Boundary conditions
-
Constraints defining the limits of a system. In the case of morphogenesis these are typically the physical boundary of a tissue or an embryo.
- Lateral line
-
A sensory system comprising clusters of mechanosensory epithelial cells (neuromasts) arranged as rosettes with their apical surface facing a shared lumen. The lateral line is initially established by a migratory group of cells, called a primordium, that deposits neuromasts at stereotyped locations along the surface of the fish.
- Mena/VASP
-
Members of the VASP (vasodilator-stimulated phosphoprotein) family of proteins regulating the dynamics of the cortical actin cytoskeleton as downstream effectors of the Rho-family small G proteins Rac and Cdc42.
- Shroom family proteins
-
Family of proteins characterized by a specific arrangement of an N-terminal PDZ ___domain, a central ASD1 (Apx/Shrm Domain 1) motif and a C-terminal ASD2 motif. ASD1 is required for targeting actin, while ASD2 is capable of eliciting an actomyosin constriction event.
- Vitelline membrane
-
Structure surrounding the outer surface of blastoderm cells in embryos of several animals including birds and insects.
- Convergence–extension
-
The process by which a tissue changes shape by narrowing (converging) in one direction and extending along a perpendicular axis.
- Traction forces
-
Forces used to generate motion between a body and a tangential surface, through the use of friction or adhesion. Contractile systems anchored to a rigid body can generate traction forces to move cells or cellular objects.
- Notochord
-
A small flexible rod made from cells from the mesoderm and oriented head to tail in embryos of organisms of the phylum Chordata. As it is composed of stiffer tissue, it allows for skeletal support of the embryo during development.
- Germband
-
Blastoderm tissue corresponding to the ventrolateral region of the embryo in Drosophila melanogaster and other insects.
- Primitive streak
-
Transient structure that forms in the blastula during the early stages of avian, reptilian and mammalian embryonic development. It forms on the dorsal (back) face of the embryo, towards the caudal or posterior end.
- Marginal zone
-
Region corresponding to the equator between the two hemispheres in amphibian embryos.
- Pair-rule genes
-
Group of genes expressed in stripes during segmentation of the embryo in arthropods. In Drosophila pair-rule genes form seven dorsoventrally oriented stripes disposed along the antero-posterior axis.
- Toll receptors
-
A class of single-pass transmembrane receptors involved in patterning and immunity.
- Parasegment
-
The fundamental unit of Drosophila melanogaster development, which is made up of portions of two adjacent segments along the body of the embryo.
- Planar cell polarity
-
The coordinated polarization of a field of cells within the plane of a cell sheet. The axis of planar polarity is typically orthogonal to that of the apico-basal polarity of epithelial cells.
- Septins
-
Cytoskeletal components that upon binding to GTP can polymerize into ordered structures such as rings and filaments, which can function as scaffolds or diffusion barriers.
- Advection
-
The transport of a substance or physical quantity by the movement of the surrounding environment.
- Dissipation timescale
-
Characteristic time at which the internal mechanical stress is reduced by a certain amount by viscous flow.
- Bandpass filter
-
A filter or device that passes frequencies within a certain range and rejects frequencies outside that range.
- Epiblast
-
Also known as primitive ectoderm. One of two distinct layers arising from the inner cell mass in the mammalian blastocyst or the blastodisc in avian and reptile embryos. In mammals, the epiblast sits between the trophectoderm and the hypoblast (or primitive endoderm).
- Primitive endoderm
-
Also known as hypoblast. One of two layers arising from the inner cell mass in the mammalian blastocyst. The primitive endoderm sits between the epiblast and the blastocoel.
- PVD neurons
-
Sensory neurons responding to harsh touch and cold temperatures with a highly elaborate dendritic arborization in the nematode Caenorhabditis elegans.
- Vpda class I neurons
-
Sensory neurons of the peripheral nervous system of Drosophila melanogaster embryos and larvae. The classification is based on the morphology of the dendritic arborization with class I being the simplest morphology and class IV the most complex.
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Collinet, C., Lecuit, T. Programmed and self-organized flow of information during morphogenesis. Nat Rev Mol Cell Biol 22, 245–265 (2021). https://doi.org/10.1038/s41580-020-00318-6
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DOI: https://doi.org/10.1038/s41580-020-00318-6
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