Epithelial morphogenesis is regulated by the modulation of intercellular junctions, which are known as adherens junctions (AJs).
AJs comprise cadherin receptors and associated proteins, including actomyosin filaments.
The interactions between cadherins and actomyosin are mediated by α-catenin and vinculin (VCL) through complex mechanisms.
The control of AJ-associated actomyosin by small GTPases is important for maintaining and remodelling the AJ.
RHO-associated protein kinase (ROCK), an effector of RHO GTPases, induces the contraction of AJ-linked actomyosin networks, which leads to various forms of epithelial remodelling.
AJs are also modulated by other mechanisms, including cadherin turnover, sliding of the junctions and transcriptional control of junction regulators.
Epithelial cells display dynamic behaviours, such as rearrangement, movement and shape changes, particularly during embryonic development and in equivalent processes in adults. Accumulating evidence suggests that the remodelling of cell junctions, especially adherens junctions (AJs), has major roles in controlling these behaviours. AJs comprise cadherin adhesion receptors and cytoplasmic proteins that associate with them, including catenins and actin filaments, and exhibit various forms, such as linear or punctate. Remodelling of AJs induces epithelial reshaping in various ways, including by planar-polarized apical constriction that is driven by the contraction of AJ-associated actomyosin and that occurs during neural plate bending and germband extension. RHO GTPases and their effectors regulate actin polymerization and actomyosin contraction at AJs during the epithelial reshaping processes.
Subscribe to Journal
Get full journal access for 1 year
only $4.92 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Lecuit, T. Adhesion remodeling underlying tissue morphogenesis. Trends Cell Biol. 15, 34–42 (2005).
Baum, B. & Georgiou, M. Dynamics of adherens junctions in epithelial establishment, maintenance, and remodeling. J. Cell Biol. 192, 907–917 (2011).
Oda, H. & Takeichi, M. Evolution: structural and functional diversity of cadherin at the adherens junction. J. Cell Biol. 193, 1137–1146 (2011).
Hirano, S. & Takeichi, M. Cadherins in brain morphogenesis and wiring. Physiol. Rev. 92, 597–634 (2012).
Lynch, A. M. et al. A genome-wide functional screen shows MAGI 1 is an L1CAM dependent stabilizer of apical junctions in C. elegans. Curr. Biol. 22, 1891–1899 (2012).
Truong Quang, B. A., Mani, M., Markova, O., Lecuit, T. & Lenne, P. F. Principles of E-adherin supramolecular organization in vivo. Curr. Biol. 23, 2197–2207 (2013).
Farquhar, M. G. & Palade, G. E. Junctional complexes in various epithelia. J. Cell Biol. 17, 375–412 (1963).
Taguchi, K., Ishiuchi, T. & Takeichi, M. Mechanosensitive EPLIN-dependent remodeling of adherens junctions regulates epithelial reshaping. J. Cell Biol. 194, 643–656 (2011). Shows that epithelial linear junctions are converted into a punctate type at colony peripheries, where radial actin filaments prevent EPLIN from binding to AJs.
Kametani, Y. & Takeichi, M. Basal to apical cadherin flow at cell junctions. Nature Cell Biol. 9, 92–98 (2007).
Brasch, J., Harrison, O. J., Honig, B. & Shapiro, L. Thinking outside the cell: how cadherins drive adhesion. Trends Cell Biol. 22, 299–310 (2012).
Hirano, S., Kimoto, N., Shimoyama, Y., Hirohashi, S. & Takeichi, M. Identification of a neural α-catenin as a key regulator of cadherin function and multicellular organization. Cell 70, 293–301 (1992).
Watabe-Uchida, M. et al. α-Catenin-vinculin interaction functions to organize the apical junctional complex in epithelial cells. J. Cell Biol. 142, 847–857 (1998).
Rimm, D. L., Koslov, E. R., Kebriaei, P., Cianci, C. D. & Morrow, J. S. α1(E)-catenin is an actin-binding and -bundling protein mediating the attachment of F actin to the membrane adhesion complex. Proc. Natl Acad. Sci. USA 92, 8813–8817 (1995).
Hansen, S. D. et al. αE-catenin actin-binding domain alters actin filament conformation and regulates binding of nucleation and disassembly factors. Mol. Biol. Cell 24, 3710–3720 (2013).
Yamada, S., Pokutta, S., Drees, F., Weis, W. I. & Nelson, W. J. Deconstructing the cadherin–catenin–actin complex. Cell 123, 889–901 (2005). Reports the unexpected finding that, despite the well-known ability of α-catenin to bind to F-actin, the cadherin–α-catenin complex does not do so.
Drees, F., Pokutta, S., Yamada, S., Nelson, W. J. & Weis, W. I. α-catenin is a molecular switch that binds E cadherin-β-catenin and regulates actin-filament assembly. Cell 123, 903–915 (2005).
Kwiatkowski, A. V. et al. In vitro and in vivo reconstitution of the cadherin–catenin–actin complex from Caenorhabditis elegans. Proc. Natl Acad. Sci. USA 107, 14591–14596 (2010).
Yonemura, S. Wada, Y., Watanabe, T., Nagafuchi, A. & Shibata, M. α-Catenin as a tension transducer that induces adherens junction development. Nature Cell Biol. 12, 533–542 (2010). First to propose that the binding of VCL to α-catenin is inhibited by self-folding of α-catenin, and this autoinhibited state is reversed by a force that pulls the α-catenin C-terminal domain.
Desai, R. et al. Monomeric α-catenin links cadherin to the actin cytoskeleton. Nature Cell Biol. 15, 261–273 (2013).
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).
Twiss, F. et al. Vinculin-dependent cadherin mechanosensing regulates efficient epithelial barrier formation. Biol. Open 1, 1128–1140 (2012).
Ishiyama, N. et al. An autoinhibited structure of α-catenin and its implications for vinculin recruitment to adherens junctions. J. Biol. Chem. 288, 15913–15925 (2013).
Maddugoda, M. P., Crampton, M. S., Shewan, A. M. & Yap, A. S. Myosin VI and vinculin cooperate during the morphogenesis of cadherin cell cell contacts in mammalian epithelial cells. J. Cell Biol. 178, 529–540 (2007).
Peng, X., Cuff, L. E., Lawton, C. D. & DeMali, K. A. Vinculin regulates cell-surface E cadherin expression by binding to β-catenin. J. Cell Sci. 123, 567–577 (2010).
Rangarajan, E. S. & Izard, T. Dimer asymmetry defines α-catenin interactions. Nature Struct. Mol. Biol. 20, 188–193 (2013).
Choi, H. J. et al. αE catenin is an autoinhibited molecule that coactivates vinculin. Proc. Natl Acad. Sci. USA 109, 8576–8581 (2012).
Rangarajan, E. S. & Izard, T. The cytoskeletal protein α-catenin unfurls upon binding to vinculin. J. Biol. Chem. 287, 18492–18499 (2012).
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).
Imamura, Y., Itoh, M., Maeno, Y., Tsukita, S. & Nagafuchi, A. Functional domains of α-catenin required for the strong state of cadherin-based cell adhesion. J. Cell Biol. 144, 1311–1322 (1999).
Huveneers, S. et al. Vinculin associates with endothelial VE cadherin junctions to control force-dependent remodeling. J. Cell Biol. 196, 641–652 (2012).
Millan, J. et al. Adherens junctions connect stress fibres between adjacent endothelial cells. BMC Biol. 8, 11 (2010).
Maul, R. S. et al. EPLIN regulates actin dynamics by cross-linking and stabilizing filaments. J. Cell Biol. 160, 399–407 (2003).
Abe, K. & Takeichi, M. EPLIN mediates linkage of the cadherin–catenin complex to F actin and stabilizes the circumferential actin belt. Proc. Natl Acad. Sci. USA 105, 13–19 (2008).
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).
Chervin-Petinot, A. et al. Epithelial protein lost in neoplasm (EPLIN) interacts with α-catenin and actin filaments in endothelial cells and stabilizes vascular capillary network in vitro. J. Biol. Chem. 287, 7556–7572 (2012).
Palacios, F., Schweitzer, J. K., Boshans, R. L. & D'Souza-Schorey, C. ARF6 GTP recruits Nm23 H1 to facilitate dynamin-mediated endocytosis during adherens junctions disassembly. Nature Cell Biol. 4, 929–936 (2002).
Delva, E. & Kowalczyk, A. P. Regulation of cadherin trafficking. Traffic 10, 259–267 (2009).
Davis, M. A., Ireton, R. C. & Reynolds, A. B. A core function for p120 catenin in cadherin turnover. J. Cell Biol. 163, 525–534 (2003).
Xiao, K. et al. Cellular levels of p120 catenin function as a set point for cadherin expression levels in microvascular endothelial cells. J. Cell Biol. 163, 535–545 (2003).
Myster, S. H., Cavallo, R., Anderson, C. T., Fox, D. T. & Peifer, M. Drosophila p120catenin plays a supporting role in cell adhesion but is not an essential adherens junction component. J. Cell Biol. 160, 433–449 (2003).
Pettitt, J., Cox, E. A., Broadbent, I. D., Flett, A. & Hardin, J. The Caenorhabditis elegans p120 catenin homologue, JAC 1, modulates cadherin-catenin function during epidermal morphogenesis. J. Cell Biol. 162, 15–22 (2003).
Nanes, B. A. et al. p120 catenin binding masks an endocytic signal conserved in classical cadherins. J. Cell Biol. 199, 365–380 (2012). Dissects the core p120 catenin-binding region of VE-cadherin and identifies a specific sequence that is masked by this catenin to prevent cadherin internalization.
Miyashita, Y. & Ozawa, M. Increased internalization of p120 uncoupled E cadherin and a requirement for a dileucine motif in the cytoplasmic domain for endocytosis of the protein. J. Biol. Chem. 282, 11540–11548 (2007).
de Beco, S., Gueudry, C., Amblard, F. & Coscoy, S. Endocytosis is required for E cadherin redistribution at mature adherens junctions. Proc. Natl Acad. Sci. USA 106, 7010–7015 (2009).
Troyanovsky, R. B., Sokolov, E. P. & Troyanovsky, S. M. Endocytosis of cadherin from intracellular junctions is the driving force for cadherin adhesive dimer disassembly. Mol. Biol. Cell 17, 3484–3493 (2006).
Kane, D. A., McFarland, K. N. & Warga, R. M. Mutations in half baked/E cadherin block cell behaviors that are necessary for teleost epiboly. Development 132, 1105–1116 (2005).
Song, S. et al. Pou5f1-dependent EGF expression controls E cadherin endocytosis, cell adhesion, and zebrafish epiboly movements. Dev. Cell 24, 486–501 (2013).
Ulrich, F. et al. Wnt11 functions in gastrulation by controlling cell cohesion through Rab5c and E cadherin. Dev. Cell 9, 555–564 (2005).
Arboleda-Estudillo, Y. et al. Movement directionality in collective migration of germ layer progenitors. Curr. Biol. 20, 161–169 (2010).
Tang, V. W. & Brieher, W. M. α-Actinin 4/FSGS1 is required for Arp2/3 dependent actin assembly at the adherens junction. J. Cell Biol. 196, 115–130 (2012).
Verma, S. et al. A WAVE2 Arp2/3 actin nucleator apparatus supports junctional tension at the epithelial zonula adherens. Mol. Biol. Cell 23, 4601–4610 (2012).
Georgiou, M., Marinari, E., Burden, J. & Baum, B. Cdc42, Par6, and aPKC regulate Arp2/3 mediated endocytosis to control local adherens junction stability. Curr. Biol. 18, 1631–1638 (2008).
Kovacs, E. M. et al. N-WASP regulates the epithelial junctional actin cytoskeleton through a non-canonical post-nucleation pathway. Nature Cell Biol. 13, 934–943 (2011).
Otani, T., Ichii, T., Aono, S. & Takeichi, M. Cdc42 GEF Tuba regulates the junctional configuration of simple epithelial cells. J. Cell Biol. 175, 135–146 (2006).
Ivanov, A. I. et al. A unique role for nonmuscle myosin heavy chain IIa in regulation of epithelial apical junctions. PLoS ONE 2, e658 (2007).
McCormack, J., Welsh, N. J. & Braga, V. M. Cycling around cell–cell adhesion with Rho GTPase regulators. J. Cell Sci. 126, 379–391 (2013).
Citi, S., Spadaro, D., Schneider, Y., Stutz, J. & Pulimeno, P. Regulation of small GTPases at epithelial cell–cell junctions. Mol. Membr. Biol. 28, 427–444 (2011).
Goode, B. L. & Eck, M. J. Mechanism and function of formins in the control of actin assembly. Annu. Rev. Biochem. 76, 593–627 (2007).
Carramusa, L., Ballestrem, C., Zilberman, Y. & Bershadsky, A. D. Mammalian diaphanous-related formin Dia1 controls the organization of E cadherin-mediated cell–cell junctions. J. Cell Sci. 120, 3870–3882 (2007).
Sahai, E. & Marshall, C. J. ROCK and Dia have opposing effects on adherens junctions downstream of Rho. Nature Cell Biol. 4, 408–415 (2002).
Homem, C. C. & Peifer, M. Diaphanous regulates myosin and adherens junctions to control cell contractility and protrusive behavior during morphogenesis. Development 135, 1005–1018 (2008).
Kobielak, A., Pasolli, H. A. & Fuchs, E. Mammalian formin 1 participates in adherens junctions and polymerization of linear actin cables. Nature Cell Biol. 6, 21–30 (2004).
Riento, K. & Ridley, A. J. Rocks: multifunctional kinases in cell behaviour. Nature Rev. Mol. Cell Biol. 4, 446–456 (2003).
Ishiuchi, T. & Takeichi, M. Willin and Par3 cooperatively regulate epithelial apical constriction through aPKC-mediated ROCK phosphorylation. Nature Cell Biol. 13, 860–866 (2011).
Fanning, A. S., Van Itallie, C. M. & Anderson, J. M. Zonula occludens 1 and -2 regulate apical cell structure and the zonula adherens cytoskeleton in polarized epithelia. Mol. Biol. Cell 23, 577–590 (2012).
Ayollo, D. V., Zhitnyak, I. Y., Vasiliev, J. M. & Gloushankova, N. A. Rearrangements of the actin cytoskeleton and E cadherin-based adherens junctions caused by neoplasic transformation change cell–cell interactions. PLoS ONE 4, e8027 (2009).
Smutny, M. et al. Myosin II isoforms identify distinct functional modules that support integrity of the epithelial zonula adherens. Nature Cell Biol. 12, 696–702 (2010).
Warner, S. J. & Longmore, G. D. Distinct functions for Rho1 in maintaining adherens junctions and apical tension in remodeling epithelia. J. Cell Biol. 185, 1111–1125 (2009).
Herder, C. et al. ArhGEF18 regulates RhoA Rock2 signaling to maintain neuro-epithelial apico–basal polarity and proliferation. Development 140, 2787–2797 (2013).
Nakajima, H. & Tanoue, T. Lulu2 regulates the circumferential actomyosin tensile system in epithelial cells through p114RhoGEF. J. Cell Biol. 195, 245–261 (2011).
Terry, S. J. et al. Spatially restricted activation of RhoA signalling at epithelial junctions by p114RhoGEF drives junction formation and morphogenesis. Nature Cell Biol. 13, 159–166 (2011).
Ngok, S. P. et al. TEM4 is a junctional Rho GEF required for cell–cell adhesion, monolayer integrity and barrier function. J. Cell Sci. 126, 3271–3277 (2013).
Ratheesh, A. et al. Centralspindlin and α-catenin regulate Rho signalling at the epithelial zonula adherens. Nature Cell Biol. 14, 818–828 (2012). Describes an unforeseen pathway that recruits a RHO GEF to cell junctions; this pathway includes proteins that are involved in cytokinesis.
Aijaz, S., D'Atri, F., Citi, S., Balda, M. S. & Matter, K. Binding of GEF H1 to the tight junction-associated adaptor cingulin results in inhibition of Rho signaling and G1/S phase transition. Dev. Cell 8, 777–786 (2005).
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).
Nishimura, T., Honda, H. & Takeichi, M. Planar cell polarity links axes of spatial dynamics in neural-tube closure. Cell 149, 1084–1097 (2012). Demonstrates that, in the bending neural plate, the cadherin CELSR1 induces contraction of AJ-associated actomyosin along the mediolateral axis, which leads to the polarized bending of the plate.
Zebda, N. et al. Interaction of p190RhoGAP with C terminal domain of p120 catenin modulates endothelial cytoskeleton and permeability. J. Biol. Chem. 288, 18290–18299 (2013).
Holeiter, G. et al. The RhoGAP protein Deleted in Liver Cancer 3 (DLC3) is essential for adherens junctions integrity. Oncogenesis 1, e13 (2012).
Tripathi, V., Popescu, N. C. & Zimonjic, D. B. DLC1 interaction with α-catenin stabilizes adherens junctions and enhances DLC1 antioncogenic activity. Mol. Cell. Biol. 32, 2145–2159 (2012).
Sousa, S. et al. ARHGAP10 is necessary for α-catenin recruitment at adherens junctions and for Listeria invasion. Nature Cell Biol. 7, 954–960 (2005).
Kooistra, M. R., Dube, N. & Bos, J. L. Rap1: a key regulator in cell–cell junction formation. J. Cell Sci. 120, 17–22 (2007).
Hogan, C. et al. Rap1 regulates the formation of E cadherin-based cell–cell contacts. Mol. Cell. Biol. 24, 6690–6700 (2004).
Price, L. S. et al. Rap1 regulates E cadherin-mediated cell–cell adhesion. J. Biol. Chem. 279, 35127–35132 (2004).
Knox, A. L. & Brown, N. H. Rap1 GTPase regulation of adherens junction positioning and cell adhesion. Science 295, 1285–1288 (2002).
Dube, N. et al. The RapGEF PDZ GEF2 is required for maturation of cell–cell junctions. Cell. Signal. 20, 1608–1615 (2008).
Ando, K. et al. Rap1 potentiates endothelial cell junctions by spatially controlling myosin II activity and actin organization. J. Cell Biol. 202, 901–916 (2013).
Mandai, K., Rikitake, Y., Shimono, Y. & Takai, Y. Afadin/AF 6 and canoe: roles in cell adhesion and beyond. Prog. Mol. Biol. Transl. Sci. 116, 433–454 (2013).
Hoshino, T. et al. Regulation of E cadherin endocytosis by nectin through afadin, Rap1, and p120ctn. J. Biol. Chem. 280, 24095–24103 (2005).
Hildebrand, J. D. & Soriano, P. Shroom, a PDZ domain-containing actin-binding protein, is required for neural tube morphogenesis in mice. Cell 99, 485–497 (1999).
Ernst, S. et al. Shroom3 is required downstream of FGF signalling to mediate proneuromast assembly in zebrafish. Development 139, 4571–4581 (2012).
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).
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).
Bolinger, C., Zasadil, L., Rizaldy, R. & Hildebrand, J. D. Specific isoforms of drosophila shroom define spatial requirements for the induction of apical constriction. Dev. Dyn. 239, 2078–2093 (2010).
Plageman, T. F. et al. Pax6 dependent Shroom3 expression regulates apical constriction during lens placode invagination. Development 137, 405–415 (2010).
Hildebrand, J. D. Shroom regulates epithelial cell shape via the apical positioning of an actomyosin network. J. Cell Sci. 118, 5191–5203 (2005).
Mohan, S. et al. Structure of Shroom domain 2 reveals a three-segmented coiled-coil required for dimerization, Rock binding, and apical constriction. Mol. Biol. Cell 23, 2131–2142 (2012).
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).
Simoes Sde, M., Mainieri, A. & Zallen, J. A. Rho GTPase and Shroom direct planar polarized actomyosin contractility during convergent extension. J. Cell Biol. 204, 575–589 (2014).
Plageman, T. F. Jr et al. A Trio-RhoA Shroom3 pathway is required for apical constriction and epithelial invagination. Development 138, 5177–5188 (2011).
Chu, C. W., Gerstenzang, E., Ossipova, O. & Sokol, S. Y. Lulu regulates Shroom-induced apical constriction during neural tube closure. PLoS ONE 8, e81854 (2013).
Wei, L. et al. Rho kinases play an obligatory role in vertebrate embryonic organogenesis. Development 128, 2953–2962 (2001).
Ybot-Gonzalez, P. et al. Convergent extension, planar-cell-polarity signalling and initiation of mouse neural tube closure. Development 134, 789–799 (2007).
Kinoshita, N., Sasai, N., Misaki, K. & Yonemura, S. Apical accumulation of Rho in the neural plate is important for neural plate cell shape change and neural tube formation. Mol. Biol. Cell 19, 2289–2299 (2008).
Usui, T. et al. Flamingo, a seven-pass transmembrane cadherin, regulates planar cell polarity under the control of Frizzled. Cell 98, 585–595 (1999).
Gray, R. S., Roszko, I. & Solnica-Krezel, L. Planar cell polarity: coordinating morphogenetic cell behaviors with embryonic polarity. Dev. Cell 21, 120–133 (2011).
Dawes-Hoang, R. E. et al. folded gastrulation, cell shape change and the control of myosin localization. Development 132, 4165–4178 (2005).
Barrett, K., Leptin, M. & Settleman, J. The Rho GTPase and a putative RhoGEF mediate a signaling pathway for the cell shape changes in Drosophila gastrulation. Cell 91, 905–915 (1997).
Martin, A. C., Kaschube, M. & Wieschaus, E. F. Pulsed contractions of an actin-myosin network drive apical constriction. Nature 457, 495–499 (2009). Reports that the actomyosin networks at the apical cortex of cells display pulsed contractions, which eventually induce apical constriction of the cells owing to a linkage between cortical actomyosins and cell edges.
Spahn, P., Ott, A. & Reuter, R. The PDZ-GEF protein Dizzy regulates the establishment of adherens junctions required for ventral furrow formation in Drosophila. J. Cell Sci. 125, 3801–3812 (2012).
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).
Blanchard, G. B., Murugesu, S., Adams, R. J., Martinez-Arias, A. & Gorfinkiel, N. Cytoskeletal dynamics and supracellular organisation of cell shape fluctuations during dorsal closure. Development 137, 2743–2752 (2010).
David, D. J., Tishkina, A. & Harris, T. J. The PAR complex regulates pulsed actomyosin contractions during amnioserosa apical constriction in Drosophila. Development 137, 1645–1655 (2010).
Roh-Johnson, M. et al. Triggering a cell shape change by exploiting preexisting actomyosin contractions. Science 335, 1232–1235 (2012).
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).
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).
Kurita, S., Yamada, T., Rikitsu, E., Ikeda, W. & Takai, Y. Binding between the junctional proteins afadin and PLEKHA7 and implication in the formation of adherens junction in epithelial cells. J. Biol. Chem. 288, 29356–29368 (2013).
Bertet, C., Sulak, L. & Lecuit, T. Myosin-dependent junction remodelling controls planar cell intercalation and axis elongation. Nature 429, 667–671 (2004).
Zallen, J. A. & Wieschaus, E. Patterned gene expression directs bipolar planar polarity in Drosophila. Dev. Cell 6, 343–355 (2004).
Simoes Sde, M. et al. Rho-kinase directs Bazooka/Par 3 planar polarity during Drosophila axis elongation. Dev. Cell 19, 377–388 (2010).
Blankenship, J. T., Backovic, S. T., Sanny, J. S. P., Weitz, O. & Zallen, J. A. Multicellular rosette formation links planar cell polarity to tissue morphogenesis. Dev. Cell 11, 459–470 (2006).
Fernandez-Gonzalez, R., Simoes, S. D., Roper, J. C., Eaton, S. & Zallen, J. A. Myosin, I. I. Dynamics are regulated by tension in intercalating cells. Dev. Cell 17, 736–743 (2009).
Simoes, S. et al. The role of Bazooka/Par3 in epithelial intercalation - a live imaging approach. Mech. Dev. 126, S81 (2009).
Rauzi, M., Lenne, P. F. & Lecuit, T. Planar polarized actomyosin contractile flows control epithelial junction remodelling. Nature 468, 1110–1114 (2010).
Levayer, R. & Lecuit, T. Oscillation and polarity of E cadherin asymmetries control actomyosin flow patterns during morphogenesis. Dev. Cell 26, 162–175 (2013). Links two subcellular events, movement of cortical actomyosin networks and E-cadherin endocytosis, to explain how the shrinkage of polarized junctions occurs.
Levayer, R., Pelissier-Monier, A. & Lecuit, T. Spatial regulation of Dia and Myosin II by RhoGEF2 controls initiation of E cadherin endocytosis during epithelial morphogenesis. Nature Cell Biol. 13, 529–540 (2011).
Tamada, M., Farrell, D. L. & Zallen, J. A. Abl regulates planar polarized junctional dynamics through β-catenin tyrosine phosphorylation. Dev. Cell 22, 309–319 (2012).
Walck-Shannon, E. & Hardin, J. Cell intercalation from top to bottom. Nature Rev. Mol. Cell Biol. 15, 34–48 (2013).
Wang, Y. C., Khan, Z., Kaschube, M. & Wieschaus, E. F. Differential positioning of adherens junctions is associated with initiation of epithelial folding. Nature 484, 390–393 (2012). Finds a novel mechanism for epithelial reshaping in which the sliding, rather than contraction, of AJs is crucial for apical constriction in certain tissues.
Wang, Y. C., Khan, Z. & Wieschaus, E. F. Distinct Rap1 activity states control the extent of epithelial invagination via α-catenin. Dev. Cell 25, 299–309 (2013).
Taniguchi, K. et al. Chirality in planar cell shape contributes to left-right asymmetric epithelial morphogenesis. Science 333, 339–341 (2011).
Davis, N. M. et al. The chirality of gut rotation derives from left-right asymmetric changes in the architecture of the dorsal mesentery. Dev. Cell 15, 134–145 (2008).
Kurpios, N. A. et al. The direction of gut looping is established by changes in the extracellular matrix and in cell:cell adhesion. Proc. Natl Acad. Sci. USA 105, 8499–8506 (2008).
Plageman, T. F., Zacharias, A. L., Gage, P. J. & Lang, R. A. Shroom3 and a Pitx2 N cadherin pathway function cooperatively to generate asymmetric cell shape changes during gut morphogenesis. Dev. Biol. 357, 227–234 (2011).
Welsh, I. C. et al. Integration of left-right Pitx2 transcription and Wnt signaling drives asymmetric gut morphogenesis via Daam2. Dev. Cell 26, 629–644 (2013).
Wu, S. K. et al. Cortical F actin stabilization generates apical-lateral patterns of junctional contractility that integrate cells into epithelia. Nature Cell Biol. 16, 167–178 (2014).
Alatortsev, V. E., Kramerova, I. A., Frolov, M. V., Lavrov, S. A. & Westphal, E. D. Vinculin gene is non-essential in Drosophila melanogaster. FEBS Lett. 413, 197–201 (1997).
Xu, W., Baribault, H. & Adamson, E. D. Vinculin knockout results in heart and brain defects during embryonic development. Development 125, 327–337 (1998).
Torres, M. et al. An α E catenin gene trap mutation defines its function in preimplantation development. Proc. Natl Acad. Sci. USA 94, 901–906 (1997).
Mason, F. M., Tworoger, M. & Martin, A. C. Apical domain polarization localizes actin-myosin activity to drive ratchet-like apical constriction. Nature Cell Biol. 15, 926–936 (2013).
Stehbens, S. J. et al. Dynamic microtubules regulate the local concentration of E cadherin at cell–cell contacts. J. Cell Sci. 119, 1801–1811 (2006).
Meng, W., Mushika, Y., Ichii, T. & Takeichi, M. Anchorage of microtubule minus ends to adherens junctions regulates epithelial cell–cell contacts. Cell 135, 948–959 (2008).
Harris, T. J. & Peifer, M. aPKC controls microtubule organization to balance adherens junction symmetry and planar polarity during development. Dev. Cell 12, 727–738 (2007).
Shaw, R. M. et al. Microtubule plus-end-tracking proteins target gap junctions directly from the cell interior to adherens junctions. Cell 128, 547–560 (2007).
Ligon, L. A., Karki, S., Tokito, M. & Holzbaur, E. L. Dynein binds to β-catenin and may tether microtubules at adherens junctions. Nature Cell Biol. 3, 913–917 (2001).
Bellett, G. et al. Microtubule plus-end and minus-end capture at adherens junctions is involved in the assembly of apico–basal arrays in polarised epithelial cells. Cell Motil. Cytoskeleton 66, 893–908 (2009).
Harrison, O. J. et al. The extracellular architecture of adherens junctions revealed by crystal structures of type I cadherins. Structure 19, 244–256 (2011).
Ishiyama, N. et al. Dynamic and static interactions between p120 catenin and E-cadherin regulate the stability of cell–cell adhesion. Cell 141, 117–128 (2010).
Huber, A. H. & Weis, W. I. The structure of the β-catenin/E cadherin complex and the molecular basis of diverse ligand recognition by β-catenin. Cell 105, 391–402 (2001).
Pokutta, S. & Weis, W. I. Structure of the dimerization and β-catenin-binding region of α-catenin. Mol. Cell 5, 533–543 (2000).
The author thanks T. Nishimura for critical comments on the manuscript, K. Taguchi for editing the movie and S. Ito for suggestions on references. The author's laboratory is supported by the programme Grants-in-Aid for Specially Promoted Research of the Ministry of Education, Science, Sports and Culture of Japan.
The author declares no competing financial interests.
EPLIN stabilizes the ZA. Time-lapse images of DLD1 cells transfected with Kusabira Orange-tagged epithelial cadherin (E‑cadherin). Left, control cells; right, epithelial protein lost in neoplasm (EPLIN)-depleted cells. Frames were taken every minute for 20 minutes. Frame rate, 2.1 frames/s. Scale bar, 20 µm. Details of the experiments are described in REF. 1. Adherens junctions (AJs) in EPLIN-depleted cells are much more unstable and more mobile than those in wild-type cells, which suggests that EPLIN-mediated actin assembly is essential for the formation of the zonula adherens (ZA). © 2011 Taguchi et al. Journal of Cell Biology. 194:643–656. 10.1083/jcb.201104124. 1. Taguchi, K., Ishiuchi, T. & Takeichi, M. Mechanosensitive EPLIN-dependent remodeling of adherens junctions regulates epithelial reshaping. J. Cell Biol. 194, 643–656 (2011). (MOV 3128 kb)
- Apical constriction
A process to induce the bending of epithelial sheets during various morphogenetic processes, such as gastrulation and neural tube formation. Epithelial cells shrink specifically at the apical ends in response to external or internal signals, and as a result their sheets bend towards the apical ends.
- Zonula adherens
(ZA). A cell–cell adherens junction that forms a circumferential belt around the apical pole of epithelial cells.
- Tight junctions
Circumferential rings at the apex of epithelial cells that seal adjacent cells to one another. Tight junctions regulate solute and ion flux between adjacent epithelial cells.
Junctional structures that are formed by transmembrane proteins that are homologous to cadherins and are called desmocollins and desmogleins. These are linked to plakoglobin and desmoplakin, and are anchored to intermediate filaments.
- Actomyosin cables
Subcellular structures that consist of accumulated actin filaments and myosin II. Sliding of myosin II motors along the actin filaments provides a force to contract the cables. These contracting cables have various roles in cell shape changes, which are dependent on where the cables are anchored.
- Guanine nucleotide exchange factors
(GEFs). Proteins that facilitate the exchange of GDP for GTP in the nucleotide-binding pocket of a GTP-binding protein.
- GTPase-activating proteins
(GAPs). Proteins that inactivate small GTP-binding proteins, such as RAS family members, by increasing their rate of GTP hydrolysis.
- Planar cell polarity
A mechanism of cellular organization by which cells acquire information about their orientation within the tissue in the plane of the epithelium. It is distinct from apical–basal polarity.
- Amnioserosa cells
Cells that form the amnioserosa, an extra-embryonic epithelial sheet that covers the dorsal side of fly embryos at the blastoderm stage.
- Dorsal folds
Epithelial structures that form on the dorsal side of the gastrulating Drosophila melanogaster embryo. Dorsal epithelial sheets are folded at an anterior and posterior portion of the embryo during development. The anterior fold is shallower than the posterior fold.
About this article
Cite this article
Takeichi, M. Dynamic contacts: rearranging adherens junctions to drive epithelial remodelling. Nat Rev Mol Cell Biol 15, 397–410 (2014). https://doi.org/10.1038/nrm3802
Targeting desmosomal adhesion and signalling for intestinal barrier stabilization in inflammatory bowel diseases—Lessons from experimental models and patients
Acta Physiologica (2021)
Communications Biology (2021)
Activated nanoscale actin-binding domain motion in the catenin–cadherin complex revealed by neutron spin echo spectroscopy
Proceedings of the National Academy of Sciences (2021)
G3 Genes|Genomes|Genetics (2021)
Autoregulatory “Multitasking” at Endothelial Cell Junctions by Junction-Associated Intermittent Lamellipodia Controls Barrier Properties
Frontiers in Physiology (2021)