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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
IUBMB Life. Author manuscript; available in PMC 2010 April 1.
Published in final edited form as:
PMCID: PMC2708117

Regulation and roles for claudin-family tight junction proteins


Transmembrane proteins known as claudins play a critical role in tight junctions by regulating paracellular barrier permeability. The control of claudin assembly into tight junctions requires a complex interplay between several classes of claudins, other transmembrane proteins and scaffold proteins. Claudins are also subject to regulation by post-translational modifications including phosphorylation and palmitoylation. Several human diseases have been linked to claudin mutations, underscoring the physiologic function of these proteins. Roles for claudins in regulating cell phenotype and growth control also are beginning to emerge, suggesting a multifaceted role for claudins in regulation of cells beyond serving as a simple structural element of tight junctions.

Keywords: tight junction, claudin, occludin, zonula occludens, tetraspanin, epithelium, cancer


Epithelial and endothelial cell monolayers form barriers to separate organs into functional subcompartments. This distinct compartmentalization and isolation from the external environment is crucial for the function of organ systems in multicellular organisms. Cells control these selective barriers by regulating the movement of water, ions and proteins across the monolayer, thus generating polarity of cellular structure and function (1). The movement of ions and molecules between cells is known as paracellular permeability and is regulated by sites of cell-cell contact known as tight junctions, a complex of transmembrane and peripheral proteins that is tethered to the cytoskeleton. While tight junctions require the coordinated activity of several different proteins, the specificity of tight junction permeability is regulated by transmembrane proteins known as claudins (2), a name derived from the Latin claudere, which means “to close”. The primary role of claudins is the regulation of paracellular selectivity to small ions. Claudins form what are functionally equivalent to charge selective pores which promote specific ion permeability, although there are also examples of claudins which more generally increase rather than restrict paracellular permeability as well (3). The mechanisms which regulate claudin assembly and function are just beginning to be elucidated. Here we summarize current progress in understanding how claudins control paracellular permeability.

Claudin Structure

To date, 23 distinct human claudins have been identified (4). Claudins are 20–27kDa transmembrane proteins which span the bilayer four times, where the N- and C- termini are oriented towards the cytoplasm and there are two extracellular loop domains (5). For at least two claudins, claudin-10 and claudin-18, alternative splicing can also generate claudin isoforms with different properties (6, 7).

Little is known about function of the relatively short, 7-amino acid, N-terminus. The cytoplasmic C-terminus sequence varies considerably in length (from 21–63 residues) and sequence between isoforms. All claudins have C-terminal PDZ binding motifs that enables direct interaction with tight junction cytoplasmic proteins such as ZO-1,-2, and -3, multi-PDZ domain protein (MUPP)-1 and PALS-1 associated TJ protein (PATJ). In particular, interactions with the cytoplasmic scaffolding proteins ZO-1 and ZO-2 indirectly link claudins to the actin cytoskeleton which stabilizes the tight junction and is required to maintain their permeability characteristics (8).

The first extracellular loop of claudins is approximately 52 residues and influences paracellular charge selectivity (9). The first extracellular loop domain has a signature motif (Gly-Leu-Trp-x-x-Cys-(8-10aa)-Cys) which is highly conserved and is found in closely related proteins, such as epithelial membrane proteins (EMPs, PMP22, MP20) (4). The cysteines in this signature motif may form an intramolecular disulfide bond to stabilize protein conformation, however, this remains to be determined. The second extracellular loop of claudins is shorter (16–33 residues) and less well characterized, although molecular modeling suggests the second extracellular loop is folded in a helix-turn-helix motif which participates in claudin-claudin interactions (10).

Incorporation into tight junctions

Freeze fracture immunogold analysis has shown that tight junctions are composed of a network of strands of which claudins are the major transmembrane constituent (11). These strands are roughly the diameter of a gap junction channel, which has led to the suggestion that claudins are organized into a basic hexameric unit, similar to connexins in gap junctions. Consistent with this, claudins have been shown to form ladders of stable oligomers in native gel electrophoresis increasing in molecular mass up to a hexamer (12, 13). However, to date little is known about the mechanisms which regulate claudin oligomerization and whether this is a process which occurs prior to assembly into tight junction strands.


Signature di-cysteine palmitoylation motifs are conserved throughout the claudin protein family and were found to enhance incorporation of claudins into tight junctions (14). Palmitoylated claudins more efficiently portion into detergent resistant membranes as compared to non-palmitoylated mutants, suggesting lipid rafts play a role in tight junction assembly (15). Tetraspanins, such as CD9 and CD81, are also palmitoylated which is critical to their function as membrane chaperones and organizers of higher order membrane domains (16). A link between tetraspanins and claudins is suggested by the demonstration that CD9 directly interacts with claudin-1 in a cholesterol sensitive manner (17). Whether CD81 plays a role in regulating claudin assembly is not yet known, however, CD81 acts as a co-receptor with claudin-1, -6 or 9 to facilitate Hepatitis C binding and entry into cells (18). Also, it is unknown where claudins oligomerize however, since palmitoylation and tetraspanin oligomerization occurs predominantly in the Golgi Apparatus, this suggests the Golgi Apparatus as a potential location for claudin oligomerization. This would be analogous to the connexin family of gap junction proteins which oligomerize in the Trans Golgi Network (4).

Claudin-claudin interactions

Epithelia and endothelia typically express multiple claudin isoforms in a tissue specific manner, suggesting the potential of claudin isoforms to intermix in tight junction strands. The interactions between different classes of claudins has the potential to control barrier permeability through the formation of tight junctions composed of multiple claudins. Claudins interact in two different ways: laterally in the plane of the membrane (heteromeric interactions) or head to head binding between adjacent cells (heterotypic interactions). Very little is known about claudin heteromeric binding. Claudin homomultimers composed of up to six monomers have been observed biochemically (12, 13, 19) and lateral interactions of claudin-5 within cells has been detected using fluorescence resonance energy transfer (FRET) (20).

Cells with claudin-null backgrounds have helped define specificity in claudin hetertypic interactions (12, 19, 21). Using this approach, heterotypic and heteromeric compatibility have been found to be determined by distinct mechanisms. For example, claudin-3 and claudin-4 are heteromerically compatible when expressed in the same cell, however, they do not heterotypically interact despite having extracellular loop domains that are highly conserved at the amino acid level (21). By contrast, claudin-3 heterotypically interacts with other claudins, including claudin-1 and claudin-5 (12, 19, 21). However, a single point mutation in the first extracellular domain of claudin-3 to convert Asn(44) to the corresponding amino acid in claudin-4 (Thr) produced a claudin capable of heterotypic binding to claudin-4 while still retaining the ability to bind to claudin-1 and claudin-5. Thus far, the only claudin found to be capable of heterotypic binding appears to be claudin-3, suggesting that heterotypic interactions may be rare and that homotypic claudin interactions may be more common. Whether this is the case will require analysis of claudins beyond the handful already studied in detail.

Claudin-occludin interactions

Occludin is another tetraspan transmembrane protein associated with tight junctions (22). For many years occludin was assumed to be the protein which formed the physical basis of the tight junction barrier; however, transgenic occludin deficient mice are viable and have normal barrier function (23) which ultimately lead to the identification of claudins as the barrier forming component of tight junctions (24). Although occludin-deficiency does not impair barrier function, peptides that mimic the occludin extracellular loop domains can disrupt epithelial barrier function by enhancing junctional disassembly suggesting that occludin associates with tight junctions in a regulatory capacity (25). Regulation of barrier function is also suggested by studies where cells transfected with a C-terminal truncated occludin construct show increased paracellular flux of uncharged molecules (26). In this case, transepithelial resistance was not altered, suggesting the increase in paracellular flux was due to increased turnover (or treadmilling) of the tight junction strands (1), most likely due to a disruption of occludin-scaffold protein interactions.

Recently, a claudin-1 extracellular loop peptide was also found to disrupt tight junction assembly and function (27). Interestingly, the claudin-1 peptide was found to bind to occludin, suggesting a direct interaction (27). The structural basis for this interaction as well as whether this reflects a potential heterotypic or heteromeric interaction between claudins and occludin remains to be determined.

Claudin-scaffold protein interactions

Claudin-ZO protein interactions are required for tight junction assembly (8). This was demonstrated using cultured epithelial cells in where expression of both ZO-1 and ZO-2 was suppressed. In ZO-1/ZO-2 null cells, claudins failed to localize to tight junctions and the cells had poor barrier function. However, transfection with either ZO-1 or ZO-2 enabled proper claudin localization and function (8). Thus, ZO-1 and ZO-2 have overlapping function, which suggests that linking claudins to the actin cytoskeleton through these scaffold proteins is essential for tight junction formation.

Since most claudins have conserved C-terminal YV domain, they are likely to interact with zona occludens (ZO) -1, -2 and 3, as demonstrated for claudin-1 through claudin-8 (28). Claudin-1, -5 and 8 have also been shown to interact with MUPP1 and claudin-1 can bind PATJ, underscoring the multiplicity of scaffold proteins capable of interacting with claudins (29, 30). Claudin-scaffold interactions are also specified by other motifs in the C-terminal domain. This was demonstrated using a series of tail swap constructs, where tight junction strand count and morphology correlate with the claudin tail motif, whereas ion permeability characteristics depended on the remainder of the protein, most notably, the extracellular loop domains (31).


Claudin phosphorylation is another mode of regulating paracellular permeability. Phosphorylation has been linked to both increases and decreases in tight junction assembly and function. For instance, Protein Kinase A-mediated phosphorylation has been shown to decrease assembly of claudin-3 into tight junctions (32), yet is necessary for claudin-16 assembly and function (33). Claudin-1 and Claudin-4 phosphorylation by Protein Kinase C-theta is required for assembly into intestinal epithelial tight junctions (34). Conversely, PP2A phosphatase activity has been shown to decrease claudin-1 phosphorylation and barrier function, although in this case, ZO-1 phosphorylation was also reduced, so this may be reflect altered interactions between claudin-1 and ZO-1 (35). Consistent with the ability of a phosphatase to impair interactions with ZO-1 and the cytoskeleton, claudin-1 detergent solubility is enhanced by PP2A.

Conversely, phosphorylation via myosin light chain kinase (MLCK) and rho kinase is more frequently associated with tight junction disassembly and increased paracellular permeability associated with inflammaton and in response to environmental stresses such as chronic alcohol abuse (36, 37). In addition to increased permeability, it seems possible that increased junction turnover induced by MLCK and/or rho kinase might be a contributing factor in the changes in claudin composition observed under these conditions, although this remains to be determined (38, 39).

Claudin phosphorylation associated with tight junction disassembly is also enhanced by EphA1, which is recruited to bind to claudin-4 by forming a complex with ephrin-B1 (40). A direct role for EphA1 sand ephrin-B1 in interfering with claudin assembly is intriguing, given the role of ephrins in inducing epithelial to mesenchyme transition (EMT) (41), a process where tight junction protein expression is downregulated (42, 43).

Claudin remodeling and endocytosis

Although the network of tight junctions is constantly remodeled, the different components have different stability within the strands themselves (44, 45). Based on fluorescence recovery after photobleaching (FRAP), the majority of claudins are stably integrated into tight junctions, reflecting the strength of intercellular binding and association with the cytoskeleton (44, 45). In contrast, occludin and ZO-1 are more dynamic (45). The mobile fraction of occludin is nearly three times greater than that of claudin. ZO-1 is highly dynamic as well, although instead of diffusing along the plane of the membrane, it instead dissociates from tight junctions and diffuses into the cytoplasm. This is consistent with models where ZO-1 interactions with occludin and claudins are required to stabilize these proteins in junctions, in fact, the rate of diffusion for the rapidly mobile fractions of claudins and occludin are comparable (45).

Tight junctions are also remodeled at a more macroscopic level through strand breaks and reformation (44). Clathrin-mediated endocytosis plays an important role in this process (46, 47). Claudins are internalized by a unique mechanism, where the tightly opposed membranes of the tight junction are endocytosed together into one of the adjoining cells, a mechanism also used for gap junction internalization (48). During internalization, the claudins separate away from occludin, JAM and ZO-1 and generate claudin-enriched vesicles, which has the potential to regulate the claudin composition of tight junctions.

Effect of inflammation on claudins

In response to pathogens and during inflammation, increased claudin internalization can result in a deleterious increase in paracellular permeability. Toxins such as E.coli cytotoxic necrotizing factor-1, H. Pylori associated factors, and Clostridium perfringens enterotoxin (CPE) have been found to induce claudin internalization (49, 50). The effect of CPE is due to a direct interaction with the second extracellular loop domain of claudin-3 and claudin-4, which can also inhibit tight junction reformation in addition to enhancing endocytosis (51).

Host factors and cytokines can also influence tight junction turnover and claudin expression, presumably in part to accommodate neurophil migration across epithelial barriers (52). For instance, interferon-γ increases claudin endocytosis and tight junction permeability (53). In addition to interferon-γ, other inflammatory cytokines such as tumor-necrosis factor (TNF)-α and interleukin (IL)-13 downregulate claudins and induce a marked increase in paracellular permeability by epithelial cells in culture (39, 54). The notion of pathogenic consequences for claudin misregulation are underscored by the findings that inflammatory bowel disease (IBD) causes a strong increase in the expression of claudin-2 while normal colons have low levels of claudin-2 (39, 55). A role for altered claudin expression in predisposition to IBD is also suggested by studies of JAM-A- deficient mice, which show increased claudin-10 and claudin-15 expression in the colonic mucosa and increased sensitivity to dextran sulfate sodium-induced colitis (56). Alterations in claudin expression has also correlated with pathological failures in barrier function in other systems. For instance, in chronic alcohol abuse, the lung barrier is compromised which is accompanied by decreased claudin-1 and claudin-7 and increased claudin-5 expression (38, 57). In these examples, the correlation of increased claudin content reducing the tight junction barrier is consistent with the notion of “leaky” claudins (3), although the mechanism underlying this remains obscure at present.

Physiologic roles for claudins

The function of specific claudins can be elucidated from mutations leading to human diseases, many which have been mimicked by mouse knockout models. For instance, mutations in claudin-1 cause neonatal icthyosis and sclerosing cholangitis, consistent with high levels of claudin-1 expression in the skin and in cholangiocytes of the bile duct (58) and claudin-1 null mice have a neonatal lethal phenotype due to skin permeability defects (58, 59). Claudin-14 mutations are associated with nonsyndromic deafness in humans and mice (60), which suggests a role for claudin-14 in the cation-restrictive barrier that maintains the normal endolymph ionic concentration that bathes the outer hair cells of the cochlea in the ear (61).

Mutations in claudin-16 (also known as paracellulin-1) lead to recessive renal hypomagnesaemia (62) and claudin-19 mutations show a similar deficiency in regulating magnesium reabsorption (63). Although it was initially hypothesized that claudin-16 directly regulated paracellular magnesium permeability, in fact claudin-16 interacts with claudin-19 to increase paracellular selectivity to sodium (64). This leads to a model where a gradient of claudin-16/claudin-19 expression in the thick ascending limb of the kidney establishes a NaCl concentration gradient which creates a lumen positive transepithelial diffusion potential that drives paracellular magnesium reabsorption, as opposed to a direct change in magnesium permeability, underscoring that roles for claudins in regulating ion permeability can be indirect (65).

Epithelial-mesenchymal transformation, cancer and claudin expression

Epithelial-mesenchymal transition (EMT) is a process where epithelial cells differentiate towards a more fibroblastic phenotype. EMT is involved in normal embryonic development and repair of epithelial injury, however it is also implicated in carcinogenesis (66, 67). Consistent with a decrease in epithelial gene expression, EMT is associated with changes in claudin expression and regulation. For instance, overexpression of the transcription factor SNAIL induces EMT and downregulates transcription of tight junction proteins including claudins and occludin (42, 43). This is due in part to E-box motifs in their promoter where SNAIL can bind and repress transcription (43, 68). SNAIL has also been shown to downregulate claudin-1 without affecting transcription (69), however, this may be a cell specific effect, since repression of claudin-1 transcription has been observed in other systems, suggesting that it might also regulate claudin translation (68).

Histological screens of human tumors have revealed changes in claudin expression associated with tumor phenotype. This is consistent with EMT as a contributing factor in tumorigenesis, since tight junctions are often altered in human carcinomas (70). Some trends are beginning to emerge for some claudins. For instance, tumors seem to be generally associated with decreased claudin-7 expression (71, 72). However, there are studies linking both decreased (73) or increased (74) claudin-1 expression by tumors. Claudin-3 and claudin-4 seem to be generally upregulated, particularly in aggressively metatastic tumors, although this is not always the case (72, 75). The physiologic role for upregulated claudin-3 and 4 in some human tumors is not fully understood, but this provides a fortuitous target, since it enables drugs based on Clostridium toxin which binds to the extracellular loop of these claudins to be developed as potential anti-tumor agents (7678). Since the structure of the loop binding domain of Clostridium toxin is now known, this could potentially enable other claudin specific interacting proteins to be designed (79).


Functional roles for claudins in regulating tissue barrier function have are becoming more solidly established, with claudin heterogeneity having both direct and indirect influences on paracellular ion permeability. There also remain several open questions in understanding how claudins are regulated. Some of the critical factors required for claudin assembly into tight junctions, such as roles for occludin and scaffold proteins are now known, but the mechanism of claudin assembly is not well understood. In particular, the intracellular compartments where claudins are assembled have not been identified and determinants which influence heteromeric and heterotypic claudin-claudin interactions have just begun to be elucidated. This last point is particularly critical to define how claudin multiplicity can influence paracellular permeability.

Finally, correlations between claudin expression and metastasis raise the intriguing notion of roles for claudins in cell growth control. Whether changes in claudin expression are a cause or a consequence of carcinogenesis is not clear. The observed discrepancies in claudin expression in tumorigenesis could be related to tissue-specific differences in claudin function or tissue microenvironmental features. Nonetheless, a role for claudins in regulation of cell growth would be comparable to roles found for other cell adhesion molecules in cancer, such as cadherins. Whether this is the case, remains to be determined, but this suggests that claudins play a multiple roles beyond acting as a simple paracellular barrier.


Supported by Emory Alcohol and Lung Biology Center (NIH P50-AA013757), NIH HL083120 (MK) and NIH T32GM008367 (MKF).

Contributor Information

Mary K. Findley, Division of Pulmonary, Allergy and Critical Care Medicine, Department of Medicine.

Michael Koval, Division of Pulmonary, Allergy and Critical Care Medicine, Department of Medicine and Department of Cell Biology, Emory University School of Medicine Atlanta, GA 30322.


1. Schneeberger EE, Lynch RD. The tight junction: a multifunctional complex. Am J Physiol Cell Physiol. 2004;286:C1213–1228. [PubMed]
2. Angelow S, Ahlstrom R, Yu AS. Biology of Claudins. Am J Physiol Renal Physiol. 2008;295:F867–F876. [PubMed]
3. Turksen K, Troy TC. Barriers built on claudins. J Cell Sci. 2004;117:2435–2447. [PubMed]
4. Koval M. Claudins--key pieces in the tight junction puzzle. Cell Commun Adhes. 2006;13:127–138. [PubMed]
5. 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 U S A. 1999;96:511–516. [PubMed]
6. Van Itallie CM, Rogan S, Yu A, Vidal LS, Holmes J, Anderson JM. Two splice variants of claudin-10 in the kidney create paracellular pores with different ion selectivities. Am J Physiol Renal Physiol. 2006;291:F1288–1299. [PubMed]
7. Niimi T, Nagashima K, Ward JM, Minoo P, Zimonjic DB, Popescu NC, Kimura S. claudin-18, a novel downstream target gene for the T/EBP/NKX2.1 homeodomain transcription factor, encodes lung- and stomach-specific isoforms through alternative splicing. Mol Cell Biol. 2001;21:7380–7390. [PMC free article] [PubMed]
8. Umeda K, Ikenouchi J, Katahira-Tayama S, Furuse K, Sasaki H, Nakayama M, Matsui T, Tsukita S, Furuse M. ZO-1 and ZO-2 independently determine where claudins are polymerized in tight-junction strand formation. Cell. 2006;126:741–754. [PubMed]
9. Van Itallie CM, Anderson JM. Claudins and epithelial paracellular transport. Annu Rev Physiol. 2006;68:403–429. [PubMed]
10. Piontek J, Winkler L, Wolburg H, Muller SL, Zuleger N, Piehl C, Wiesner B, Krause G, Blasig IE. Formation of tight junction: determinants of homophilic interaction between classic claudins. Faseb J. 2008;22:146–158. [PubMed]
11. Tsukita S, Furuse M. Claudin-based barrier in simple and stratified cellular sheets. Curr Opin Cell Biol. 2002;14:531–536. [PubMed]
12. Coyne CB, Gambling TM, Boucher RC, Carson JL, Johnson LG. Role of claudin interactions in airway tight junctional permeability. Am J Physiol Lung Cell Mol Physiol. 2003;285:L1166–1178. [PubMed]
13. Mitic LL, Unger VM, Anderson JM. Expression, solubilization, and biochemical characterization of the tight junction transmembrane protein claudin-4. Protein Sci. 2003;12:218–227. [PubMed]
14. Van Itallie CM, Gambling TM, Carson JL, Anderson JM. Palmitoylation of claudins is required for efficient tight-junction localization. J Cell Sci. 2005;118:1427–1436. [PubMed]
15. Nusrat A, Parkos CA, Verkade P, Foley CS, Liang TW, Innis-Whitehouse W, Eastburn KK, Madara JL. Tight junctions are membrane microdomains. J Cell Sci. 2000;113( Pt 10):1771–1781. [PubMed]
16. Hemler ME. Tetraspanin functions and associated microdomains. Nat Rev Mol Cell Biol. 2005;6:801–811. [PubMed]
17. Kovalenko OV, Yang XH, Hemler ME. A novel cysteine cross-linking method reveals a direct association between claudin-1 and tetraspanin CD9. Mol Cell Proteomics. 2007;6:1855–1867. [PubMed]
18. Evans MJ, von Hahn T, Tscherne DM, Syder AJ, Panis M, Wolk B, Hatziioannou T, McKeating JA, Bieniasz PD, Rice CM. Claudin-1 is a hepatitis C virus co-receptor required for a late step in entry. Nature. 2007;446:801–805. [PubMed]
19. Furuse M, Sasaki H, Tsukita S. Manner of interaction of heterogeneous claudin species within and between tight junction strands. J Cell Biol. 1999;147:891–903. [PMC free article] [PubMed]
20. Blasig IE, Winkler L, Lassowski B, Mueller SL, Zuleger N, Krause E, Krause G, Gast K, Kolbe M, Piontek J. On the self-association potential of transmembrane tight junction proteins. Cell Mol Life Sci. 2006;63:505–514. [PubMed]
21. Daugherty BL, Ward C, Smith T, Ritzenthaler JD, Koval M. Regulation of heterotypic claudin compatibility. J Biol Chem. 2007;282:30005–30013. [PubMed]
22. Tsukita S, Furuse M. Occludin and claudins in tight-junction strands: leading or supporting players? Trends Cell Biol. 1999;9:268–273. [PubMed]
23. Saitou M, Furuse M, Sasaki H, Schulzke JD, Fromm M, Takano H, Noda T, Tsukita S. Complex phenotype of mice lacking occludin, a component of tight junction strands. Mol Biol Cell. 2000;11:4131–4142. [PMC free article] [PubMed]
24. Furuse M, Fujita K, Hiiragi T, Fujimoto K, Tsukita S. Claudin-1 and -2: novel integral membrane proteins localizing at tight junctions with no sequence similarity to occludin. J Cell Biol. 1998;141:1539–1550. [PMC free article] [PubMed]
25. Nusrat A, Brown GT, Tom J, Drake A, Bui TT, Quan C, Mrsny RJ. Multiple protein interactions involving proposed extracellular loop domains of the tight junction protein occludin. Mol Biol Cell. 2005;16:1725–1734. [PMC free article] [PubMed]
26. Balda MS, Whitney JA, Flores C, Gonzalez S, Cereijido M, Matter K. 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. 1996;134:1031–1049. [PMC free article] [PubMed]
27. Mrsny RJ, Brown GT, Gerner-Smidt K, Buret AG, Meddings JB, Quan C, Koval M, Nusrat A. A key claudin extracellular loop domain is critical for epithelial barrier integrity. Am J Pathol. 2008;172:905–915. [PubMed]
28. Itoh M, Furuse M, Morita K, Kubota K, Saitou M, Tsukita S. Direct binding of three tight junction-associated MAGUKs, ZO-1, ZO-2, and ZO-3, with the COOH termini of claudins. J Cell Biol. 1999;147:1351–1363. [PMC free article] [PubMed]
29. Poliak S, Matlis S, Ullmer C, Scherer SS, Peles E. Distinct claudins and associated PDZ proteins form different autotypic tight junctions in myelinating Schwann cells. J Cell Biol. 2002;159:361–372. [PMC free article] [PubMed]
30. Roh MH, Makarova O, Liu CJ, Shin K, Lee S, Laurinec S, Goyal M, Wiggins R, Margolis B. The Maguk protein, Pals1, functions as an adapter, linking mammalian homologues of Crumbs and Discs Lost. J Cell Biol. 2002;157:161–172. [PMC free article] [PubMed]
31. Van Itallie CM, Colegio OR, Anderson JM. The cytoplasmic tails of claudins can influence tight junction barrier properties through effects on protein stability. J Membr Biol. 2004;199:29–38. [PubMed]
32. D’Souza T, Agarwal R, Morin PJ. Phosphorylation of claudin-3 at threonine 192 by cAMP-dependent protein kinase regulates tight junction barrier function in ovarian cancer cells. J Biol Chem. 2005;280:26233–26240. [PubMed]
33. Ikari A, Matsumoto S, Harada H, Takagi K, Hayashi H, Suzuki Y, Degawa M, Miwa M. Phosphorylation of paracellin-1 at Ser217 by protein kinase A is essential for localization in tight junctions. J Cell Sci. 2006;119:1781–1789. [PubMed]
34. Banan A, Zhang LJ, Shaikh M, Fields JZ, Choudhary S, Forsyth CB, Farhadi A, Keshavarzian A. theta Isoform of protein kinase C alters barrier function in intestinal epithelium through modulation of distinct claudin isotypes: a novel mechanism for regulation of permeability. J Pharmacol Exp Ther. 2005;313:962–982. [PubMed]
35. Nunbhakdi-Craig V, Machleidt T, Ogris E, Bellotto D, White CL, 3rd, Sontag E. Protein phosphatase 2A associates with and regulates atypical PKC and the epithelial tight junction complex. J Cell Biol. 2002;158:967–978. [PMC free article] [PubMed]
36. Turner JR, Angle JM, Black ED, Joyal JL, Sacks DB, Madara JL. PKC-dependent regulation of transepithelial resistance: roles of MLC and MLC kinase. Am J Physiol. 1999;277:C554–562. [PubMed]
37. Haorah J, Heilman D, Knipe B, Chrastil J, Leibhart J, Ghorpade A, Miller DW, Persidsky Y. Ethanol-induced activation of myosin light chain kinase leads to dysfunction of tight junctions and blood-brain barrier compromise. Alcohol Clin Exp Res. 2005;29:999–1009. [PubMed]
38. Fernandez AL, Koval M, Fan X, Guidot DM. Chronic alcohol ingestion alters claudin expression in the alveolar epithelium of rats. Alcohol. 2007;41:371–379. [PMC free article] [PubMed]
39. Prasad S, Mingrino R, Kaukinen K, Hayes KL, Powell RM, MacDonald TT, Collins JE. Inflammatory processes have differential effects on claudins 2, 3 and 4 in colonic epithelial cells. Lab Invest. 2005;85:1139–1162. [PubMed]
40. Tanaka M, Kamata R, Sakai R. EphA2 phosphorylates the cytoplasmic tail of Claudin-4 and mediates paracellular permeability. J Biol Chem. 2005;280:42375–42382. [PubMed]
41. Pasquale EB. Eph receptor signalling casts a wide net on cell behaviour. Nat Rev Mol Cell Biol. 2005;6:462–475. [PubMed]
42. Carrozzino F, Soulie P, Huber D, Mensi N, Orci L, Cano A, Feraille E, Montesano R. Inducible expression of Snail selectively increases paracellular ion permeability and differentially modulates tight junction proteins. Am J Physiol Cell Physiol. 2005;289:C1002–1014. [PubMed]
43. Ikenouchi J, Matsuda M, Furuse M, Tsukita S. Regulation of tight junctions during the epithelium-mesenchyme transition: direct repression of the gene expression of claudins/occludin by Snail. J Cell Sci. 2003;116:1959–1967. [PubMed]
44. Sasaki H, Matsui C, Furuse K, Mimori-Kiyosue Y, Furuse M, Tsukita S. Dynamic behavior of paired claudin strands within apposing plasma membranes. Proc Natl Acad Sci U S A. 2003;100:3971–3976. [PubMed]
45. Shen L, Weber CR, Turner JR. The tight junction protein complex undergoes rapid and continuous molecular remodeling at steady state. J Cell Biol. 2008;181:683–695. [PMC free article] [PubMed]
46. Ivanov AI, Nusrat A, Parkos CA. The epithelium in inflammatory bowel disease: potential role of endocytosis of junctional proteins in barrier disruption. Novartis Found Symp. 2004;263:115–124. discussion 124–132, 211-118. [PubMed]
47. Daugherty BL, Mateescu M, Patel AS, Wade K, Kimura S, Gonzales LW, Guttentag S, Ballard PL, Koval M. Developmental regulation of claudin localization by fetal alveolar epithelial cells. Am J Physiol Lung Cell Mol Physiol. 2004;287:L1266–1273. [PubMed]
48. Jordan K, Chodock R, Hand AR, Laird DW. The origin of annular junctions: a mechanism of gap junction internalization. J Cell Sci. 2001;114:763–773. [PubMed]
49. Fedwick JP, Lapointe TK, Meddings JB, Sherman PM, Buret AG. Helicobacter pylori activates myosin light-chain kinase to disrupt claudin-4 and claudin-5 and increase epithelial permeability. Infect Immun. 2005;73:7844–7852. [PMC free article] [PubMed]
50. Hopkins AM, Walsh SV, Verkade P, Boquet P, Nusrat A. Constitutive activation of Rho proteins by CNF-1 influences tight junction structure and epithelial barrier function. J Cell Sci. 2003;116:725–742. [PubMed]
51. Fujita K, Katahira J, Horiguchi Y, Sonoda N, Furuse M, Tsukita S. Clostridium perfringens enterotoxin binds to the second extracellular loop of claudin-3, a tight junction integral membrane protein. FEBS Lett. 2000;476:258–261. [PubMed]
52. Walsh SV, Hopkins AM, Nusrat A. Modulation of tight junction structure and function by cytokines. Adv Drug Deliv Rev. 2000;41:303–313. [PubMed]
53. Chiba H, Kojima T, Osanai M, Sawada N. The significance of interferon-gamma-triggered internalization of tight-junction proteins in inflammatory bowel disease. Sci STKE. 2006:pe1. [PubMed]
54. Tedelind S, Ericson LE, Karlsson JO, Nilsson M. Interferon-gamma down-regulates claudin-1 and impairs the epithelial barrier function in primary cultured human thyrocytes. Eur J Endocrinol. 2003;149:215–221. [PubMed]
55. Heller F, Florian P, Bojarski C, Richter J, Christ M, Hillenbrand B, Mankertz J, Gitter AH, Burgel N, Fromm M, et al. Interleukin-13 is the key effector Th2 cytokine in ulcerative colitis that affects epithelial tight junctions, apoptosis, and cell restitution. Gastroenterology. 2005;129:550–564. [PubMed]
56. Laukoetter MG, Nava P, Lee WY, Severson EA, Capaldo CT, Babbin BA, Williams IR, Koval M, Peatman E, Campbell JA, et al. JAM-A regulates permeability and inflammation in the intestine in vivo. J Exp Med. 2007;204:3067–3076. [PMC free article] [PubMed]
57. Wang F, Daugherty B, Keise LL, Wei Z, Foley JP, Savani RC, Koval M. Heterogeneity of claudin expression by alveolar epithelial cells. Am J Respir Cell Mol Biol. 2003;29:62–70. [PubMed]
58. Hadj-Rabia S, Baala L, Vabres P, Hamel-Teillac D, Jacquemin E, Fabre M, Lyonnet S, De Prost Y, Munnich A, Hadchouel M, et al. Claudin-1 gene mutations in neonatal sclerosing cholangitis associated with ichthyosis: a tight junction disease. Gastroenterology. 2004;127:1386–1390. [PubMed]
59. Furuse M, Hata M, Furuse K, Yoshida Y, Haratake A, Sugitani Y, Noda T, Kubo A, Tsukita S. Claudin-based tight junctions are crucial for the mammalian epidermal barrier: a lesson from claudin-1-deficient mice. J Cell Biol. 2002;156:1099–1111. [PMC free article] [PubMed]
60. Wattenhofer M, Reymond A, Falciola V, Charollais A, Caille D, Borel C, Lyle R, Estivill X, Petersen MB, Meda P, et al. Different mechanisms preclude mutant CLDN14 proteins from forming tight junctions in vitro. Hum Mutat. 2005;25:543–549. [PubMed]
61. Ben-Yosef T, Belyantseva IA, Saunders TL, Hughes ED, Kawamoto K, Van Itallie CM, Beyer LA, Halsey K, Gardner DJ, Wilcox ER, et al. Claudin 14 knockout mice, a model for autosomal recessive deafness DFNB29, are deaf due to cochlear hair cell degeneration. Hum Mol Genet. 2003;12:2049–2061. [PubMed]
62. Weber S, Schneider L, Peters M, Misselwitz J, Ronnefarth G, Boswald M, Bonzel KE, Seeman T, Sulakova T, Kuwertz-Broking E, et al. Novel paracellin-1 mutations in 25 families with familial hypomagnesemia with hypercalciuria and nephrocalcinosis. J Am Soc Nephrol. 2001;12:1872–1881. [PubMed]
63. Konrad M, Schaller A, Seelow D, Pandey AV, Waldegger S, Lesslauer A, Vitzthum H, Suzuki Y, Luk JM, Becker C, et al. Mutations in the tight-junction gene claudin 19 (CLDN19) are associated with renal magnesium wasting, renal failure, and severe ocular involvement. Am J Hum Genet. 2006;79:949–957. [PubMed]
64. Hou J, Renigunta A, Konrad M, Gomes AS, Schneeberger EE, Paul DL, Waldegger S, Goodenough DA. Claudin-16 and claudin-19 interact and form a cation-selective tight junction complex. J Clin Invest. 2008;118:619–628. [PubMed]
65. Angelow S, Yu AS. Claudins and paracellular transport: an update. Curr Opin Nephrol Hypertens. 2007;16:459–464. [PubMed]
66. Yang J, Weinberg RA. Epithelial-mesenchymal transition: at the crossroads of development and tumor metastasis. Dev Cell. 2008;14:818–829. [PubMed]
67. Radisky DC, Kenny PA, Bissell MJ. Fibrosis and cancer: do myofibroblasts come also from epithelial cells via EMT? J Cell Biochem. 2007;101:830–839. [PMC free article] [PubMed]
68. Martinez-Estrada OM, Culleres A, Soriano FX, Peinado H, Bolos V, Martinez FO, Reina M, Cano A, Fabre M, Vilaro S. The transcription factors Slug and Snail act as repressors of Claudin-1 expression in epithelial cells. Biochem J. 2006;394:449–457. [PubMed]
69. Ohkubo T, Ozawa M. The transcription factor Snail downregulates the tight junction components independently of E-cadherin downregulation. J Cell Sci. 2004;117:1675–1685. [PubMed]
70. Oliveira SS, Morgado-Diaz JA. Claudins: multifunctional players in epithelial tight junctions and their role in cancer. Cell Mol Life Sci. 2007;64:17–28. [PubMed]
71. Sauer T, Pedersen MK, Ebeltoft K, Naess O. Reduced expression of Claudin-7 in fine needle aspirates from breast carcinomas correlate with grading and metastatic disease. Cytopathology. 2005;16:193–198. [PubMed]
72. Montgomery E, Mamelak AJ, Gibson M, Maitra A, Sheikh S, Amr SS, Yang S, Brock M, Forastiere A, Zhang S, et al. Overexpression of claudin proteins in esophageal adenocarcinoma and its precursor lesions. Appl Immunohistochem Mol Morphol. 2006;14:24–30. [PubMed]
73. Tokes AM, Kulka J, Paku S, Szik A, Paska C, Novak PK, Szilak L, Kiss A, Bogi K, Schaff Z. Claudin-1, -3 and -4 proteins and mRNA expression in benign and malignant breast lesions: a research study. Breast Cancer Res. 2005;7:R296–305. [PMC free article] [PubMed]
74. Dhawan P, Singh AB, Deane NG, No Y, Shiou SR, Schmidt C, Neff J, Washington MK, Beauchamp RD. Claudin-1 regulates cellular transformation and metastatic behavior in colon cancer. J Clin Invest. 2005;115:1765–1776. [PubMed]
75. Cunningham SC, Kamangar F, Kim MP, Hammoud S, Haque R, Iacobuzio-Donahue CA, Maitra A, Ashfaq R, Hustinx S, Heitmiller RE, et al. Claudin-4, mitogen-activated protein kinase kinase 4, and stratifin are markers of gastric adenocarcinoma precursor lesions. Cancer Epidemiol Biomarkers Prev. 2006;15:281–287. [PubMed]
76. Kominsky SL. Claudins: emerging targets for cancer therapy. Expert Rev Mol Med. 2006;8:1–11. [PubMed]
77. Kominsky SL, Vali M, Korz D, Gabig TG, Weitzman SA, Argani P, Sukumar S. Clostridium perfringens enterotoxin elicits rapid and specific cytolysis of breast carcinoma cells mediated through tight junction proteins claudin 3 and 4. Am J Pathol. 2004;164:1627–1633. [PubMed]
78. Michl P, Buchholz M, Rolke M, Kunsch S, Lohr M, McClane B, Tsukita S, Leder G, Adler G, Gress TM. Claudin-4: a new target for pancreatic cancer treatment using Clostridium perfringens enterotoxin. Gastroenterology. 2001;121:678–684. [PubMed]
79. Van Itallie CM, Betts L, Smedley JG, 3rd, McClane BA, Anderson JM. Structure of the claudin-binding domain of Clostridium perfringens enterotoxin. J Biol Chem. 2008;283:268–274. [PubMed]