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Tight junctions: from simple barriers to multifunctional molecular gates

Nature Reviews Molecular Cell Biology volume 17, pages 564–580 (2016)Cite this article

Subjects

Key Points

  • Tight junctions are intercellular adhesion complexes in epithelia and endothelia that control paracellular permeability. This paracellular diffusion barrier is semipermeable: it is size- and charge-selective.

  • Paracellular ion permeability at tight junctions is largely determined by their claudin composition. Claudins are a family of transmembrane proteins that are thought to form gated ion-selective paracellular pores through the paracellular diffusion barrier.

  • Tight junctions form the border between the apical and basolateral cell surface domains in polarized epithelia, and support the maintenance of cell polarity by restricting intermixing of apical and basolateral transmembrane components.

  • Tight junctions are an integral component of the evolutionarily conserved signalling mechanisms that control epithelial-cell polarization and the formation of morphologically and functionally distinct apical domains.

  • Tight junctions form bidirectional signalling platforms that receive signals from the cell interior, which regulate their assembly and function, and that transduce signals to the cell interior to control cell proliferation, migration, differentiation and survival.

  • Tight junctions are part of an interconnected network of adhesion complexes that also includes adherens junctions and focal adhesions. These adhesion complexes crosstalk through direct protein–protein interactions as well as by transmitting signals to each other that influence their assembly and function.

Abstract

Epithelia and endothelia separate different tissue compartments and protect multicellular organisms from the outside world. This requires the formation of tight junctions, selective gates that control paracellular diffusion of ions and solutes. Tight junctions also form the border between the apical and basolateral plasma-membrane domains and are linked to the machinery that controls apicobasal polarization. Additionally, signalling networks that guide diverse cell behaviours and functions are connected to tight junctions, transmitting information to and from the cytoskeleton, nucleus and different cell adhesion complexes. Recent advances have broadened our understanding of the molecular architecture and cellular functions of tight junctions.

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Figure 1: The junctional complex and tight junctions.
Figure 2: Models of the structure and function of tight junctions.
Figure 3: Structure of claudins and intercellular pore formation.
Figure 4: Junction assembly and crosstalk between adhesion complexes.

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References

  1. Cereijido, M., Contreras, R. G. & Shoshani, L. Cell adhesion, polarity, and epithelia in the dawn of metazoans. Physiol. Rev. 84, 1229–1262 (2004).

    CAS  PubMed  Google Scholar 

  2. Farquhar, M. G. & Palade, G. E. Junctional complexes in various epithelia. J. Cell Biol. 17, 375–412 (1963).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. Claude, P. & Goodenough, D. A. Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia. J. Cell Biol. 58, 390–400 (1973).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. Staehelin, L. A., Mukherjee, T. M. & Williams, A. W. Freeze-etch appearance of the tight junctions in the epithelium of small and large intestine of mice. Protoplasma 67, 165–184 (1969).

    CAS  PubMed  Google Scholar 

  5. Furuse, M. et al. Overexpression of occludin, a tight junction integral membrane protein, induces the formation of intracellular multilamellar bodies bearing tight junction-like structures. J. Cell Sci. 109, 429–435 (1996).

    CAS  PubMed  Google Scholar 

  6. Furuse, M., Sasaki, H., Fujimoto, K. & Tsukita, S. A single gene product, claudin-1 or -2, reconstitutes tight junction strands and recruits occludin in fibroblasts. J. Cell Biol. 143, 391–401 (1998). Demonstration that expression of claudins is sufficient for intramembrane strand formation in cells that lack tight junctions.

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Morita, K., Furuse, M., Fujimoto, K. & Tsukita, S. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proc. Natl Acad. Sci. USA 96, 511–516 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  8. Kubota, K. et al. Ca2+-independent cell-adhesion activity of claudins, a family of integral membrane proteins localized at tight junctions. Curr. Biol. 9, 1035–1038 (1999).

    CAS  PubMed  Google Scholar 

  9. Van Itallie, C. M. & Anderson, J. M. Occludin confers adhesiveness when expressed in fibroblasts. J. Cell Sci. 110, 1113–1121 (1997).

    CAS  PubMed  Google Scholar 

  10. Osler, M. E., Chang, M. S. & Bader, D. M. Bves modulates epithelial integrity through an interaction at the tight junction. J. Cell Sci. 118, 4667–4678 (2005).

    CAS  PubMed  Google Scholar 

  11. Luissint, A. C., Nusrat, A. & Parkos, C. A. JAM-related proteins in mucosal homeostasis and inflammation. Semin. Immunopathol. 36, 211–226 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. Martin-Padura, I. et al. Junctional adhesion molecule, a novel member of the immunoglobulin superfamily that distributes at intercellular junctions and modulates monocyte transmigration. J. Cell Biol. 142, 117–127 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Cohen, C. J. et al. The coxsackievirus and adenovirus≈receptor is a transmembrane component of the tight junction. Proc. Natl Acad. Sci. USA 98, 15191–15196 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Higashi, T. et al. Analysis of the 'angulin' proteins LSR, ILDR1 and ILDR2—tricellulin recruitment, epithelial barrier function and implication in deafness pathogenesis. J. Cell Sci. 126, 966–977 (2013).

    CAS  PubMed  Google Scholar 

  15. Masuda, S. et al. LSR defines cell corners for tricellular tight junction formation in epithelial cells. J. Cell Sci. 124, 548–555 (2011).

    CAS  PubMed  Google Scholar 

  16. Lemmers, C. et al. hINADl/PATJ, a homolog of discs lost, interacts with crumbs and localizes to tight junctions in human epithelial cells. J. Biol. Chem. 277, 25408–25415 (2002).

    CAS  PubMed  Google Scholar 

  17. Makarova, O., Roh, M. H., Liu, C. J., Laurinec, S. & Margolis, B. Mammalian Crumbs3 is a small transmembrane protein linked to protein associated with Lin-7 (Pals1). Gene 302, 21–29 (2003).

    CAS  PubMed  Google Scholar 

  18. Van Itallie, C. M. & Anderson, J. M. Architecture of tight junctions and principles of molecular composition. Semin. Cell Dev. Biol. 36, 157–165 (2014).

    CAS  PubMed  Google Scholar 

  19. Stevenson, B. R., Siliciano, J. D., Mooseker, M. S. & Goodenough, D. A. Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J. Cell Biol. 103, 755–766 (1986). Identification of the first tight junction protein.

    CAS  PubMed  Google Scholar 

  20. Rodgers, L. S., Beam, M. T., Anderson, J. M. & Fanning, A. S. Epithelial barrier assembly requires coordinated activity of multiple domains of the tight junction protein ZO-1. J. Cell Sci. 126, 1565–1575 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. Balda, M. S. & Anderson, J. M. Two classes of tight junctions are revealed by ZO-1 isoforms. Am. J. Physiol. 264, C918–C924 (1993).

    CAS  PubMed  Google Scholar 

  22. Fanning, A. S., Jameson, B. J., Jesaitis, L. A. & Anderson, J. M. The tight junction protein ZO-1 establishes a link between the transmembrane protein occludin and the actin cytoskeleton. J. Biol. Chem. 273, 29745–29753 (1998).

    CAS  PubMed  Google Scholar 

  23. Balda, M. S. & Matter, K. The tight junction protein ZO-1 and an interacting transcription factor regulate ErbB-2 expression. EMBO J. 19, 2024–2033 (2000). Identification of the first transcription factor that is regulated by tight junctions.

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Tsapara, A., Matter, K. & Balda, M. S. The heat-shock protein Apg-2 binds to the tight junction protein ZO-1 and regulates transcriptional activity of ZONAB. Mol. Biol. Cell 17, 1322–1330 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. Schmidt, A. et al. Occludin binds to the SH3-hinge-GuK unit of zonula occludens protein 1: potential mechanism of tight junction regulation. Cell. Mol. Life Sci. 61, 1354–1365 (2004).

    CAS  PubMed  Google Scholar 

  26. Lye, M. F., Fanning, A. S., Su, Y., Anderson, J. M. & Lavie, A. Insights into regulated ligand binding sites from the structure of ZO-1 Src homology 3-guanylate kinase module. J. Biol. Chem. 285, 13907–13917 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Fanning, A. S. et al. The unique-5 and -6 motifs of ZO-1 regulate tight junction strand localization and scaffolding properties. Mol. Biol. Cell 18, 721–731 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  28. Spadaro, D. et al. ZO proteins redundantly regulate the transcription factor DbpA/ZONAB. J. Biol. Chem. 289, 22500–22511 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. Gumbiner, B., Lowenkopf, T. & Apatira, D. Identification of a 160-kDa polypeptide that binds to the tight junction protein ZO-1. Proc. Natl Acad. Sci. USA 88, 3460–3464 (1991).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Balda, M. S., Gonzalez-Mariscal, L., Matter, K., Cereijido, M. & Anderson, J. M. Assembly of the tight junction: the role of diacylglycerol. J. Cell Biol. 123, 293–302 (1993).

    CAS  PubMed  Google Scholar 

  31. Haskins, J., Gu, L., Wittchen, E. S., Hibbard, J. & Stevenson, B. R. ZO-3, a novel member of the MAGUK protein family found at the tight junction, interacts with ZO-1 and occludin. J. Cell Biol. 141, 199–208 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. Ide, N. et al. Localization of membrane-associated guanylate kinase (MAGI)-1/BAI-associated protein (BAP) 1 at tight junctions of epithelial cells. Oncogene 18, 7810–7815 (1999).

    CAS  PubMed  Google Scholar 

  33. Dobrosotskaya, I., Guy, R. K. & James, G. L. MAGI-1, a membrane-associated guanylate kinase with a unique arrangement of protein-protein interaction domains. J. Biol. Chem. 272, 31589–31597 (1997).

    CAS  PubMed  Google Scholar 

  34. Poliak, S., Matlis, S., Ullmer, C., Scherrer, S. S. & Peles, E. Distinct claudins and associated PDZ proteins form different autotypic tight junctions in myelinating Schwann cells. J. Cell Biol. 159, 361–372 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. Roh, M. H. et al. The Maguk protein, Pals1, functions as an adapter, linking mammalian homologues of Crumbs and Discs Lost. J. Cell Biol. 157, 161–172 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  36. Hamazaki, Y., Itoh, M., Sasaki, H., Furuse, M. & Tsukita, S. Multi-PDZ domain protein 1 (MUPP1) is concentrated at tight junctions through its possible interaction with claudin-1 and junctional adhesion molecule. J. Biol. Chem. 277, 455–461 (2002).

    CAS  PubMed  Google Scholar 

  37. Citi, S., Pulimeno, P. & Paschoud, S. Cingulin, paracingulin, and PLEKHA7: signaling and cytoskeletal adaptors at the apical junctional complex. Ann. NY Acad. Sci. 1257, 125–132 (2012).

    CAS  PubMed  Google Scholar 

  38. Yano, T., Matsui, T., Tamura, A., Uji, M. & Tsukita, S. The association of microtubules with tight junctions is promoted by cingulin phosphorylation by AMPK. J. Cell Biol. 203, 605–614 (2013). Demonstration of a direct link between tight junctions and microtubules.

    CAS  PubMed  PubMed Central  Google Scholar 

  39. Stevenson, B. R., Heintzelman, M. B., Anderson, J. M., Citi, S. & Mooseker, M. S. ZO-1 and cingulin: tight junction proteins with distinct identities and localizations. Am. J. Physiol. 257, C621–C628 (1989).

    CAS  PubMed  Google Scholar 

  40. Cordenonsi, M. et al. Cingulin contains globular and coiled-coil domains and interacts with ZO-1, ZO-2, ZO-3, and myosin. J. Cell Biol. 147, 1569–1582 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Steed, E. et al. MarvelD3 couples tight junctions to the MEKK1–JNK pathway to regulate cell behavior and survival. J. Cell Biol. 204, 821–838 (2014). Elucidation of a mechanism connecting tight junctions and JNK signalling that regulates the cellular stress response.

    CAS  PubMed  PubMed Central  Google Scholar 

  42. Fredriksson, K. et al. Proteomic analysis of proteins surrounding occludin and claudin-4 reveals their proximity to signaling and trafficking networks. PLoS ONE 10, e0117074 (2015).

    PubMed  PubMed Central  Google Scholar 

  43. Zihni, C., Balda, M. S. & Matter, K. Signalling at tight junctions during epithelial differentiation and microbial pathogenesis. J. Cell Sci. 127, 3401–3413 (2014).

    CAS  PubMed  Google Scholar 

  44. Gonzalez-Mariscal, L. et al. Tight junctions and the regulation of gene expression. Semin. Cell Dev. Biol. 36, 213–223 (2014).

    CAS  PubMed  Google Scholar 

  45. Quiros, M. & Nusrat, A. RhoGTPases, actomyosin signaling and regulation of the epithelial apical junctional complex. Semin. Cell Dev. Biol. 36, 194–203 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  46. Adachi, M. et al. Normal establishment of epithelial tight junctions in mice and cultured cells lacking expression of ZO-3, a tight-junction MAGUK protein. Mol. Cell. Biol. 26, 9003–9015 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. Guillemot, L. et al. Cingulin is dispensable for epithelial barrier function and tight junction structure, and plays a role in the control of claudin-2 expression and response to duodenal mucosa injury. J. Cell Sci. 125, 5005–5014 (2012).

    CAS  PubMed  Google Scholar 

  48. Xu, J. et al. Early embryonic lethality of mice lacking ZO-2, but Not ZO-3, reveals critical and nonredundant roles for individual zonula occludens proteins in mammalian development. Mol. Cell. Biol. 28, 1669–1678 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  49. Katsuno, T. et al. Deficiency of ZO-1 causes embryonic lethal phenotype associated with defected yolk sac angiogenesis and apoptosis of embryonic cells. Mol. Biol. Cell 19, 2465–2475 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  50. Tokuda, S., Higashi, T. & Furuse, M. ZO-1 knockout by TALEN-mediated gene targeting in MDCK cells: involvement of ZO-1 in the regulation of cytoskeleton and cell shape. PLoS ONE 9, e104994 (2014).

    PubMed  PubMed Central  Google Scholar 

  51. Kiener, T. K., Selptsova-Friedrich, I. & Hunziker, W. Tjp3/zo-3 is critical for epidermal barrier function in zebrafish embryos. Dev. Biol. 316, 36–49 (2008).

    CAS  PubMed  Google Scholar 

  52. Mir, H. et al. Occludin deficiency promotes ethanol-induced disruption of colonic epithelial junctions, gut barrier dysfunction and liver damage in mice. Biochim. Biophys. Acta 1860, 765–774 (2016). References 48–52 provide striking examples of specific physiological roles for tight junction proteins that are often erroneously considered as being redundant or not crucial for barrier function.

    CAS  PubMed  Google Scholar 

  53. Suzuki, H. et al. Crystal structure of a claudin provides insight into the architecture of tight junctions. Science 344, 304–307 (2014). Determination of the structure of a claudin, enabling more-detailed modelling of the structure of tight junctions and the possible route of ion permeation.

    CAS  PubMed  Google Scholar 

  54. Suzuki, H., Tani, K., Tamura, A., Tsukita, S. & Fujiyoshi, Y. Model for the architecture of claudin-based paracellular ion channels through tight junctions. J. Mol. Biol. 427, 291–297 (2015).

    CAS  PubMed  Google Scholar 

  55. Furuse, M., Sasaki, H. & Tsukita, S. Manner of interaction of heterogeneous claudin species within and between tight junction strands. J. Cell Biol. 147, 891–903 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  56. Tsukita, S. & Furuse, M. Occludin and claudins in tight-junction strands: leading or supporting players? Trends Cell Biol. 9, 268–273 (1999).

    CAS  PubMed  Google Scholar 

  57. Haseloff, R. F., Dithmer, S., Winkler, L., Wolburg, H. & Blasig, I. E. Transmembrane proteins of the tight junctions at the blood–brain barrier: structural and functional aspects. Semin. Cell Dev. Biol. 38, 16–25 (2015).

    CAS  PubMed  Google Scholar 

  58. Shen, L., Weber, C. R. & Turner, J. R. The tight junction protein complex undergoes rapid and continuous molecular remodeling at steady state. J. Cell Biol. 181, 683–695 (2008). Demonstration that tight junctions are not a rigid complex and that different junctional proteins have distinct dynamic properties.

    CAS  PubMed  PubMed Central  Google Scholar 

  59. Pinto da Silva, P. & Kachar, B. On tight junction structure. Cell 28, 441–450 (1982).

    CAS  PubMed  Google Scholar 

  60. Chernomordik, L. V. & Kozlov, M. M. Membrane hemifusion: crossing a chasm in two leaps. Cell 123, 375–382 (2005).

    CAS  PubMed  Google Scholar 

  61. Kachar, B. & Reese, T. S. Evidence for the lipidic nature of tight junction strands. Nature 296, 464–466 (1982).

    CAS  PubMed  Google Scholar 

  62. Kan, F. W. Cytochemical evidence for the presence of phospholipids in epithelial tight junction strands. J. Histochem. Cytochem. 41, 649–656 (1993).

    CAS  PubMed  Google Scholar 

  63. Fujimoto, K. Freeze-fracture replica electron microscopy combined with SDS digestion for cytochemical labeling of intergral membrane proteins: aplication to the immunogold labeling of intercellular junctional complex. J. Cell Sci. 108, 3443–3449 (1995).

    CAS  PubMed  Google Scholar 

  64. Cording, J. et al. In tight junctions, claudins regulate the interactions between occludin, tricellulin and marvelD3, which, inversely, modulate claudin oligomerization. J. Cell Sci. 126, 554–564 (2013).

    CAS  PubMed  Google Scholar 

  65. van Meer, G., Gumbiner, B. & Simons, K. The tight junction does not allow lipid molecules to diffuse from one epithelial cell to the next. Nature 322, 639–641 (1986).

    CAS  PubMed  Google Scholar 

  66. Grebenkamper, K. & Galla, H. J. Translational diffusion measurements of a fluorescent phospholipid between MDCK-I cells support the lipid model of the tight junctions. Chem. Phys. Lipids 71, 133–143 (1994).

    CAS  PubMed  Google Scholar 

  67. Nusrat, A. et al. Tight junctions are membrane microdomains. J. Cell Sci. 113, 1771–1781 (2000).

    CAS  PubMed  Google Scholar 

  68. Lambert, D., O'Neill, C. A. & Padfield, P. J. Depletion of Caco-2 cell cholesterol disrupts barrier function by altering the detergent solubility and distribution of specific tight-junction proteins. Biochem. J. 387, 553–560 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Lynch, R. D. et al. Cholesterol depletion alters detergent-specific solubility profiles of selected tight junction proteins and the phosphorylation of occludin. Exp. Cell Res. 313, 2597–2610 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  70. Calderon, V. et al. Tight junctions and the experimental modifications of lipid content. J. Membr. Biol. 164, 59–69 (1998).

    CAS  PubMed  Google Scholar 

  71. Francis, S. A. et al. Rapid reduction of MDCK cell cholesterol by methyl-beta-cyclodextrin alters steady state transepithelial electrical resistance. Eur. J. Cell Biol. 78, 473–484 (1999).

    CAS  PubMed  Google Scholar 

  72. Larre, I., Ponce, A., Franco, M. & Cereijido, M. The emergence of the concept of tight junctions and physiological regulation by ouabain. Semin. Cell Dev. Biol. 36, 149–156 (2014).

    CAS  PubMed  Google Scholar 

  73. Yu, A. S. Claudins and the kidney. J. Am. Soc. Nephrol. 26, 11–19 (2015).

    CAS  PubMed  Google Scholar 

  74. Yu, A. S. et al. Molecular basis for cation selectivity in claudin-2-based paracellular pores: identification of an electrostatic interaction site. J. Gen. Physiol. 133, 111–127 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. Lingaraju, A. et al. Conceptual barriers to understanding physical barriers. Semin. Cell Dev. Biol. 42, 13–21 (2015).

    PubMed  PubMed Central  Google Scholar 

  76. Steed, E., Balda, M. S. & Matter, K. Dynamics and functions of tight junctions. Trends Cell Biol. 20, 142–149 (2010).

    CAS  PubMed  Google Scholar 

  77. Simon, D. B. et al. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption. Science 285, 103–106 (1999). Identification of a tight junction component required for ion-specific paracellular diffusion.

    CAS  PubMed  Google Scholar 

  78. McCarthy, K. M. et al. Inducible expression of claudin-1-myc but not occludin-VSV-G results in aberrant tight junction strand formation in MDCK cells. J. Cell Sci. 113, 3387–3398 (2000).

    CAS  PubMed  Google Scholar 

  79. Furuse, M. et al. Claudin-based tight junctions are crucial for the mammalian epidermal barrier: a lesson from claudin-1-deficient mice. J. Cell Biol. 156, 1099–1111 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. Amasheh, S. et al. Claudin-2 expression induces cation-selective channels in tight junctions of epithelial cells. J. Cell Sci. 115, 4969–4976 (2002).

    CAS  PubMed  Google Scholar 

  81. Furuse, M., Furuse, K., Sasaki, H. & Tsukita, S. Conversion of zonulae occludentes from tight to leaky strand type by introducing claudin-2 into Madin–Darby canine kidney I cells. J. Cell Biol. 153, 263–272 (2001). Demonstration that the claudin composition of a junction determines transepithelial electrical resistance.

    CAS  PubMed  PubMed Central  Google Scholar 

  82. Van Itallie, C. M. et al. Two splice variants of claudin-10 in the kidney create paracellular pores with different ion selectivities. Am. J. Physiol. Renal Physiol. 291, F1288–F1299 (2006).

    CAS  PubMed  Google Scholar 

  83. Gunzel, D. et al. Claudin-10 exists in six alternatively spliced isoforms that exhibit distinct localization and function. J. Cell Sci. 122, 1507–1517 (2009).

    PubMed  Google Scholar 

  84. Krug, S. M. et al. Claudin-17 forms tight junction channels with distinct anion selectivity. Cell. Mol. Life Sci. 69, 2765–2778 (2012).

    CAS  PubMed  Google Scholar 

  85. Krug, S. M., Schulzke, J. D. & Fromm, M. Tight junction, selective permeability, and related diseases. Semin. Cell Dev. Biol. 36, 166–176 (2014).

    CAS  PubMed  Google Scholar 

  86. Colegio, O. R., Van Itallie, C. M., McCrea, H. J., Rahner, C. & Anderson, J. M. Claudins create charge-selective channels in the paracellular pathway between epithelial cells. Am. J. Physiol. Cell Physiol. 283, C142–C147 (2002). Establishes that the claudin repertoire of a cell is important for the ion selectivity of the paracellular pathway and provides evidence for the roles of the extracellular domains.

    CAS  PubMed  Google Scholar 

  87. Angelow, S. & Yu, A. S. Structure-function studies of claudin extracellular domains by cysteine-scanning mutagenesis. J. Biol. Chem. 284, 29205–29217 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  88. Gunzel, D. & Yu, A. S. Claudins and the modulation of tight junction permeability. Physiol. Rev. 93, 525–569 (2013).

    PubMed  PubMed Central  Google Scholar 

  89. Weber, C. R. et al. Claudin-2-dependent paracellular channels are dynamically gated. eLife 4, e09906 (2015). Examination of paracellular claudin pores using the patch clamp approach.

    PubMed  PubMed Central  Google Scholar 

  90. Tamura, A. et al. Loss of claudin-15, but not claudin-2, causes Na+ deficiency and glucose malabsorption in mouse small intestine. Gastroenterology 140, 913–923 (2011).

    CAS  PubMed  Google Scholar 

  91. Wada, M., Tamura, A., Takahashi, N. & Tsukita, S. Loss of claudins 2 and 15 from mice causes defects in paracellular Na+ flow and nutrient transport in gut and leads to death from malnutrition. Gastroenterology 144, 369–380 (2013).

    CAS  PubMed  Google Scholar 

  92. Wilcox, E. R. et al. Mutations in the gene encoding tight junction claudin-14 cause autosomal recessive deafness DFNB29. Cell 104, 165–172 (2001).

    CAS  PubMed  Google Scholar 

  93. Wen, H., Watry, D. D., Marcondes, M. C. & Fox, H. S. Selective decrease in paracellular conductance of tight junctions: role of the first extracellular domain of claudin-5. Mol. Cell. Biol. 24, 8408–8417 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  94. Krause, G., Protze, J. & Piontek, J. Assembly and function of claudins: Structure–function relationships based on homology models and crystal structures. Semin. Cell Dev. Biol. 42, 3–12 (2015).

    CAS  PubMed  Google Scholar 

  95. Kahle, K. T. et al. Paracellular Cl− permeability is regulated by WNK4 kinase: insight into normal physiology and hypertension. Proc. Natl Acad. Sci. USA 101, 14877–14882 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. Wilson, F. H. et al. Human hypertension caused by mutations in WNK kinases. Science 293, 1107–1112 (2001).

    CAS  PubMed  Google Scholar 

  97. Yamauchi, K. et al. Disease-causing mutant WNK4 increases paracellular chloride permeability and phosphorylates claudins. Proc. Natl Acad. Sci. USA 101, 4690–4694 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  98. Ohta, A. et al. Overexpression of human WNK1 increases paracellular chloride permeability and phosphorylation of claudin-4 in MDCKII cells. Biochem. Biophys. Res. Commun. 349, 804–808 (2006).

    CAS  PubMed  Google Scholar 

  99. Tatum, R. et al. WNK4 phosphorylates ser(206) of claudin-7 and promotes paracellular Cl(-) permeability. FEBS Lett. 581, 3887–3891 (2007).

    CAS  PubMed  Google Scholar 

  100. Dragsen, P. R., Blumenthal, R. & Handler, J. S. Membrane asymmetry in epithelia: is the tight junction a barrier to diffusion in the plasma membrane? Nature 294, 718–722 (1981).

    Google Scholar 

  101. van Meer, G. & Simons, K. The function of tight junctions in maintaining differences in lipid composition between the apical and the basolateral cell surface domains of MDCK cells. EMBO J. 5, 1455–1464 (1986).

    CAS  PubMed  PubMed Central  Google Scholar 

  102. Mandel, L. J., Bacallao, R. & Zampighi, G. Uncoupling of the molecular 'fence' and paracellular 'gate' functions in epithelial tight junctions. Nature 361, 552–555 (1993).

    CAS  PubMed  Google Scholar 

  103. Nava, P., Lopez, S., Arias, C. F., Islas, S. & Gonzalez-Mariscal, L. The rotavirus surface protein VP8 modulates the gate and fence function of tight junctions in epithelial cells. J. Cell Sci. 117, 5509–5519 (2004).

    CAS  PubMed  Google Scholar 

  104. Balda, M. S. et al. Functional dissociation of paracellular permeability and transepithelial electrical resistance and disruption of the apical–basolateral intramembrane diffusion barrier by expression of a mutant tight junction membrane protein. J. Cell Biol. 134, 1031–1049 (1996).

    CAS  PubMed  Google Scholar 

  105. Umeda, K. et al. ZO-1 and ZO-2 independently determine where claudins are polymerized in tight-junction strand formation. Cell 126, 741–754 (2006).

    CAS  PubMed  Google Scholar 

  106. Ikenouchi, J. et al. Lipid polarity is maintained in absence of tight junctions. J. Biol. Chem. 287, 9525–9533 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  107. Cereijido, M., Robbins, E. S., Dolan, W. J., Rotunno, C. A. & Sabatini, D. D. Polarized monolayers formed by epithelial cells on a permeable and translucent support. J. Cell Biol. 77, 853–880 (1978). Establishes the now commonly used model of culturing epithelial cells on a permeable support.

    CAS  PubMed  Google Scholar 

  108. Balda, M. S. et al. Assembly and sealing of tight junctions: possible participation of G-proteins, phospholipase C, protein kinase C and calmodulin. J. Mem. Biol. 122, 193–202 (1991).

    CAS  Google Scholar 

  109. Rajasekaran, A. K., Hojo, M., Huima, T. & Rodriguez-Boulan, E. Catenins and zonula occludens-1 form a complex during early stages in the assembly of tight junctions. J. Cell Biol. 132, 451–463 (1996).

    CAS  PubMed  Google Scholar 

  110. Maiers, J. L., Peng, X., Fanning, A. S. & DeMali, K. A. ZO-1 recruitment to α-catenin — a novel mechanism for coupling the assembly of tight junctions to adherens junctions. J. Cell Sci. 126, 3904–3915 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. Fukuhara, A. et al. Involvement of nectin in the localization of junctional adhesion molecule at tight junctions. Oncogene 21, 7642–7655 (2002).

    CAS  PubMed  Google Scholar 

  112. Garrido-Urbani, S., Bradfield, P. F. & Imhof, B. A. Tight junction dynamics: the role of junctional adhesion molecules (JAMs). Cell Tissue Res. 355, 701–715 (2014).

    CAS  PubMed  Google Scholar 

  113. Herrmann, J. R. & Turner, J. R. Beyond Ussing's chambers: contemporary thoughts on integration of transepithelial transport. Am. J. Physiol. Cell Physiol. 310, C423–C431 (2016).

    PubMed  PubMed Central  Google Scholar 

  114. Terry, S. J. et al. Spatially restricted activation of RhoA signalling at epithelial junctions by p114RhoGEF drives junction formation and morphogenesis. Nat. Cell Biol. 13, 159–166 (2011). Identification of a molecular mechanism that mediates tight junction-specific RHOA and myosin activation, and thereby the formation of functional barriers.

    CAS  PubMed  PubMed Central  Google Scholar 

  115. Itoh, M., Tsukita, S., Yamazaki, Y. & Sugimoto, H. Rho GTP exchange factor ARHGEF11 regulates the integrity of epithelial junctions by connecting ZO-1 and RhoA–myosin II signaling. Proc. Natl Acad. Sci. USA 109, 9905–9910 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  116. Tornavaca, O. et al. ZO-1 controls endothelial adherens junctions, cell–cell tension, angiogenesis, and barrier formation. J. Cell Biol. 208, 821–838 (2015). Demonstration that tight junctions regulate angiogenesis and tension on adherens junctions.

    CAS  PubMed  PubMed Central  Google Scholar 

  117. 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).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Oda, Y., Otani, T., Ikenouchi, J. & Furuse, M. Tricellulin regulates junctional tension of epithelial cells at tricellular contacts through Cdc42. J. Cell Sci. 127, 4201–4212 (2014). Determination of a regulatory function of tricellulin involving CDC42 and the control of tension between tricellular corners.

    CAS  PubMed  Google Scholar 

  119. Schluter, M. A. & Margolis, B. Apicobasal polarity in the kidney. Exp. Cell Res. 318, 1033–1039 (2012).

    PubMed  PubMed Central  Google Scholar 

  120. Itoh, M. et al. Junctional adhesion molecule (JAM) binds to PAR-3: a possible mechanism for the recruitment of PAR-3 to tight junctions. J. Cell Biol. 154, 491–497 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. Ebnet, K., Iden, S., Gerke, V. & Suzuki, A. Regulation of epithelial and endothelial junctions by PAR proteins. Front. Biosci. 13, 6520–6536 (2008).

    CAS  PubMed  Google Scholar 

  122. Liu, X. F., Ishida, H., Raziuddin, R. & Miki, T. Nucleotide exchange factor ECT2 interacts with the polarity protein complex Par6/Par3/protein kinase Cζ (PKCζ) and regulates PKCζ activity. Mol. Cell. Biol. 24, 6665–6675 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  123. Wells, C. D. et al. A Rich1/Amot complex regulates the Cdc42 GTPase and apical-polarity proteins in epithelial cells. Cell 125, 535–548 (2006).

    CAS  PubMed  Google Scholar 

  124. Elbediwy, A. et al. Epithelial junction formation requires confinement of Cdc42 activity by a novel SH3BP1 complex. J. Cell Biol. 198, 677–693 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  125. Zihni, C. et al. Dbl3 drives Cdc42 signaling at the apical margin to regulate junction position and apical differentiation. J. Cell Biol. 204, 111–127 (2014). Elucidation of the molecular mechanism that drives polarized CDC42 activation at the apical pole.

    CAS  PubMed  PubMed Central  Google Scholar 

  126. Morais- de-Sa, E., Mirouse, V. & St Johnston, D. aPKC phosphorylation of Bazooka defines the apical/lateral border in Drosophila epithelial cells. Cell 141, 509–523 (2010).

    Google Scholar 

  127. Walther, R. F. & Pichaud, F. Crumbs/DaPKC-dependent apical exclusion of Bazooka promotes photoreceptor polarity remodeling. Curr. Biol. 20, 1065–1074 (2010).

    CAS  PubMed  Google Scholar 

  128. Michel, D. et al. PATJ connects and stabilizes apical and lateral components of tight junctions in human intestinal cells. J. Cell Sci. 118, 4049–4057 (2005).

    CAS  PubMed  Google Scholar 

  129. Adachi, M. et al. Similar and distinct properties of MUPP1 and Patj, two homologous PDZ domain-containing tight-junction proteins. Mol. Cell. Biol. 29, 2372–2389 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  130. Roh, M. H., Liu, C. J., Laurinec, S. & Margolis, B. The carboxyl terminus of zona occludens-3 binds and recruits a mammalian homologue of discs lost to tight junctions. J. Biol. Chem. 277, 27501–27509 (2002).

    CAS  PubMed  Google Scholar 

  131. Lemmers, C. et al. CRB3 binds directly to Par6 and regulates the morphogenesis of the tight junctions in mammalian epithelial cells. Mol. Biol. Cell 15, 1324–1333 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  132. Ellenbroek, S. I., Iden, S. & Collard, J. G. Cell polarity proteins and cancer. Semin. Cancer Biol. 22, 208–215 (2012).

    CAS  PubMed  Google Scholar 

  133. Zhao, B. et al. Angiomotin is a novel Hippo pathway component that inhibits YAP oncoprotein. Genes Dev. 25, 51–63 (2011).

    PubMed  PubMed Central  Google Scholar 

  134. Oka, T. et al. Functional complexes between YAP2 and ZO-2 are PDZ domain-dependent, and regulate YAP2 nuclear localization and signalling. Biochem. J. 432, 461–472 (2010).

    CAS  PubMed  Google Scholar 

  135. Cravo, A. S. et al. Hippo pathway elements co-localize with occludin: a possible sensor system in pancreatic epithelial cells. Tissue Barriers 3, e1037948 (2015).

    PubMed  PubMed Central  Google Scholar 

  136. Lv, X. B. et al. PARD3 induces TAZ activation and cell growth by promoting LATS1 and PP1 interaction. EMBO Rep. 16, 975–985 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  137. Yi, C. et al. A tight junction-associated Merlin–angiomotin complex mediates Merlin's regulation of mitogenic signaling and tumor suppressive functions. Cancer Cell 19, 527–540 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  138. Zhang, N. et al. The Merlin/NF2 tumor suppressor functions through the YAP oncoprotein to regulate tissue homeostasis in mammals. Dev. Cell 19, 27–38 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  139. Balda, M. S., Garrett, M. D. & Matter, K. The ZO-1-associated Y-box factor ZONAB regulates epithelial cell proliferation and cell density. J. Cell Biol. 160, 423–432 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  140. Nie, M., Aijaz, S., Leefa Chong San, I. V., Balda, M. S. & Matter, K. The Y-box factor ZONAB/DbpA associates with GEF-H1/Lfc and mediates Rho-stimulated transcription. EMBO Rep. 10, 1125–1131 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  141. Kavanagh, E. et al. Functional interaction between the ZO-1-interacting transcription factor ZONAB/DbpA and the RNA processing factor symplekin. J. Cell Sci. 119, 5098–5105 (2006).

    CAS  PubMed  Google Scholar 

  142. Frankel, P. et al. RalA interacts with ZONAB in a cell density-dependent manner and regulates its transcriptional activity. EMBO J. 24, 54–62 (2005).

    CAS  PubMed  Google Scholar 

  143. Ikari, A. et al. Nuclear distribution of claudin-2 increases cell proliferation in human lung adenocarcinoma cells. Biochim. Biophys. Acta 1843, 2079–2088 (2014).

    CAS  PubMed  Google Scholar 

  144. Pannequin, J. et al. Phosphatidylethanol accumulation promotes intestinal hyperplasia by inducing ZONAB-mediated cell density increase in response to chronic ethanol exposure. Mol. Cancer Res. 5, 1147–1157 (2007).

    CAS  PubMed  Google Scholar 

  145. Buchert, M. et al. Symplekin promotes tumorigenicity by up-regulating claudin-2 expression. Proc. Natl Acad. Sci. USA 107, 2628–2633 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  146. Sourisseau, T. et al. Regulation of PCNA and cyclin D1 expression and epithelial morphogenesis by the ZO-1-regulated transcription factor ZONAB/DbpA. Mol. Cell. Biol. 26, 2387–2398 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  147. Ruan, Y. C. et al. CFTR interacts with ZO-1 to regulate tight junction assembly and epithelial differentiation through the ZONAB pathway. J. Cell Sci. 127, 4396–4408 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  148. Russ, P. K. et al. Bves modulates tight junction associated signaling. PLoS ONE 6, e14563 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  149. Liu, L. B. et al. Bradykinin increased the permeability of BTB via NOS/NO/ZONAB-mediating down-regulation of claudin-5 and occludin. Biochem. Biophys. Res. Commun. 464, 118–125 (2015).

    CAS  PubMed  Google Scholar 

  150. Dominguez-Calderon, A. et al. ZO-2 silencing induces renal hypertrophy through a cell cycle mechanism and the activation of YAP and the mTOR pathway. Mol. Biol. Cell 27, 1581–1595 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  151. Li, D. & Mrsny, R. J. Oncogenic Raf-1 disrupts epithelial tight junctions via downregulation of occludin. J. Cell Biol. 148, 791–800 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Nusrat, A. et al. The coiled-coil domain of occludin can act to organize structural and functional elements of the epithelial tight junction. J. Biol. Chem. 275, 29816–29822 (2000).

    CAS  PubMed  Google Scholar 

  153. Barrios-Rodiles, M. et al. High-throughput mapping of a dynamic signaling network in mammalian cells. Science 307, 1621–1625 (2005).

    CAS  PubMed  Google Scholar 

  154. Lockwood, C., Zaidel-Bar, R. & Hardin, J. The C. elegans zonula occludens ortholog cooperates with the cadherin complex to recruit actin during morphogenesis. Curr. Biol. 18, 1333–1337 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Nie, M., Balda, M. S. & Matter, K. Stress- and Rho-activated ZO-1-associated nucleic acid binding protein binding to p21 mRNA mediates stabilization, translation, and cell survival. Proc. Natl Acad. Sci. USA 109, 10897–10902 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  156. Monteiro, A. C. et al. Trans-dimerization of JAM-A regulates Rap2 and is mediated by a domain that is distinct from the cis-dimerization interface. Mol. Biol. Cell 25, 1574–1585 (2014).

    PubMed  PubMed Central  Google Scholar 

  157. Severson, E. A., Lee, W. Y., Capaldo, C. T., Nusrat, A. & Parkos, C. A. Junctional adhesion molecule A interacts with Afadin and PDZ-GEF2 to activate Rap1A, regulate β1 integrin levels, and enhance cell migration. Mol. Biol. Cell 20, 1916–1925 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  158. Cera, M. R. et al. JAM-A promotes neutrophil chemotaxis by controlling integrin internalization and recycling. J. Cell Sci. 122, 268–277 (2009).

    CAS  PubMed  Google Scholar 

  159. McSherry, E. A., Brennan, K., Hudson, L., Hill, A. D. K. & Hopkins, A. M. Breast cancer cell migration is regulated through junctional adhesion molecule-A-mediated activation of Rap1 GTPase. Breast Cancer Res. 13, R31 (2011). References 156–159 establish JAMA as a regulator of focal adhesions and adherens junctions through RAP GTPases.

    CAS  PubMed  PubMed Central  Google Scholar 

  160. Kuo, J. C. et al. Analysis of the myosin-II-responsive focal adhesion proteome reveals a role for β-Pix in negative regulation of focal adhesion maturation. Nat. Cell Biol. 13, 383–393 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  161. Ren, Y., Li, R., Zheng, Y. & Busch, H. Cloning and characterization of GEF-H1, a microtubule-associated guanine nucleotide exchange factor for Rac and Rho GTPases. J. Biol. Chem. 273, 34954–34960 (1998).

    CAS  PubMed  Google Scholar 

  162. Krendel, M., Zenke, F. T. & Bokoch, G. M. Nucleotide exchange factor GEF-H1 mediates cross-talk between microtubules and the actin cytoskeleton. Nat. Cell Biol. 4, 294–301 (2002).

    CAS  PubMed  Google Scholar 

  163. 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). Identification of a mechanism by which tight junctions contribute to the downregulation of cellular RHOA signalling on formation of mature monolayers.

    CAS  PubMed  Google Scholar 

  164. Huang, I. H. et al. GEF-H1 controls focal adhesion signaling that regulates mesenchymal stem cell lineage commitment. J. Cell Sci. 127, 4186–4200 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  165. Guilluy, C. et al. The Rho GEFs LARG and GEF-H1 regulate the mechanical response to force on integrins. Nat. Cell Biol. 13, 722–727 (2011).

    PubMed  PubMed Central  Google Scholar 

  166. Tiwari-Woodruff, S. K. et al. OSP/claudin-11 forms a complex with a novel member of the tetraspanin super family and β1 integrin and regulates proliferation and migration of oligodendrocytes. J. Cell Biol. 153, 295–305 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  167. Lu, Z. et al. A non-tight junction function of claudin-7-Interaction with integrin signaling in suppressing lung cancer cell proliferation and detachment. Mol. Cancer 14, 120 (2015).

    PubMed  PubMed Central  Google Scholar 

  168. Dhawan, P. et al. Claudin-1 regulates cellular transformation and metastatic behavior in colon cancer. J. Clin. Invest. 115, 1765–1776 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  169. Peddibhotla, S. S. et al. Tetraspanin CD9 links junctional adhesion molecule-A to αvβ3 integrin to mediate basic fibroblast growth factor-specific angiogenic signaling. Mol. Biol. Cell 24, 933–944 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  170. Naik, M. U. & Naik, U. P. Junctional adhesion molecule-A-induced endothelial cell migration on vitronectin is integrin αvβ3 specific. J. Cell Sci. 119, 490–499 (2006).

    CAS  PubMed  Google Scholar 

  171. Izumi, Y. & Furuse, M. Molecular organization and function of invertebrate occluding junctions. Semin. Cell Dev. Biol. 36, 186–193 (2014).

    CAS  PubMed  Google Scholar 

  172. Simske, J. S. & Hardin, J. Claudin family proteins in Caenorhabditis elegans. Methods Mol. Biol. 762, 147–169 (2011).

    CAS  PubMed  Google Scholar 

  173. Wu, V. M., Schulte, J., Hirschi, A., Tepass, U. & Beitel, G. J. Sinuous is a Drosophila claudin required for septate junction organization and epithelial tube size control. J. Cell Biol. 164, 313–323 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  174. Wu, V. M. et al. Drosophila Varicose, a member of a new subgroup of basolateral MAGUKs, is required for septate junctions and tracheal morphogenesis. Development 134, 999–1009 (2007).

    CAS  PubMed  Google Scholar 

  175. Behr, M., Riedel, D. & Schuh, R. The claudin-like megatrachea is essential in septate junctions for the epithelial barrier function in Drosophila. Dev. Cell 5, 611–620 (2003).

    CAS  PubMed  Google Scholar 

  176. Genova, J. L. & Fehon, R. G. Neuroglian, Gliotactin, and the Na+/K+ ATPase are essential for septate junction function in Drosophila. J. Cell Biol. 161, 979–989 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  177. Nelson, K. S., Furuse, M. & Beitel, G. J. The Drosophila claudin Kune-kune is required for septate junction organization and tracheal tube size control. Genetics 185, 831–839 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  178. Asano, A., Asano, K., Sasaki, H., Furuse, M. & Tsukita, S. Claudins in Caenorhabditis elegans: their distribution and barrier function in the epithelium. Curr. Biol. 13, 1042–1046 (2003).

    CAS  PubMed  Google Scholar 

  179. Suzuki, A. & Ohno, S. The PAR–aPKC system: lessons in polarity. J. Cell Sci. 119, 979–987 (2006).

    CAS  PubMed  Google Scholar 

  180. Matter, K. & Balda, M. S. Signalling to and from tight junctions. Nat. Rev. Mol. Cell. Biol. 4, 225–236 (2003).

    CAS  PubMed  Google Scholar 

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Acknowledgements

The authors are supported by the UK Medical Research Council, the UK Biotechnology and Biological Sciences Research Council, Fight for Sight and the Wellcome Trust.

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Authors and Affiliations

  1. Department of Cell Biology, UCL Institute of Ophthalmology, University College London, Bath Street, London, EC1V 9EL, UK

    Ceniz Zihni, Clare Mills, Karl Matter & Maria S. Balda

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  1. Ceniz Zihni
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Correspondence to Karl Matter or Maria S. Balda.

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Glossary

Desmosomes

Adhesive structures, also known as maculae adhaerentes, formed from dense protein plaques of two adjacent cells, with associated intermediate filaments and transmembrane proteins, which belong to cadherin family.

MARVEL domain

A four-transmembrane helix module that has been identified in proteins of various families, many of which are associated with cholesterol-rich membrane microdomains.

Immunoglobulin-like domains

Protein domains consisting of a double-layer sandwich of seven to nine antiparallel β-stands arranged in two β-sheets.

Osmoregulation

A process used by cells and simple organisms to maintain fluid and electrolyte balance with their immediate environment.

Lipid micelles

Lipid molecules arranged in a spherical form in aqueous solutions as a result of the amphipathic nature of fatty acids, meaning that they contain a hydrophilic, polar head group and a long hydrophobic chain.

Patch clamp approach

An electrophysiology technique that allows the study of single and multiple ion channels in membranes.

Homology models

Comparative modelling of a protein through construction of an atomic-resolution model of the 'test' protein from its amino acid sequence and a resolved three-dimensional structure of a related homologous protein that is used as a template.

Brush border

The specialized apical membrane of absorptive epithelial cells, such as enterocytes. It is covered with regularly shaped microvilli: finger-like plasma membrane projections with a core formed by the actin cytoskeleton.

Focal adhesions

Large, dynamic protein complexes that link the cytoskeleton of a cell to the extracellular matrix.

Small GTPases

Small, monomeric proteins that are homologous to RAS. They exist in an inactive GDP-bound form and an active GTP-bound form in which they activate other signalling proteins.

Heterotrimeric GTPases

(Also called G proteins). These consist of three subunits: the GTP-binding α-subunit and the smaller β- and γ-subunits, which have regulatory and signalling functions.

Guanine nucleotide exchange factors

(GEFs). Proteins that activate monomeric GTPases by stimulating the dissociation of GDP, thereby permitting binding of GTP.

Hypertonic stress

A phenomenon experienced by cells and tissues when the extracellular-fluid osmolarity exceeds that of the intracellular fluid.

Stress fibres

Contractile actin bundles in non-muscle cells. They consist of actin microfilaments, myosin II and crosslinkers such as α-actinin.

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Zihni, C., Mills, C., Matter, K. et al. Tight junctions: from simple barriers to multifunctional molecular gates. Nat Rev Mol Cell Biol 17, 564–580 (2016). https://doi.org/10.1038/nrm.2016.80

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