Tight Junctions: Molecular Architecture and Function

Tight Junctions: Molecular Architecture and Function Saima Aijaz, Maria S. Balda, and Karl Matter Division of Cell Biology, Institute of Ophthalmology, University College London, London, United Kingdom

Tight junctions are the most apical component of the epithelial junctional complex and are crucial for the formation and functioning of epithelial and endothelial barriers. They regulate selective diffusion of ions and solutes along the paracellular pathway and restrict apical/basolateral intramembrane diffusion of lipids. Research over the past years provided much insight into the molecular composition of tight junctions, and we are starting to understand the mechanisms that permit selective paracellular diffusion. Moreover, a complex network of proteins has been identified at tight junctions that is based on cytoskeleton‐linked adaptors that recruit and thereby often regulate different types of signaling components that regulate epithelial proliferation, differentiation, and polarization. KEYWORDS: Intercellular junctions, Tight junctions, Adhesion, Differentiation, Cell polarity, Signal transduction, Oncogenesis, Permeability. ß 2006 Elsevier Inc.

I. Introduction Epithelia and endothelia form selective barriers between tissues and diVerent body compartments. This requires that they polarize (i.e., they develop an apical and a basolateral cell surface domain) and that they adhere to each other through adhesive complexes between the cells. These intercellular junctional complexes carry out adhesive functions but also contain crucial components of signaling pathways that regulate cell proliferation, polarization, and diVerentiation. Vertebrate epithelial cells are joined to each other via a set of intercellular junctions that consists of gap junctions, desmosomes, adherens junctions, and tight junctions or the zonula occludens. The latter three junctions are International Review of Cytology, Vol. 248 Copyright 2006, Elsevier Inc. All rights reserved.

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0074-7696/06 $35.00 DOI: 10.1016/S0074-7696(06)48005-0

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often referred to as the ‘‘epithelial junctional complex’’ (Cereijido et al., 2004; Farquhar and Palade, 1963). Tight junctions encircle cells at the apical end of the lateral membrane and are important for the barrier properties of epithelia and endothelia (Kachar and Reese, 1982; Staehelin, 1973). Adherens junctions and desmosomes are adhesive junctions and are linked to the actin cytoskeleton and intermediate filaments, respectively. Depending on the epithelial cell type, adherens junction components can be concentrated close to tight junctions and form a morphologically clearly distinct junction that colocalizes with a prominent actin belt, or they can be distributed over the entire lateral membrane. Desmosomes are generally found all along the lateral membrane. Gap junctions form intercellular pores that allow the exchange of small hydrophilic molecules between cells. In endothelia, adherens and tight junctions are not as morphologically distinct as they are in epithelia, and classical desmosomes are absent (Bazzoni and Dejana, 2004; Vestweber, 2000; Wolburg and Lippoldt, 2002). Although the basic architecture of epithelial and endothelial tight and adherens junctions is the same, certain junctional components are preferentially expressed by either endothelial or epithelial cells. Tight junctions have a striking ultrastructural appearance. In ultrathin sections, they appear as very close contacts between the plasma membranes of two neighboring cells (Fig. 1). These close contacts sometimes seem to contain focal hemifusions of the two membranes, which has led to the speculation that the junctional barrier might be of a lipidic nature (Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982). In freeze‐fracture replicas, a technique that allows the visualization of the ultrastructure of the hydrophobic membrane surfaces, the contact sites appear as a net‐like meshwork of intramembrane fibrils. In the lipid model, it was suggested that these intramembrane fibrils represented inverted cylindrical micelles. Today, it is generally assumed that they represent polymers of interacting junctional transmembrane components, although a contribution from lipids and specialized lipid structures cannot be ruled out (Tsukita et al., 2001). Tight junctions possess multiple functions. They are essential for the barrier function of epithelia and endothelia by restricting paracellular diVusion. The junctional diVusion barrier is not an absolute one, however, but is semipermeable as it allows the selective passage of certain solutes but not others (Anderson et al., 2004; Cereijido et al., 2004). Furthermore, paracellular permeability diVers from epithelia to epithelia and can change due to physiological and pathological stimuli (Balda et al., 1992; Bentzel et al., 1991; Claude, 1978; Madara, 1988). Tight junctions also contribute to the maintenance of cell surface polarity by forming an intramembrane diVusion fence that restricts diVusion of lipids in the exoplasmic leaflet of the plasma membrane (Dragsten et al., 1981; van Meer and Simons, 1986). More recently, tight junctions have been shown to harbor evolutionarily conserved protein

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FIG. 1 Epithelial and endothelial intercellular junctions. Electron micrographs of human retinal pigment epithelial cells (A) and brain endothelial cells (B) were taken from ultrathin sections derived from Araldite‐embedded retinas and primary cultures, respectively. TJ, tight junction; AJ, adherens junction; POS, photoreceptor outer segment; M, melanosome. (Courtesy of Peter Munro, Institute of Ophthalmology, University College London, London.)

complexes that regulate polarization and junction assembly and to recruit signaling molecules that participate in the regulation of cell proliferation, diVerentiation, and gene expression (Matter and Balda, 2003b). II. Molecular Composition Tight junctions possess the characteristic molecular architecture of an adhesion complex (Table I). They consist of a set of diVerent types of transmembrane proteins that functions in barrier formation and regulation,

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TABLE I Components of Tight Junctions and Their Interactions Protein Occludin

Claudins

Protein type and functions

Interacting proteins

Transmembrane protein Cell–cell adhesion

JAM‐A ZO‐1, ‐2, ‐3, cingulin

Regulation of paracellular permeability

c‐yes, PI3 kinase TGF‐b‐receptor type I

Modulation of signaling pathways: Raf‐1, RhoA, TGF‐b

E3 ubiquitin ligase—Itch f‐actin

Transmembrane proteins Cell–cell adhesion

ZO‐1, ‐2, ‐3 MUPP‐1, PATJ

Regulation of paracellular permeability

WNK4 kinase OAP‐1

Mediation of ion‐selective diVusion

C. perfringens enterotoxin

Regulation of cell migration JAMs, CAR, ESAM, CLMP

Transmembrane proteins Cell–cell adhesion Regulation of leukocyte transmigration Regulation of junction assembly

Occludin ZO‐1, cingulin, Par3, MUPP‐1, MAGI‐1, AF‐6 CASK LFA‐1 PICK‐1 H. pylori CagA Reovirus, adenovirus, coxsackievirus

CRB3

Transmembrane protein (localizes along entire apical membrane) Regulation of polarization and junction assembly

Pals1, PATJ Par6

ZO‐1

Cytoplasmic adaptor protein Regulation of junction assembly

Occludin, claudins, JAMs ZO‐2, ‐3, cingulin, f‐actin, AF‐6, a‐catenin, ARVCF Connexins‐30/36/43/45/ 47/50 ZONAB

Regulation of gene expression Regulation of cell proliferation

Ga12, ZAK a‐Dystrobrevin (continued )

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TIGHT JUNCTIONS TABLE I (continued) Protein ZO‐2

Protein type and functions

Interacting proteins

Cytoplasmic adaptor protein Regulation of gene expression

Occludin, claudins, ZO‐1, f‐actin, cingulin

Binds and inhibits viral oncogenes

a‐Catenin, 4.1R SAF‐B, C/EBP, Jun, Fos ARVCF Papillomavirus E6 Adenovirus type 9 E4‐ORF1

ZO‐3

Cytoplasmic adaptor protein

Occludin, claudins ZO‐1, cingulin, f‐actin, PATJ

MAGI‐1

Cytoplasmic adaptor protein Binds and inhibits viral oncogenes

JAMs Synaptopodin a‐Actinin 4

AF‐6, p120 catenin

Rap GEP Adenovirus E4‐ORF1 Papillomavirus E6 MAGI‐2

Cytoplasmic adaptor protein

PTEN b1‐Adrenergic receptor

MAGI‐3

Cytoplasmic adaptor protein

PTEN RPTPb pro‐TGF‐a Leukemia virus type 1 Tax Oncoprotein

MUPP‐1

Cytoplasmic adaptor protein

Claudins, JAMs, CAR c‐Kit Papillomavirus E6 Adenovirus type 9 E4‐ORF1

Pals1

Cytoplasmic adaptor protein Regulation of junction assembly and polarization

CRB3 PATJ, Par6

PATJ

Cytoplasmic adaptor protein Regulation of junction assembly and polarization

Pals1 ZO‐3

Cingulin

Cytoplasmic adaptor protein Regulation of RhoA signaling and gene expression

JAM ZO‐1, ‐2, ‐3 f‐Actin, myosin

JACOP

Cytoplasmic adaptor protein (also localizes to adherens junctions)

GEF‐H1/Lfc f‐Actin

(continued)

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TABLE I (continued) Protein Par3/ASIP

Protein type and functions Cytoplasmic adaptor protein Regulation of junction assembly and polarization

Interacting proteins JAMs Par6, aPKC 14–3‐3, TIAM1, PTEN Phospholipase C b, nectin

Par6

Cytoplasmic adaptor protein Regulation of junction assembly and polarization

CRB3 Par3, aPKC, Cdc42 Pals1, mLgl TGF‐b receptors E3 ubiquitin ligase Smurf1 Phospholipase C b ECT2

aPKC

Cytoplasmic protein kinase Regulation of junction assembly and polarization

Par3, Par6 Protein phosphatase 2A

PTEN

Cytoplasmic lipid phosphatase

MAGI‐1, ‐2, ‐3 Par3

GEF‐H1/Lfc

Cytoplasmic guanine nucleotide exchange factor for Rho

Rho Cingulin

Regulation of Rho signaling, paracellular permeability, and G1/S phase progression

PAK1, PAK4 14–3‐3

G proteins

Cytoplasmic trimeric GTP binding proteins Regulation of junction assembly

ZO‐1 (Ga12)

Rab13

Cytoplasmic monomeric GTP binding protein Regulation of junction assembly and PKA signaling

PKA

Rab3b

Cytoplasmic monomeric GTP binding protein Cytoplasmic protein phosphatase Regulation of junction assembly

Protein phosphatase 2A

aPKC

WNK4

Cytoplasmic protein kinase Regulation of paracellular ion permeability

Claudins

c‐Yes

Cytoplasmic tyrosine kinase Regulation of junction assembly

Occludin

Cdk4

NACo: localizes to tight junctions and nucleus; cell division kinase Regulates G1/S phase progression

ZONAB D‐type cyclins

ZONAB

NACo: localizes to tight junctions and nucleus; transcription factor

ZO‐1 Cdk4

Regulates gene expression

RalA (continued)

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TIGHT JUNCTIONS TABLE I (continued) Protein Symplekin

Protein type and functions

Interacting proteins

NACo: localizes to tight junctions and nucleus; interacts with polyadenylation machinery Regulation of polyadenylation

CstF CPSF

AP‐1

NACo: localizes to tight junctions and nucleus; transcription factor Regulation of gene expression

ZO‐2

HuASH1 Sec6/8 complex

NACo: localizes to tight junctions and nucleus; transcription factor Cytoplasmic protein complex Regulation of vesicular traYc to the basolateral membrane

7H6/barmotin

Cytoplasmic protein

JEAP

Cytoplasmic protein of exocrine cells Cytoplasmic protein

Pilt LYRIC

HSF1

Cytoplasmic protein transiently associated with tight junctions

adhesion, and signal transduction. These transmembrane components are linked to a cytoplasmic plaque that consists of a scaVold of adaptor proteins that anchors the junction to the actin cytoskeleton. This protein network recruits multiple signaling proteins that regulate tight junction function and cell proliferation and diVerentiation. An important class of recruited signaling proteins consists of dual localization proteins that appear to shuttle between the junction and the nucleus, where they function in the regulation of gene expression, and have hence been named NACos for proteins associated with the Nucleus and Adhesion Complexes (Balda and Matter, 2003). The localization of proteins to tight junctions is not always straightforward. Because tight and adherens junctions are very close to each other, such a localization is optimally based on immunoelectron microscopy. However, this technique often fails because many antibodies do not work for this application and/or the investigated protein is expressed at low levels, as is the case for many signaling components. Therefore, colocalization with tight junction markers by confocal microscopy in cells with a clearly distinct distribution of adherens junctions, such as in the mouse trophectoderm or Madin–Darby canine kidney (MDCK) cells, combined with biochemical interactions with known tight junction components, is generally taken as suYcient evidence for association with tight junctions.

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Occludin is the first transmembrane protein of tight junctions that had been identified and has been linked to various junctional functions (Feldman et al., 2005; Furuse et al., 1993; Matter and Balda, 1999). Occludin is a polytopic membrane protein with four transmembrane domains, N‐ and C‐termini in the cytosol and, consequently, two extracellular loops. This originally predicted membrane topology is supported by various experimental evidence that includes binding sites of interaction partners such as ZO‐1 or extracellular binding partners, N‐glycosylation of sites introduced into the extracellular loops, and accessibility of epitopes in nonpermeabilized cells (Furuse et al., 1994; Matter and Balda, 1998; Nusrat et al., 2005; Van Itallie and Anderson, 1997). Although occludin is a component of the intramembrane strands observed in freeze‐fracture replicas, increased or reduced expression of occludin or expression of various occludin mutants does not aVect tight junction morphology, suggesting that it is not a critical structural component of tight junctions (Balda et al., 1996b, 2000; Fujimoto, 1995; McCarthy et al., 1996; Saitou et al., 1998, 2000; Yu et al., 2005). Occludin is expressed by all vertebrates tested, and expression is driven by a single occludin gene. Nevertheless, five diVerent isoforms have been described, which arise by alternative splicing. Occludin1B contains an insertion that results in an additional 56 amino acids at the N‐terminus. It also is associated with tight junctions and has a broad tissue expression (Muresan et al., 2000). Although is not clear whether this insertion aVects occludin function, it is possible that the modified N‐terminus diVers in cytoplasmic interactions and aVects the transmigration of leukocytes since the N‐terminal cytoplasmic domain of occludin is important for the regulation of transepithelial neutrophil migration (Huber et al., 2000). The N‐terminal domain also interacts with the E3 ubiquitin ligase itch, which regulates occludin turnover and may regulate tight junction assembly in Sertoli cells (Lui and Lee, 2005; Traweger et al., 2002a). However, it is not known whether itch regulates neutrophil transmigration. One type of alternative splicing event eliminates the fourth transmembrane domain, resulting in an isoform that exposes its C‐terminal domain to the exterior and not the cytosol (Ghassemifar et al., 2002; Mankertz et al., 2002). As expression of this isoform was observed at the periphery of subconfluent islands of cells, it was suggested that its expression might be related to epithelial wound repair (Ghassemifar et al., 2002). Mice deficient in occludin expression do not exhibit obvious junctional defects but are unable to reproduce and have a complex phenotype, which includes defects in epithelial diVerentiation in various tissues (Saitou et al., 2000; Schulzke et al., 2005). Although the molecular mechanisms by which occludin deficiency causes these defects in diVerentiation are unknown, occludin has been linked to several diVerent signaling pathways and junctional regulation (see below).

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The second type of tight junction‐associated transmembrane protein consists of the claudins. The founding members of the claudin family, claudin‐1 and ‐2, were discovered since they copurified with occludin in junctional fractions from chick hepatocytes (Furuse et al., 1998a). Today more than 20 human claudin genes are known. Although claudins appear to be central components of vertebrate tight junctions, they are also expressed in other metazoa such as Drosophila, where they localize to septate junctions (Behr et al., 2003; Wu et al., 2004). Claudins have four transmembrane domains and expose both termini to the cytosol as does occludin. Expression of a single claudin type is suYcient to induce the appearance of tight junction‐like intramembrane strands in fibroblasts, suggesting that they are important structural components of tight junctions (Furuse et al., 1998b). This is supported by the disappearance of junctional intramembrane strands in central nervous system (CNS) myelin and Sertoli cells in claudin‐11 null mice (Gow et al., 1999). The extracellular loops show the most remarkable diVerences between various claudin family members. This has led to the speculation that claudins are not only important for barrier formation but that they are responsible for the selective permeability of the paracellular pathway (Tsukita et al., 2001). Indeed, multiple evidence now suggests that claudins are important for junctional ion selectivity and that the expressed claudin repertoire of an epithelium determines the ion selectivity of the paracellular pathway (Furuse et al., 2001; Tsukita et al., 2001; Van Itallie and Anderson, 2004; Van Itallie et al., 2001). Tight junctions also contain diVerent types of single‐span transmembrane proteins. Several members of the CTX/ VH‐C2 family of adhesion proteins also associate with tight junctions in epithelia and endothelia (Table I). These proteins contain two immunoglobulin folds in the extracellular domain, one of the VH and one of the C2 type (Bazzoni, 2003). Examples are the junctional adhesion molecules (JAMs). Four JAMs––JAM‐A, ‐B, ‐C, and ‐4 (D)––have been identified and all four are able to mediate cell–cell adhesion. Several of the JAM proteins as well as the related proteins CLMP and the endothelial‐ specific adhesion protein ESAM associate with tight junctions and interact with adaptor proteins of the cytoplasmic plaque (Bazzoni et al., 2000; Ebnet et al., 2000, 2003; Raschperger et al., 2004; Wegmann et al., 2004). Coxsackie virus and adenovirus receptor (CAR) is another member of the CTX family that associates with epithelial junctions (Cohen et al., 2001; Walters et al., 2002). It forms homotypic adhesions and was reported to be associated with the junctional intramembrane strands (Coyne and Bergelson, 2005). Its C‐terminal cytoplasmic domain interacts with the junctional plaque adaptor ZO‐1 as well as a range of other cytoplasmic proteins (Coyne and Bergelson, 2005; Cohen et al., 2001). These interactions are likely to be responsible for the observed disruption of tight junctions upon binding

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of adenovirus fibers to CAR, which results in apical escape of the virus after basolateral secretion (Walters et al., 2002). JAM‐A also functions as a receptor for pathogens by interacting with reovirus (Barton et al., 2001). Moreover, the Helicobactor pylori CagA protein associates with a ZO‐1/ JAM‐A complex, which is thought to induce corruption of the gastric epithelial barrier (Amieva et al., 2003). A second type of single‐span protein associated with tight junctions represents CRB‐3, a 25‐kDa type I membrane protein. CRB‐3 is the only one of the three vertebrate homologues of the Drosophila melanogastor crumbs protein that has been demonstrated to associate with tight junctions in epithelial cells (Lemmers et al., 2004; Makarova et al., 2003; Roh et al., 2003). CRB‐3 is spread over the entire apical membrane and only a minor fraction of the entire cellular pool is associated with tight junctions. The cytoplasmic domain of CRB‐3 interacts with cytosolic adaptor proteins, such as Pals1, linking it to the cellular machinery that regulates epithelial polarization. The scaVold of the cytoplasmic plaque at tight junctions is formed by adaptor proteins, such as ZO‐1/2/3, PATJ, Pals1, PAR‐3, and PAR‐6 (Table I). A large number of protein–protein interactions have been characterized between these proteins, suggesting that they form an intricate protein network. Many of these proteins interact with transmembrane components as well as f‐actin, resulting in a molecular bridge between the adhesion proteins and the cytoskeleton (Fanning, 2001; Schneeberger and Lynch, 2004). Several of these proteins have only been identified recently and little is known about their function (Table I) (Britt et al., 2004; Kawabe et al., 2001; Muto et al., 2000; Nishimura et al., 2002). ZO‐1, the first identified tight junction component, illustrates the architectural principle of the junctional plaque nicely (Stevenson et al., 1986). ZO‐1 has several protein–protein interaction domains: three PDZ (for PSD‐95, DlgA, and ZO‐1 homology) domains, one SH3 domain, and one with homology to yeast guanylate kinase (GUK domain). Proteins containing these types of domains are called MAGUKs (membrane‐associated guanylase kinase proteins); several members of this protein family associate with tight junctions (Table I) (Tsukita et al., 2001; Willott et al., 1993; Woods and Bryant, 1993). ZO‐1 can interact with the transmembrane proteins JAMs, claudins, and occludin, form stable complexes with either ZO‐2 or ZO‐3 via a PDZ–PDZ domain mediated interaction, bind to other adaptors such as cingulin, and contain a discrete actin‐binding domain in its C‐terminal half (Bazzoni et al., 2000; Cordenonsi et al., 1999; Ebnet et al., 2000; Fanning and Anderson, 1998; Furuse et al., 1994; Itoh et al., 1999). Moreover, its SH3 domain functions in intracellular signaling mechanisms by binding to the serine protein kinase ZAK and the Y‐box transcription factor ZONAB, which in turn binds to the cell cycle regulator CDK4 (Balda and Matter,

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2000a; Balda et al., 1996a, 2003). During junction assembly or in cell types lacking tight junctions, ZO‐1 has also been reported to interact with the adherens junction components AF‐6, a‐catenin, and ARVCF (Itoh et al., 1997; Kausalya et al., 2004; Rajasekaran et al., 1996; Yamamoto et al., 1997). ZO‐1 can also interact with multiple connexins and has been localized to gap junctions in various cell types such as cardiac myocytes, oligodendrocytes, and astrocytes; this interaction is also thought to be important for the close association of certain connexins with tight junctions (Giepmans and Moolenaar, 1998; Kausalya et al., 2001; Kojima et al., 2001; Laing et al., 2001; Li et al., 2004a,b; Nielsen et al., 2003; Penes et al., 2005; Toyofuku et al., 1998). Moreover, diVerent isoforms of ZO‐1 containing diVerent alternatively spliced domains have been described; expression of one of which, the a‐ domain, has been shown to correlate with the plasticity of tight junctions and is developmentally regulated (Balda and Anderson, 1993; Sheth et al., 1997). It is clear that some of the interactions of ZO‐1 occur only during junction assembly or in specific cell types. Moreover, ZO‐1, as other tight junction‐associated proteins, is likely to be part of diVerent types of complexes in fully assembled tight junctions. A diVerent type of junctional adaptor protein is cingulin, a dimeric actin‐ binding protein that does not have classical protein–protein interaction motifs (Citi et al., 1988; Cordenonsi et al., 1999; D’Atri and Citi, 2001). Nevertheless, it interacts with many diVerent junctional proteins (see Table I). As cingulin can form short filaments in vitro, it might form a supporting junctional scaVold that intertwines diVerent junctional subcomplexes (D’Atri and Citi, 2002). However, a partial knockout of the cingulin gene, which resulted in the expression of an N‐terminally truncated protein, did not aVect tight junction morphology, suggesting that it is either not critically contributing to tight junction structure or that its architectural function is redundant (Guillemot et al., 2004). A candidate for a protein with a similar function is JACOB, a protein homologous to cingulin that associates with diVerent types of intercellular junctions including tight junctions (Ohnishi et al., 2004). The junctional plaque also contains diVerent types of signaling proteins, which can be divided into two diVerent classes: classical signaling proteins such as G proteins and kinases, and dual localization proteins that not only localize to tight junctions but that also have functions at other cellular sites such as the nucleus (Table I). The class of classical signaling proteins includes a variety of diVerent proteins ranging from protein kinases and protein and lipid phosphatases to heterotrimeric and small GTP‐binding proteins (Gonzalez‐Mariscal et al., 2003; Kohler and Zahraoui, 2005; Schneeberger and Lynch, 2004). In many cases, it has been concluded that these proteins associate with tight junctions because of immunofluorescence data and/or protein–protein interactions.

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Immunoelectron microscopy has often proven diYcult as these proteins are frequently not expressed at high levels. Functionally, they have been associated with roles in the regulation of junction assembly and disassembly as well as of junctional functions such as selective paracellular permeability. Moreover, some of these tight junction‐associated signaling proteins participate in the regulation of cell proliferation and diVerentiation. The second class of proteins is the dual localization proteins. As they generally localize to either sites of adhesion or the nucleus, they have also been named NACos (Balda and Matter, 2003). The majority of these proteins are transcription factors such as the Y‐box factor ZONAB, huASH1, c‐fos, and c‐jun, or proteins that interact with them such as the cell division kinase CDK4 (Balda and Matter, 2000a; Balda et al., 2003; Betanzos et al., 2004; Nakamura et al., 2000). Generally, these proteins have been linked to the regulation of gene expression and/or epithelial proliferation and diVerentiation. Another intriguing example is symplekin, which also associates with the polyadenylation machinery (Barnard et al., 2004; Hofmann et al., 2002; Keon et al., 1996; Takagaki and Manley, 2000). Additionally, some of the adaptor proteins (e.g., ZO‐1 and ZO‐2) have also been reported to localize to the nucleus under certain conditions. However, the reported results are controversial and potential nuclear functions have not been defined (Balda and Matter, 2000a; Gottardi et al., 1996; Jaramillo et al., 2004; Reichert et al., 2000; Traweger et al., 2002b).

III. Junctional Diffusion Barriers A. Selective Paracellular Permeability Epithelial tight junctions form a semipermeable paracellular diVusion barrier or gate that is crucial for epithelia and endothelia to separate diVerent body compartments. Functionally, this diVusion gate has been known for over 40 years to restrict diVusion on the basis of charge and size of the solute or experimental tracer (Cereijido et al., 1978; Lindemann and Solomon, 1962; Moreno and Diamond, 1975; Tang and Goodenough, 2003; Wright and Diamond, 1968). The ion and size selectivity of the paracellular pathway diVers among epithelia (Anderson et al., 2004; Powell, 1981) and is regulated by diVerent physiological and pathological stimuli (Benais‐Pont et al., 2001; Nguyen et al., 2001; Schulzke and Fromm, 2001; Turner and Madara, 2001). A model for junctional permeability had been proposed many years ago that was based on a series of diVusion barriers containing fluctuating pores or channels and that has subsequently been modified to include compartmentalization and regulation (Balda and Matter, 2000b; Cereijido et al., 1989;

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Claude, 1978; Madara, 1998). Although the lack of insight into the biochemical composition of tight junctions has hindered progress toward an understanding of tight junction function for a long time, multiple experimental evidence now supports the existence of paracellular channels. Junctional ion permeability seems to be primarily determined by claudins. Claudins are expressed in a tissue‐specific manner and the claudin composition of a tight junction in cultured epithelial cell lines determines paracellular ion selectivity (Anderson et al., 2004; Tsukita and Furuse, 2002). The first experimental evidence for the importance of claudins for the ion specificity of a junction came from claudin‐16/paracellin‐1, which is expressed in the lung and the kidney; patients carrying a mutation in this gene suVer from hereditary hypomagnesemia (renal magnesium wasting) due to a deficiency in paracellular magnesium resorption in the kidney (Simon et al., 1999). Claudin‐16 has hence been proposed to form a paracellular channel or pore that allows the diVusion of magnesium ions. Transfection experiments with epithelial cell lines further suggested that diVerent claudins favor paracellular diVusion of specific ions (Furuse et al., 2001; Jeansonne et al., 2003; Van Itallie et al., 2001, 2003; Yu et al., 2003). These and other observations have led to a model in which claudins form homo‐ and heterooligomers that then engage in intercellular interactions to form paracellular channels or pores. The ion selectivity of these channels is determined by the types of claudins they contain (Tsukita et al., 2001; Van Itallie and Anderson, 2004). Because mutations in the extracellular loops of claudin‐4 and ‐15 reverse their respective charge selectivity, it is thought that the extracellular domains of claudins directly participate in the formation of the paracellular channels (Colegio et al., 2002). Recent reports also identified protein phosphorylation as a molecular mechanism that regulates the activity of claudin‐based channels. A threonine residue in the C‐terminal domain of several claudins has been shown to become phosphorylated in response to stimuli such as protein kinase A (PKA)‐activating compounds (D’Souza et al., 2005; Fujibe et al., 2004; Soma et al., 2004). In the majority of the studies, phosphorylation seems to interfere with junctional integration and/or favors increased ionic permeability. A pathologically important example of claudin phosphorylation involves the tight junction‐associated WNK4 kinase. Mutations in WNK4 as well as the related WNK1 kinase cause hypertension due to eVects on renal salt reabsorption and potassium ion excretion (pseudohypoaldosteronism type II) (Wilson et al., 2001). Disease‐causing WNK4 alleles carry gain of function mutations and stimulate claudin phosphorylation, resulting in increased paracellular chloride permeability (Kahle et al., 2004; Yamauchi et al., 2004). Apart from the above‐mentioned disorders, other hereditary diseases and transgenic mice models also indicate that specific claudins are important for the normal functioning of various tissues. In most cases, however, it has not been determined whether the phenotype is caused by alterations in junctional

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permeability. For example, mutations in claudin‐14 cause a form of human hereditary deafness, and claudin‐14‐deficient mice show outer hair cell degeneration in the organ of Corti (Wilcox et al., 2001). It is thus possible that hair cell degeneration in the absence of claudin‐14 is due to a switch in junctional ion selectivity. Tight junctions not only permit selective diVusion of ions along concentration gradients but also of small hydrophilic tracers (Balda and Matter, 1998). Although it was originally thought that the same channels that permit ion diVusion also mediate tracer diVusion, recent evidence suggests that permeability of ions and tracers is mediated by a diVerent mechanism. DiVusion of hydrophilic tracers is a slow process and is usually measured over a couple of hours (Matter and Balda, 2003a). Although there is some variation from one tissue to another in the size selectivity of tracer diVusion, tracers that are larger than a few kilodaltons are usually not able to diVuse across intact junctions at detectable levels. Although the actual mechanism responsible for size‐selective tracer diVusion is not known, several regulators have been identified. The transmembrane protein occludin has been shown to be important for the regulation of tracer diVusion (Matter and Balda, 1999). Overexpression of occludin was shown to increase size‐selective paracellular permeability and decrease ion conductance, suggesting not only that occludin regulates tracer diVusion but also that tracer and ion diVusion are likely to be mediated by diVerent mechanisms (Balda et al., 1996b; McCarthy et al., 1996). The activity of occludin is regulated by its C‐terminal cytoplasmic domain since expression of occludin mutants lacking this part in cultured cell lines results in stimulation of size‐selective tracer permeability in cultured epithelial cells and loss of the epithelial barrier function in Xenopus embryos (Balda et al., 1996b; Chen et al., 1997; Huber et al., 2000). It is thought that this regulatory mechanism involves phosphorylation events as well as the actin cytoskeleton since the C‐terminal domain of occludin binds to protein and lipid kinases, as well as to actin filaments and cytoskeletal linkers (Andreeva et al., 2001; Antonetti et al., 1999; Clarke et al., 2000b; Cordenonsi et al., 1997; Farshori and Kachar, 1999; Hirase et al., 2001; Muller et al., 2005; Nusrat et al., 2000a; Sakakibara et al., 1997; Schmidt et al., 2004; Tsukamoto and Nigam, 1999; Wong, 1997). Moreover, the extracellular domains of occludin have also been shown to be important for its function, as expression of proteins carrying deletions in the extracellular loops resulted in an inhibition of tracer diVusion (Balda et al., 2000). Moreover, peptides corresponding to extracellular domain sequences can cause junctional disruption if added to cultured cells (Vietor et al., 2001; Wong and Gumbiner, 1997). As similar peptides can bind to other junctional membrane components such as claudin‐1 and JAM‐A, junctional disruption might be caused by dissociating interactions (Nusrat et al., 2005).

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Regulation of the small GTPase RhoA has also been shown to modulate size‐selective paracellular diVusion. Transfection of dominant‐negative and constitutively active RhoA interferes with normal junction assembly, and inhibition of endogenous RhoA by C3 transferase disrupts tight junctions (Jou et al., 1998; Nusrat et al., 1995). However, more subtle regulations of RhoA activity by regulating the expression of a tight junction‐associated RhoA‐specific guanine nucleotide exchange factor (GEF‐H1/Lfc) or by modulating RhoA activity by activation of a prostaglandin receptor resulted in specific regulation of small tracer diVusion (Benais‐Pont et al., 2003; Hasegawa et al., 1999). How GEF‐H1/Lfc is normally activated is not known, but it is possible that occludin might play a role in this as MDCK cells with reduced occludin expression are deficient in RhoA activation in response to certain stimuli (Yu et al., 2005). The mechanism that ultimately permits selective diVusion is also not known. Nevertheless, the actinomyosin cytoskeleton seems to be a central regulator of junction assembly and permeability (Hecht et al., 1996; Nusrat et al., 2000b; Turner, 2000; Wojciak‐ Stothard and Ridley, 2002). It is thus possible that regulation of RhoA drives junctional deformations via the actinomyosin cytoskeleton that permit the slow diVusion of small hydrophilic tracers. Claudin‐5, which is preferentially expressed by endothelial cells, has also been linked to specific increases in small tracer diVusion. Mice lacking claudin‐5 exhibit increased leakage of small tracers across the brain endothelium (Nitta et al., 2003). It is not clear, however, whether ion permeability is also aVected and how the absence of this claudin causes increased selective permeability. Similarly, claudins have also been linked to the maintenance of the epidermal barrier as both knockout of claudin‐1 and overexpression of claudin‐6 result in increased permeability (Furuse et al., 2002; Turksen and Troy, 2002).

B. Restriction of Apical/Basolateral Diffusion of Lipids Tight junctions also form an intramembrane diVusion barrier or fence that prevents the intermixing of lipids in the outer leaflet of the plasma membrane (Dragsten et al., 1981; van Meer and Simons, 1986). Our understanding of the structural basis of this intramembrane diVusion fence is relatively poor. Nevertheless, expression of a dominant negative mutant of the transmembrane protein occludin resulted in a disruption of the fence, suggesting that occludin either participates in the formation of the fence or in its regulation (Balda et al., 1996b). The junctional intramembrane fence is often also suggested to be important for cell surface polarity of proteins; however, experimental disruption of the fence without disrupting the entire junctional complex results in transjunctional diVusion of lipids but not in loss in protein

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polarity (Balda et al., 1996b; Jou et al., 1998). Thus, the junctional fence is important for the maintenance of cell surface lipid but not protein polarity.

IV. Regulation of Epithelial Polarization and Proliferation A. Epithelial Polarization and Junction Formation Assembly of intercellular junctions has been primarily studied in epithelial cells using various experimental systems such as epithelial cell lines (e.g., MDCK cells or the intestinal cell lines T84 and Caco‐2) or model organisms such as D. melanogaster, Caenorhabditis elegans, and mice (Cereijido et al., 2004; Fleming et al., 2001; Matter and Balda, 2003b; Tepass et al., 2001). We will therefore primarily discuss regulation of epithelial junction assembly. The regulation of junction assembly is linked to epithelial polarization. However, adhesion as such is not required to induce cell surface polarization, and polarization precedes cell–cell adhesion during fly development (Baas et al., 2004; Harris and Peifer, 2004; Le Bivic, 2005). The fundamental principles that govern junction assembly are well conserved. Generally, the formation of calcium‐dependent primordial adherens junctions is the first step of junction assembly (Ando‐Akatsuka et al., 1999; Cereijido et al., 1978; Ebnet et al., 2001; Suzuki et al., 2002). Primordial adherens junctions are based on adhesion proteins such as E‐cadherin and nectins but also recruit proteins that in a fully assembled junctional complex are no longer part of adherens junctions such as the tight junction component ZO‐1 and JAM‐A (Irie et al., 2004). The junctional complex then matures and establishes distinct adherens and tight junctions. Similarly, disassembly of the junctional complex can be triggered by calcium removal or inactivation of E‐cadherin with antibodies (Behrens et al., 1985; Gumbiner and Simons, 1986). Cell–cell adhesion seems to stimulate tight junction formation by the activation of intracellular signaling pathways as E‐cadherin inactivation can be overcome by stimulating PKA with cyclic AMP or protein kinase C (PKC) with diacylglycerol (Balda et al., 1993; Behrens et al., 1985). Moreover, other cellular signaling components such as monomeric and heterotrimeric G proteins have also been linked to the regulation of junction assembly (Benais‐Pont et al., 2001; Kohler and Zahraoui, 2005), and evolutionarily conserved junction‐associated signaling complexes have been identified that organize junction assembly and cell polarization (Macara, 2004). PKA is one of the key regulators of tight junction assembly. Stimulation of PKA prevents the disassembly of tight junctions upon antibody‐based neutralization of E‐cadherin or calcium removal (Behrens et al., 1985;

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Nilsson et al., 1996). Nevertheless, PKA inhibits tight junction assembly in calcium switch experiments, an experimental protocol in which cells are first cultured in low calcium medium and junction formation is then triggered by the addition of calcium (Balda et al., 1991; Cereijido et al., 1978). Hence, PKA activation inhibits both disassembly and assembly of junctions, suggesting that it counteracts junctional reorganization and that it needs to be tightly controlled to coordinate its activity with junction reorganization. Regulation of PKA activity has recently been shown to involve Rab13, a small GTP‐binding protein that associates with and regulates tight junction assembly (Marzesco et al., 2002; Zahraoui et al., 1994). The GTP‐bound form of Rab13 was shown to bind and inhibit PKA, which modulates the recruitment of junctional components (Kohler et al., 2004). Heterotrimeric G proteins can be inhibitory or stimulatory. Inhibitory G‐proteins were linked to tight junction formation when pertussis toxin was shown to stimulate assembly of tight junctions in MDCK cells (Balda et al., 1991). Gai2 localizes at cell–cell contacts, where its distribution overlaps with the one of ZO‐1 (de Almeida et al., 1994; Dodane and Kachar, 1996; Nilsson et al., 1996). However, ectopically expressed Gai proteins were shown to stimulate tight junction assembly (Denker et al., 1996). The reason for the contradiction between the pharmacological and the transfection experiments is not clear, but might be caused by the inhibition of several diVerent ai proteins by pertussis toxin. Involvement of multiple G proteins in tight junction regulation is supported by the recent finding that Ga12 is recruited to tight junctions by binding to ZO‐1 and results in increased paracellular permeability, an eVect that involves activation of Src tyrosine kinase (Meyer et al., 2002, 2003). The PKC family of serine/threonine kinases consists of classical (cPKC), novel (nPKC), and atypical (aPKC) PKCs. The cPKC (a, b, and g) and nPKC isotypes (d, Z, and y) are activated by diacylglycerol (DAG) and phosphoserine. Both cPKCs and nPKCc are also activated by phorbol esters, and cPKCs require calcium for activation. In contrast, aPKCs (z, l, and PRK for PKC‐related kinases) are calcium insensitive and do not respond to phorbol esters (Mellor and Parker, 1998). Phorbol esters are potent activators of PKC signaling and have been shown to induce disassembly of tight junctions in diVerent cell lines (Benais‐Pont et al., 2001; Clarke et al., 2000a). In contrast, inhibition of PKCs blocks junction assembly and disassembly, suggesting that PKC activity is important for dynamic junctional processes but its activation needs to be carefully controlled to prevent overstimulation and, hence, disruption of tight junctions (Balda et al., 1991; Citi et al., 1994; Nigan et al., 1991). The importance of DAG‐regulated PKCs in junction assembly is supported by experiments showing that the cell‐permeable DAG analogue dioctanoylglycerol can stimulate partial junction assembly in the absence of

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E‐cadherin‐mediated cell–cell adhesion (Balda et al., 1993). The PKC isoforms whose role in the regulation of junction assembly is best understood are aPKC z and l (see below). A role for Rho GTPases in junction assembly was first implied by experiments using microinjection of C3 transferase, a toxin that inactivates Rho, which inhibits the assembly of adherens and tight junctions (Nusrat et al., 1995; Takaishi et al., 1997; Zhong et al., 1997). However, expression of constitutively activated forms of RhoA and Rac1 also perturbs tight junctions in epithelia and endothelia linking these proteins to tight junction disassembly (Braga et al., 1999; Hasegawa et al., 1999; Jou et al., 1998; Takaishi et al., 1997; Wojciak‐Stothard et al., 2001). Moreover, inactivation of Rho family GTPases in brain endothelial cells inhibits the ability of lymphocytes to migrate across them, indicating that junctional dynamics requires Rho activation (Adamson et al., 1999). Rho family GTPase activities hence require careful regulation as neither high nor low levels of Rho activity are optimal for tight junction integrity. Additionally, some of the conflicting observations might be due to the diVerent cell types used as the eVect that Rho GTPases has on adherens junctions is also aVected by the cellular context (Braga, 2000). The actin cytoskeleton has a crucial role in the regulation of tight junction assembly and several tight junction components directly interact with actin filaments (Table I) (Fanning, 2001). Given the importance of Rho family GTPases in the regulation of the actin cytoskeleton (Etienne‐Manneville and Hall, 2002), it is likely that they regulate tight junctions by inducing changes in the actin cytoskeleton and this seems to involve diVerent mechanisms. Rho signaling can directly aVect tight junction proteins. For example, it has been suggested that Rho‐stimulated phosphorylation of occludin regulates occludin’s interaction with the submembrane cytoskeleton and thereby regulates its function in selective paracellular permeability (Hirase et al., 2001). Rho also influences tight junctions by mechanisms that involve regulation of cortical actin contraction through the activation of the motor protein myosin, a pathway that involves the Rho‐associated kinase/ROCK, which inactivates myosin light chain phosphatase and hence activates myosin II, which is thought to drive tight junction assembly and disassembly (Hecht et al., 1996; Ivanov et al., 2004; Turner et al., 1997). Little is known about tight junction‐specific mechanisms that control activation of Rho GTPases. Activation of Rho GTPases requires a GEF, and a tight junction‐associated GEF for Rho has recently been identified. GEF‐H1/Lfc was shown to associate with tight junctions and to regulate tight junction function (Benais‐Pont et al., 2003). However, it is not known whether GEF‐H1 is critical for the regulation of tight junction assembly. Two evolutionarily conserved signaling complexes that regulate cell polarization and junction assembly are associated with tight junctions: the

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CRB3/Pals1/PATJ complex and the Cdc42‐interacting Par3/Par6/aPKC complex. Components of both complexes are important for the generation of polarity in diVerent cell types and for the regulation of epithelial junction assembly. Studies focusing on the molecular mechanisms that regulate polarization during asymmetric cell division in C. elegans resulted in the isolation of the Par (partitioning defective) genes (Gotta and Ahringer, 2001). Two of these proteins, Par3 and Par6, form a complex with aPKC, which regulates cell polarity in diVerent organisms. In mammalian epithelia, the Par3/Par6/ aPKC complex associates with tight junctions and regulates junction assembly and polarization (Macara, 2004; Ohno, 2001). Par‐3, which is also called ASIP, and Par‐6 are PDZ domain proteins that are recruited to the forming junctional complex because of an interaction between JAMs and Par3 (Ebnet et al., 2001; Itoh et al., 2001; Izumi et al., 1998). Par6 is an eVector of Cdc42, a Rho family GTPase that is essential for epithelial cell polarity and becomes activated during junction formation: binding of Par6 to GTP‐bound Cdc42 triggers aPKC activation, which is important for the maturation of the junctional complex and tight junction formation (Joberty et al., 2000; Kroschewski et al., 1999; Lin et al., 2000; Suzuki et al., 2001, 2002). The components of the Par3/Par6/aPKC signaling complex influence diVerent types of downstream signaling mechanisms and not all involve the entire complex. A key substrate of aPKC is Par1b (MARK2/EMK1), a protein kinase that regulates the microtubule network, polarized membrane traYc, and, thereby, epithelial polarity (Cohen et al., 2004a,b; Suzuki et al., 2004). Par3 also regulates tight junction assembly via a Par6/aPKC‐independent mechanism by binding and regulating the Rac1 GEF TIAM1 (Chen and Macara, 2005). On the other hand, Par6 has been linked to a signaling pathway that triggers loss of the epithelial phenotype. Junctional dissociation during tumor growth factor (TGF)‐b‐induced epithelial mesenchymal transition (EMT) requires Par6 phosphorylation by the TGF‐b receptor type II. Phosphorylation triggers an interaction with the ubiquitin ligase Smurf1, which has been proposed to target junction‐associated RhoA for degradation and, hence, disintegration of the junctional complex (Ozdamar et al., 2005). The Par3/Par6/aPKC complex is also regulated by other cellular signaling mechanisms. aPKC activity is negatively regulated by protein phosphatase 2A, which dephosphorylates aPKC (Nunbhakdi‐Craig et al., 2002). As protein phosphatase 2A can dephosphorylate a variety of tight junction proteins and overexpression of its catalytic subunit in MDCK cells results in increased paracellular permeability, protein phosphatase 2A is probably a negative regulator of tight junctions, suggesting that Par3/Par6/aPKC and protein phosphatase 2A represent opposing pathways that regulate tight junction assembly and disassembly.

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The second evolutionarily conserved signaling complex associated with tight junctions is the CRB3/Pals1/PATJ or crumbs complex. Crumbs had first been identified in Drosophila, where it associates with stardust (the fly homologue of Pals1) and Drosophila PATJ. In flies, this complex localizes to the subapical complex/marginal zone, the most apical junctional structure in flies (Tepass, 2002). Crumbs regulates the morphogenesis of ectodermal epithelia and regulates apical membrane biogenesis. In mammals, three crumbs homologues have been identified, but only CRB3 has thus far been localized in epithelial cells and shown to be distributed along the entire apical membrane including the apical end of tight junctions, where it forms a complex with Pals1 and PATJ (Lemmers et al., 2002; Makarova et al., 2003; Roh et al., 2002). Functional experiments in cultured cell lines using RNA interference and overexpression suggest that all three proteins regulate tight junction assembly (Fogg et al., 2005; Lemmers et al., 2002, 2004; Shin et al., 2005; Straight et al., 2004). However, defects in polarization were not observed in normal two‐dimensional culture systems but became evident when cells were grown in collagen gel experiments; hence, the CRB3/Pals1/ PATJ complex might be more critical for polarization under conditions that require a more dynamic coordination among proliferation, junction assembly, and diVerentiation. The CRB3/Pals1/PATJ and the Par3/Par6/aPKC complexes are not independent but interact with each other. Par6 binds Pals1 and CRB3, and the interaction between Par6 and Pals1 is regulated by Cdc42 (Hurd et al., 2003; Lemmers et al., 2004). In Drosophila, aPKC phoshorylates and thereby activates crumbs (Sotillos et al., 2004). Although it is not known whether mammalian aPKCs phosphorylate CRB3, it is possible that adhesion‐ induced recruitment and activation of the Par3/Par6/aPKC complex triggers activation of the CRB3/Pals1/PATJ complex by phosphorylation of the transmembrane protein.

B. Epithelial Proliferation and Gene Expression Intercellular junctions contribute to the regulation of proliferation and diVerentiation, and aVect signaling along major signal transduction pathways. Expression of several tight junction components is aVected in diVerent types of carcinomas; however, how much these alterations are a cause or a consequence of carcinogenesis is not clear. For example, it has recently been suggested that claudin‐1 promotes transformation and metastatic behavior in colon cancer (Dhawan et al., 2005). Another claudin family member, claudin‐11, had previously been shown to promote proliferation and migration by forming a complex with OAP‐1, a tetraspanin, and b1‐integrin (Tiwari‐WoodruV et al., 2001). Hence, claudins not only function in the

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junctional diVusion barrier but appear to play extrajunctional roles in the regulation of cell proliferation and migration. Cytosolic components of tight junctions have also been linked to the regulation of cell proliferation, diVerentiation, and oncogenesis. The adaptor proteins ZO‐2, MAGI‐1, and MUPP1 can bind and inactivate viral oncogenes, and oncogenes, tumor suppressors, and cell cycle regulators localize to tight junctions (Table I) (e.g., the tyrosine kinase c‐yes, the RhoA GEF GEF‐ H1/Lfc, the lipid phosphatase PTEN, and the cell division kinase CDK4) (Balda et al., 2003; Benais‐Pont et al., 2003; Chen et al., 2002; Glaunsinger et al., 2000, 2001; Lee et al., 2000; Matter and Balda, 2003b; Tolkacheva et al., 2001; Wu et al., 2000a,b). Moreover, tight junctions recruit several NACos that function in the regulation of gene expression such as the transcription factors ZONAB and huASH1 (Balda and Matter, 2000a, 2003; Nakamura et al., 2000). In general, it is thought that binding of junctional adaptors to oncogenes and stimulators of proliferation results in their inactivation, suggesting that tight junctions function as suppressors of proliferation. In many cancers, epithelial dediVerentiation correlates with deregulation of Ras signaling. Tight junctions have been directly linked to the modulation of Ras signaling pathways. Overexpression of the junctional membrane protein occludin is able to suppress transformation of salivary epithelial cells induced by Raf‐1, a common Ras eVector that stimulates cell cycle entry via ERK/MAP kinase activation (Li and Mrsny, 2000). Although the molecular mechanism that underlies this inhibitory activity of occludin is unknown, it is striking that it requires the extracellular domain of occludin that binds to TGF‐b receptor type I (Barrios‐Rodiles et al., 2005; Wang et al., 2005). TGF‐b receptor type I binding to occludin recruits the receptor to tight junctions, which is critical for disruption of the junctional complex during TGF‐b‐induced EMT (Barrios‐Rodiles et al., 2005). ERK activation is important for TGF‐b‐induced EMT (Xie et al., 2004); therefore, occludin may not only function as a suppresser of Raf‐1 but may coordinate Raf activation. A role of occludin in the modulation of signaling pathways that control epithelial diVerentiation is also suggested by the finding that deletion of the occludin gene in mice aVects the diVerentiation of some epithelial cell types (Saitou et al., 2000). Occludin has also been linked to the regulation of RhoA signaling. The small GTPase RhoA is a central regulatory switch that controls multiple cellular processes such as G1/S phase progression, cytokinesis, and cell shape changes (Coleman et al., 2004; Etienne‐Manneville and Hall, 2002; Ridley, 2004). MDCK cells with strongly reduced occludin expression fail to activate RhoA in response to certain stimuli (Yu et al., 2005). Although the molecular mechanism by which occludin stimulates RhoA activation has not been identified, it is possible that this involves GEF‐H1/Lfc, a tight

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junction‐associated member of the Dbl family of oncogenes that functions as a GEF for RhoA (Benais‐Pont et al., 2003). GEF‐H1/Lfc is a RhoA regulator that contributes to diVerent steps along the cell cycle as it has been shown to regulate G1/S phase progression as well as to promote assembly of the mitotic spindle (Aijaz et al., 2005; Bakal et al., 2005). In agreement, GEF‐H1/Lfc is down‐regulated by the c‐KIT inhibitor and antitumor drug Gleevec, suggesting that this GEF is a physiologically and pathologically important regulator of cell cycle progression (Frolov et al., 2003). GEF‐H1/Lfc is inhibited by binding to the junction‐associated adaptor cingulin (Aijaz et al., 2005). Because up‐regulation of cingulin correlates with cell cycle arrest and diVerentiation (Aijaz et al., 2005; Bordin et al., 2004), inhibition of RhoA signaling is another mechanism by which tight junctions suppress proliferation. Adhesion complexes participate in the regulation of gene expression by controlling the localization and activity of NACos, proteins that can localize to the nucleus and adhesion complexes (Balda and Matter, 2003). An interesting example of a tight junction‐associated NACo is ZO‐2, which enters the nucleus in proliferating cells and interacts with the hnRNP protein SAF‐B and transcription factors AP‐1 and C/EBP (Betanzos et al., 2004; Traweger et al., 2002b). ZO‐2 also regulates the nuclear accumulation of ARVCF, a member of the p120(ctn) family (Kausalya et al., 2004). These interactions of ZO‐2 might be of pathological relevance as its expression is deregulated in diVerent types of adenocarcinomas (Chlenski et al., 2000). ZO‐1 is a negative regulator of proliferation in epithelial cells and this activity maps to its SH3 domain (Balda et al., 2003). One of the interaction partners of ZO‐1, the Y‐box transcription factor ZONAB, binds to the SH3 domain and promotes proliferation (Balda and Matter, 2000a; Balda et al., 2003). ZO‐1 inhibits ZONAB by cytoplasmic sequestration. Although depletion of ZO‐1 does not aVect proliferation of all analyzed cell lines (Umeda et al., 2004), overexpression and depletion of ZONAB and ZO‐1 influence proliferation and the cell densities of renal and mammary epithelial cell lines (Balda et al., 2003; Fig. 2; our unpublished data). This ZONAB/ZO‐1 signaling mechanism appears to be of physiological and pathological relevance as the human homologue of canine ZONAB, DbpA, is an E2F1 target gene and is overexpressed in diVerent types of carcinomas (Arakawa et al., 2004). The mouse ZONAB homologue, Yb‐3, has also been colocalized with ZO‐1 at gap junctions of oligodendrocytes and astrocytes, suggesting that ZONAB/ ZO‐1 signaling might also be important in nonepithelial cell types (Penes et al., 2005). ZONAB promotes proliferation by regulating G1/S phase progression of the cell cycle. This involves two mechanisms: first, ZONAB interacts with the cell division kinase CDK4, a G1/S phase regulator, resulting in cosequestration of the kinase and the transcription factor in the cytoplasm when junction

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FIG. 2 Regulation of epithelial cell density by ZO‐1. ZO‐1/ZONAB signaling regulates the final cell density in MDCK cell monolayers (Balda et al., 2003). Since overexpression of ZO‐1 reduces cell density, decreased ZO‐1 expression should increase cell density. To test this, ZO‐1 was constitutively down‐regulated using RNAi by targeting two diVerent sequences (RD1: 50 – AAGATAGTTTGGCAGCAAGAG–30 ; RD2: 50 –AATGTCCCTGATCTTTCT GAC–30 ). (A) Immunoblots of confluent MDCK cells stably expressing the indicated RNA duplexes that were obtained with antibodies against ZO‐1, ‐2, and ‐3, as well as a‐tubulin. Note that only expression of ZO‐1 is aVected. (B) Cell density of confluent MDCK cells was quantified by counting. Shown are averages of four clones for ZO‐1 RNAi and three clones for control RNAi.

formation and, hence, ZO‐1 expression is upregulated (Balda et al., 2003); and second, ZONAB functions in the transcriptional regulation of cell cycle regulators (unpublished data). In confluent epithelial cells, ZONAB also interacts with the small GTPase RalA, which counteracts the transcriptional activity of the Y‐box factor (Frankel et al., 2005). RalA is part of an eVector pathway of Ras; however, RalA activation in normal MDCK cells is Ras independent. Because the amount of active RalA is not aVected by cell density but the interaction with ZONAB occurs only in dense cells, RalA does not directly aVect the localization of ZONAB but might be a switch that regulates the activation state of ZONAB. Y‐box factors are multifunctional proteins that are also thought to regulate RNA turnover and translation (Matsumoto and WolVe, 1998); hence, RalA might regulate such possible cytoplasmic functions of ZONAB. Another component of the cytoplasmic plaque, symplekin, participates in nuclear as well as cytoplasmic polyadenylation, but it is not known whether symplekin and ZONAB are functionally related (Barnard et al., 2004; Hofmann et al., 2002; Takagaki and Manley, 2000). Tight junctions might hence be important not only for transcriptional regulation of gene expression but also for processes such as the regulation of mRNA stability, localization, and translation.

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V. Concluding Remarks Research in many diVerent laboratories resulted in the discovery of an intriguingly large spectrum of proteins that associates with tight junctions. Many interactions between these proteins as well as between them and the cytoskeleton have been mapped, suggesting that tight junction components form a complex cytoskeleton‐linked protein network. For many junctional proteins, however, we do not know much about their function and how important their interactions are. Moreover, many junctional components are able to bind to so many other proteins that it is unlikely that they associate with all of them at once and recent results indeed suggest that tight junction proteins form distinct subcomplexes (Vogelmann and Nelson, 2005). It will thus be important to identify the role of particular interactions for junction assembly, to determine when they occur, and for which tight junction functions they are important. As many recent gene knockout and RNA interference studies suggest, the junctional composition is not only complex but appears to contain many redundant components. Hence, functional analysis of particular junctional components cannot rely only on single knockouts and knockdowns, but will require manipulation of several components together and the use of dominant mutants. A major challenge still remains the identification of the precise molecular structures and mechanisms that permit selective paracellular diVusion of ions and small hydrophilic tracers. Although it is clear that claudins are important for ion permeability and some of their structural features important for ion conductance have been identified, we still do not understand if and how they form the proposed paracellular channels. The mechanism that mediates permeability of small hydrophilic tracers is even more poorly defined. As tracer permeability and ion permeability are often regulated independently, it is unlikely that the two mechanisms are directly related; hence, tracer permeability may not rely on paracellular pores. In terms of the signaling function of tight junctions, many junctional components are now known that regulate epithelial polarization, but we still know very little about how these polarity complexes transduce signals and stimulate cell polarization. Moreover, it will be essential to determine how they influence more conventional cellular signaling components such as PKA and GTP‐binding proteins, and how polarity complexes together with the junctional NACos contribute to carcinogenesis and metastasis. Acknowledgments We would like to thank Peter Munro for electron micrographs. Research in the authors’ laboratories is supported by the Wellcome Trust, Fight for Sight, as well as the Medical Research Council and the Biotechnology and Biological Sciences Research Council.

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