Student or educator? Sign up with your school email and get 6 months of Pro free.
Cited by 1,496

The DOI matches its Crossref record; 8 of the 8 references it cites were found in a registry or an index.

Tight junctions: from simple barriers to multifunctional molecular gates

Zihni, Ceniz, Mills, Clare, Matter, Karl, Balda, Maria S.·2016·Nature Reviews Molecular Cell Biology

Overview

The claims and statistics of this paper haven’t been checked yet.

Run the check — free, no account needed. It reads the open-access text and checks the sources it cites.

A few free checks a day · papers already verified open instantly · results are public

Integrity signals

Registry

Crossref · 10.1038/nrm.2016.80

Cites

8 of 8 references verified0 citation errors · 0 not checkable

Statistics

Verification

Claims the paper makes

The claims and statistics of this paper haven’t been checked yet.

Run the check — free, no account needed. It reads the open-access text and checks the sources it cites.

A few free checks a day · papers already verified open instantly · results are public

Self-check

Percentages, sums and reported differences re-computed from the paper’s own numbers.

The self-check needs the paper’s full text.

Citations

Cites

Show all 8 references

Research works printed but not linked: 172

  1. Cereijido, M., Contreras, R. G. & Shoshani, L. Cell adhesion, polarity, and epithelia in the dawn of metazoans. Physiol. Rev. 84, 1229–1262 (2004). · DOI · Paper page
  2. Farquhar, M. G. & Palade, G. E. Junctional complexes in various epithelia. J. Cell Biol. 17, 375–412 (1963). · DOI · Paper page
  3. Claude, P. & Goodenough, D. A. Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia. J. Cell Biol. 58, 390–400 (1973). · DOI · Paper page
  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). · DOI · Paper page
  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). · DOI · Paper page
  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. · DOI · Paper page
  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). · DOI · Paper page
  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). · DOI · Paper page
  9. Van Itallie, C. M. & Anderson, J. M. Occludin confers adhesiveness when expressed in fibroblasts. J. Cell Sci. 110, 1113–1121 (1997). · DOI · Paper page
  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). · DOI
  11. Luissint, A. C., Nusrat, A. & Parkos, C. A. JAM-related proteins in mucosal homeostasis and inflammation. Semin. Immunopathol. 36, 211–226 (2014). · DOI
  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). · DOI · Paper page
  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). · DOI · Paper page
  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). · DOI · Paper page
  15. Masuda, S. et al. LSR defines cell corners for tricellular tight junction formation in epithelial cells. J. Cell Sci. 124, 548–555 (2011). · DOI · Paper page
  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). · DOI · Paper page
  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). · DOI · Paper page
  18. 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. · DOI · Paper page
  19. 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). · DOI
  20. Balda, M. S. & Anderson, J. M. Two classes of tight junctions are revealed by ZO-1 isoforms. Am. J. Physiol. 264, C918–C924 (1993). · DOI · Paper page
  21. 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). · DOI · Paper page
  22. 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. · DOI · Paper page
  23. 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). · DOI
  24. 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). · DOI
  25. 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). · DOI
  26. 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). · DOI
  27. Spadaro, D. et al. ZO proteins redundantly regulate the transcription factor DbpA/ZONAB. J. Biol. Chem. 289, 22500–22511 (2014). · DOI
  28. 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). · DOI · Paper page
  29. 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). · DOI · Paper page
  30. 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). · DOI · Paper page
  31. 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). · DOI
  32. 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). · DOI · Paper page
  33. 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). · DOI · Paper page
  34. 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). · DOI · Paper page
  35. 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). · DOI · Paper page
  36. 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). · DOI
  37. 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. · DOI
  38. 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). · DOI
  39. 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). · DOI · Paper page
  40. 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. · DOI
  41. 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). · DOI
  42. Zihni, C., Balda, M. S. & Matter, K. Signalling at tight junctions during epithelial differentiation and microbial pathogenesis. J. Cell Sci. 127, 3401–3413 (2014). · DOI
  43. Gonzalez-Mariscal, L. et al. Tight junctions and the regulation of gene expression. Semin. Cell Dev. Biol. 36, 213–223 (2014). · DOI
  44. Quiros, M. & Nusrat, A. RhoGTPases, actomyosin signaling and regulation of the epithelial apical junctional complex. Semin. Cell Dev. Biol. 36, 194–203 (2014). · DOI
  45. 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). · DOI
  46. 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). · DOI
  47. 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). · DOI · Paper page
  48. 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). · DOI · Paper page
  49. 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). · DOI
  50. 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). · DOI
  51. 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. · DOI
  52. 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. · DOI · Paper page
  53. 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). · DOI · Paper page
  54. 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). · DOI · Paper page
  55. Tsukita, S. & Furuse, M. Occludin and claudins in tight-junction strands: leading or supporting players? Trends Cell Biol. 9, 268–273 (1999). · DOI · Paper page
  56. 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). · DOI · Paper page
  57. 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. · DOI · Paper page
  58. Pinto da Silva, P. & Kachar, B. On tight junction structure. Cell 28, 441–450 (1982). · DOI · Paper page
  59. Chernomordik, L. V. & Kozlov, M. M. Membrane hemifusion: crossing a chasm in two leaps. Cell 123, 375–382 (2005). · DOI · Paper page
  60. Kachar, B. & Reese, T. S. Evidence for the lipidic nature of tight junction strands. Nature 296, 464–466 (1982). · DOI · Paper page
  61. Kan, F. W. Cytochemical evidence for the presence of phospholipids in epithelial tight junction strands. J. Histochem. Cytochem. 41, 649–656 (1993). · doi:10.1177/41.5.8468446
  62. 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). · DOI · Paper page
  63. 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). · DOI · Paper page
  64. 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). · DOI · Paper page
  65. 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). · doi:10.1016/0009-3084(94)90066-3
  66. Nusrat, A. et al. Tight junctions are membrane microdomains. J. Cell Sci. 113, 1771–1781 (2000). · DOI · Paper page
  67. 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). · DOI
  68. 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). · DOI
  69. Calderon, V. et al. Tight junctions and the experimental modifications of lipid content. J. Membr. Biol. 164, 59–69 (1998). · doi:10.1007/s002329900393
  70. 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). · DOI
  71. 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). · DOI
  72. Yu, A. S. Claudins and the kidney. J. Am. Soc. Nephrol. 26, 11–19 (2015). · DOI
  73. 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). · DOI · Paper page
  74. Lingaraju, A. et al. Conceptual barriers to understanding physical barriers. Semin. Cell Dev. Biol. 42, 13–21 (2015). · DOI
  75. Steed, E., Balda, M. S. & Matter, K. Dynamics and functions of tight junctions. Trends Cell Biol. 20, 142–149 (2010). · DOI · Paper page
  76. 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. · DOI · Paper page
  77. 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). · DOI · Paper page
  78. Amasheh, S. et al. Claudin-2 expression induces cation-selective channels in tight junctions of epithelial cells. J. Cell Sci. 115, 4969–4976 (2002). · DOI · Paper page
  79. 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. · DOI · Paper page
  80. 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). · DOI · Paper page
  81. 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). · DOI · Paper page
  82. Krug, S. M. et al. Claudin-17 forms tight junction channels with distinct anion selectivity. Cell. Mol. Life Sci. 69, 2765–2778 (2012). · DOI
  83. Krug, S. M., Schulzke, J. D. & Fromm, M. Tight junction, selective permeability, and related diseases. Semin. Cell Dev. Biol. 36, 166–176 (2014). · DOI · Paper page
  84. 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. · DOI · Paper page
  85. Angelow, S. & Yu, A. S. Structure-function studies of claudin extracellular domains by cysteine-scanning mutagenesis. J. Biol. Chem. 284, 29205–29217 (2009). · DOI
  86. 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. · DOI
  87. 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). · DOI · Paper page
  88. 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). · DOI · Paper page
  89. Wilcox, E. R. et al. Mutations in the gene encoding tight junction claudin-14 cause autosomal recessive deafness DFNB29. Cell 104, 165–172 (2001). · DOI · Paper page
  90. 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). · DOI · Paper page
  91. 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). · DOI
  92. 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). · DOI · Paper page
  93. Wilson, F. H. et al. Human hypertension caused by mutations in WNK kinases. Science 293, 1107–1112 (2001). · DOI · Paper page
  94. Yamauchi, K. et al. Disease-causing mutant WNK4 increases paracellular chloride permeability and phosphorylates claudins. Proc. Natl Acad. Sci. USA 101, 4690–4694 (2004). · DOI · Paper page
  95. 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). · DOI
  96. Tatum, R. et al. WNK4 phosphorylates ser(206) of claudin-7 and promotes paracellular Cl(-) permeability. FEBS Lett. 581, 3887–3891 (2007). · DOI
  97. 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). · DOI · Paper page
  98. 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). · DOI · Paper page
  99. 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). · DOI · Paper page
  100. 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). · DOI
  101. 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). · DOI · Paper page
  102. 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). · DOI · Paper page
  103. Ikenouchi, J. et al. Lipid polarity is maintained in absence of tight junctions. J. Biol. Chem. 287, 9525–9533 (2012). · DOI
  104. 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. · DOI · Paper page
  105. 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). · DOI · Paper page
  106. 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). · DOI · Paper page
  107. 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). · DOI
  108. Fukuhara, A. et al. Involvement of nectin in the localization of junctional adhesion molecule at tight junctions. Oncogene 21, 7642–7655 (2002). · DOI
  109. 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). · DOI · Paper page
  110. 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). · DOI
  111. 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. · DOI · Paper page
  112. 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). · DOI
  113. 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). · DOI · Paper page
  114. 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. · DOI
  115. Schluter, M. A. & Margolis, B. Apicobasal polarity in the kidney. Exp. Cell Res. 318, 1033–1039 (2012). · DOI
  116. 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). · DOI · Paper page
  117. Ebnet, K., Iden, S., Gerke, V. & Suzuki, A. Regulation of epithelial and endothelial junctions by PAR proteins. Front. Biosci. 13, 6520–6536 (2008). · DOI
  118. 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). · DOI
  119. 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). · DOI · Paper page
  120. Elbediwy, A. et al. Epithelial junction formation requires confinement of Cdc42 activity by a novel SH3BP1 complex. J. Cell Biol. 198, 677–693 (2012). · DOI
  121. 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. · DOI
  122. 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). · DOI · Paper page
  123. Walther, R. F. & Pichaud, F. Crumbs/DaPKC-dependent apical exclusion of Bazooka promotes photoreceptor polarity remodeling. Curr. Biol. 20, 1065–1074 (2010). · DOI
  124. 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). · DOI
  125. 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). · DOI
  126. 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). · DOI · Paper page
  127. 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). · DOI · Paper page
  128. Ellenbroek, S. I., Iden, S. & Collard, J. G. Cell polarity proteins and cancer. Semin. Cancer Biol. 22, 208–215 (2012). · DOI
  129. Zhao, B. et al. Angiomotin is a novel Hippo pathway component that inhibits YAP oncoprotein. Genes Dev. 25, 51–63 (2011). · DOI · Paper page
  130. 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). · DOI · Paper page
  131. 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). · DOI
  132. Lv, X. B. et al. PARD3 induces TAZ activation and cell growth by promoting LATS1 and PP1 interaction. EMBO Rep. 16, 975–985 (2015). · DOI
  133. 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). · DOI · Paper page
  134. 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). · DOI · Paper page
  135. 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). · DOI
  136. 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). · DOI
  137. 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). · DOI
  138. Ikari, A. et al. Nuclear distribution of claudin-2 increases cell proliferation in human lung adenocarcinoma cells. Biochim. Biophys. Acta 1843, 2079–2088 (2014). · DOI
  139. 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). · DOI
  140. Buchert, M. et al. Symplekin promotes tumorigenicity by up-regulating claudin-2 expression. Proc. Natl Acad. Sci. USA 107, 2628–2633 (2010). · DOI
  141. 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). · DOI · Paper page
  142. 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). · DOI
  143. Russ, P. K. et al. Bves modulates tight junction associated signaling. PLoS ONE 6, e14563 (2011). · DOI
  144. 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). · doi:10.1016/j.bbrc.2015.06.082
  145. 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). · DOI
  146. Li, D. & Mrsny, R. J. Oncogenic Raf-1 disrupts epithelial tight junctions via downregulation of occludin. J. Cell Biol. 148, 791–800 (2000). · DOI · Paper page
  147. 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). · DOI · Paper page
  148. Barrios-Rodiles, M. et al. High-throughput mapping of a dynamic signaling network in mammalian cells. Science 307, 1621–1625 (2005). · DOI · Paper page
  149. 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). · DOI
  150. 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). · DOI
  151. 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). · DOI
  152. 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). · DOI · Paper page
  153. Cera, M. R. et al. JAM-A promotes neutrophil chemotaxis by controlling integrin internalization and recycling. J. Cell Sci. 122, 268–277 (2009). · DOI
  154. 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. · DOI
  155. 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). · DOI · Paper page
  156. 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). · DOI
  157. 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). · DOI · Paper page
  158. 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). · DOI · Paper page
  159. 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). · DOI
  160. Dhawan, P. et al. Claudin-1 regulates cellular transformation and metastatic behavior in colon cancer. J. Clin. Invest. 115, 1765–1776 (2005). · DOI · Paper page
  161. 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). · DOI
  162. 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). · DOI
  163. Izumi, Y. & Furuse, M. Molecular organization and function of invertebrate occluding junctions. Semin. Cell Dev. Biol. 36, 186–193 (2014). · DOI
  164. Simske, J. S. & Hardin, J. Claudin family proteins in Caenorhabditis elegans. Methods Mol. Biol. 762, 147–169 (2011). · DOI
  165. 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). · DOI · Paper page
  166. 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). · DOI
  167. 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). · DOI · Paper page
  168. 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). · DOI · Paper page
  169. 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). · DOI
  170. 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). · DOI
  171. Suzuki, A. & Ohno, S. The PAR–aPKC system: lessons in polarity. J. Cell Sci. 119, 979–987 (2006). · DOI · Paper page
  172. Matter, K. & Balda, M. S. Signalling to and from tight junctions. Nat. Rev. Mol. Cell. Biol. 4, 225–236 (2003). · DOI · Paper page

Cited by

See all 1,496

Metadata

Authors
Zihni, C., Mills, C., Matter, K., & Balda, M. S.
Venue
Nature Reviews Molecular Cell Biology
Published
2016-06-29
Volume
17
Issue
9
Pages
564-580
ISSN
  • 1471-0072 (ISSN-L)
  • 1471-0080
OpenAlex
W2464915778
Cited by
1,496
References
8
FWCI
7.70
Type
journal article
Language
en
Open access
closed
Licence
www.springer.com/tdm (tdm)

About these checks

Registry, retraction, Cites and statistics results are computed, not judged, with one exception: a printed reference that no index lists is read once by a model before it counts as a citation error. Claims are extracted by a model and kept only when their quote appears verbatim in the text; each cited source is then checked, with a model’s help, for whether it says what the claim relies on. Those verdicts can be wrong — read the claim’s quote and the cited source before relying on one.

Statistics and arithmetic are recomputed, not judged.

Confidence bands are shown only for fields calibrated on our annotated test set. None is calibrated yet, so no band is shown.

What counts as a claim ›How we measure confidence ›How we check this ›

Are you an author, or do you see a problem with this page? Request a review

How to cite

Cite CoChat for the checks on this page, and the paper for its own findings.

This page
CoChat. (2026). Citation check: Tight junctions: from simple barriers to multifunctional molecular gates. https://app.cochat.ai/papers/10.1038/nrm.2016.80
The paper
Zihni, C., Mills, C., Matter, K., & Balda, M. S. (2016). Tight junctions: from simple barriers to multifunctional molecular gates. Nature Reviews Molecular Cell Biology. https://doi.org/10.1038/nrm.2016.80

Data sources

Bibliographic data: Semantic Scholar, OpenAlex, Crossref.

Contains information from the Semantic Scholar Academic Graph Datasets, made available under the ODC Attribution License (ODC-BY 1.0).

Semantic Scholar’s data is described in: Kinney, R., et al. (2023). The Semantic Scholar Open Data Platform. arXiv:2301.10140.