EDITORIAL FOCUS
Claudins form ion-selective channels in the paracellular pathway.
Focus on "Claudin extracellular domains determine paracellular charge selectively and resistance but not tight junction fibril architecture"

Eveline E. Schneeberger

Molecular Pathology Unit, Department of Pathology, Massachusetts General Hospital, Charlestown, Massachusetts 02129


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THE ABILITY OF EPITHELIA to form a diffusion barrier between cellular compartments of very different fluid and solute composition is dependent not only on asymmetrically distributed transcellular transport mechanisms (transcellular pathway) but also on structures that regulate the diffusion of ions and small, noncharged solutes through the paracellular pathway. Although those involved in regulating solute and water transport across the transcellular route are well understood, the mechanisms governing the paracellular pathway have only recently begun to be examined. At the apical end of the paracellular space, adjacent cell membranes are in close apposition, a site that was termed by early anatomists as the "terminal bar" (2) and was considered to be an impermeable barrier in the paracellular space. It was not until the elegant ultrastructural studies of Farquhar and Palade (8) that the terminal bar was shown, in fact, to consist of a junctional complex composed of an apical tight junction (zonula occludens), an intermediate junction (zonula adherens), and a desmosome (macula adherens). Analysis of freeze-fracture replicas indicated that tight junctions form a circumferential network of anastamosing strands of varying complexity located in the plane of the plasma membrane (20). Furthermore, it was suggested that the number of parallel tight junction strands might correlate with the level of measured transepithelial electrical resistance (TER) (5). When the number of strands was plotted against TER, however, it was found that the relationship between the increase in TER with each additional strand was not a linear, but an exponential, one. This led to the suggestion that tight junction strands contain pores that flicker between an open and closed conformation (4).

Electrophysiological measurements had indicated that whereas the plasma membrane contains pores of 0.4 nm radius (18), pores of 3-4 nm radius are present in the paracellular space (13). In other words, the tight junction barrier in the paracellular space is considerably more permeable to water and small solutes than the plasma membrane. Furthermore, tight junctions appeared to be capable of discriminating between ions of similar charge and, in general, to be predominantly cation selective (15). This led to the prediction that tight junctions must contain aqueous pores lined by proteins, the amino acid composition of which would determine the charge selectivity of the tight junction pores.

Initial studies designed to identify integral tight junction proteins resulted in the discovery of two important tight junction-associated cytoplasmic proteins, ZO-1 (22) and cingulin (3). It was not until 1993, however, that the molecular composition of the tight junction strands began to be clarified in a series of groundbreaking studies by the Tsukita group. The first of these was occludin (11), a ~58-kDa tetraspan phosphoprotein, which, when overexpressed in MDCK cells, resulted in an elevation of TER and a paradoxical increase in the flux of small, water-soluble solutes (1, 14). However, when occludin-null embryonic stem cells were found to differentiate into tight junction-expressing cells (16), it became clear that other protein(s) must contribute to the formation of tight junction strands and that the precise function of occludin in the tight junction remained to be established. This was further supported by the observation that occludin-null mice survived to adulthood, although they developed a complex phenotype (17).

In their quest to identify other integral tight junction proteins, the Tsukita group re-examined the junction fraction from their original chicken liver preparations. The search yielded two novel, ~23-kDa integral tight junction proteins, claudins-1 and -2 (9). They, like occludin, are tetraspan proteins; however, the claudins share no sequence homology with occludin. In addition, in contrast to occludin, when claudin-1 cDNA was transfected into fibroblasts, they formed a network of tight junction strands in the plasma membrane, indicating that claudin is necessary and sufficient to form tight junction strands (12). To date, a >20-member family of claudins has been recognized (23) and these, either singly or in combination with several different claudins, are expressed in a cell- and tissue-specific pattern. Although clearly indicating that claudin(s) are the primary component of the tight junction strands, their contribution to the formation of the predicted ion- selective pores within the tight junction strands was unclear. A significant breakthrough was achieved with the report that a cohort of kindreds suffering from a rare, autosomal recessive form of hypomagnesemia associated with a distal renal tubular defect had a variety of mutations in the claudin-16 gene (19). Claudin-16 is uniquely expressed by epithelial cells lining the thick ascending limb of Henle, where it serves as a paracellular divalent cation-selective channel. Mutations in the claudin-16 gene are associated with a defect in magnesium reabsorption, which results in turn in the urinary loss of this ion. From these data, it appears that claudin-16 forms a Mg2+-selective channel in the epithelial tight junctions of the thick ascending limb of Henle. Further evidence that claudins might be involved in forming ion-selective channels was obtained in experiments utilizing two types of Madin-Darby canine kidney (MDCK) cells. High-resistance MDCK I cells (TER >1,000 Omega .cm2) and low-resistance MDCK II cells (TER <100 Omega .cm2) are interesting in that analysis of freeze-fracture replicas indicates that the number of tight junction strands is the same in these two cell lines (21). However, when MDCK I cells that lack endogenous claudin-2 were transfected with claudin-2 cDNA, the TER fell to a level that was similar to that of low-resistance MDCK II cells that are known to express endogenous claudin-2 (10). This led to the speculation that the type and combination of claudins determines the paracellular permeability of a given epithelium.

These experiments were a prelude to a series of elegant studies conducted by the Anderson group (6, 7), including the current article in focus (Ref. 7, see p. C1336 in this issue), which examined the role of the two extracellular domains of selected claudins in the formation of ion- selective channels within tight junction strands. In the first, site-directed mutagenesis was utilized to reverse the net charge on the first extracellular loop of claudins-4 and -15 (6). These mutated claudins were then expressed under an inducible promoter in MDCK cells. When a single negative charge was substituted for a positive charge in the first extracellular domain of claudin-4, paracellular Na+ permeability was increased. Conversely, when one or more negative charges on the first extracellular loop of claudin-15 were replaced singly or in combination with positive charges, the paracellular charge selectivity changed from a Na+- to a Cl--selective one. While these are convincing and important observations, two notes of caution are raised. First, site-directed mutagenesis resulting in amino acid substitutions may lead not only to a change in net charge but also to an altered molecular conformation of the protein in question. Second, overexpression of a given protein can result in its disproportionate incorporation into the tight junction with a concomitant replacement and/or displacement of other endogenous tight junction proteins, a result that may alter paracellular ion selectivity. Finally the overexpressed protein may be expressed in an aberrant location.

The first of these concerns is neatly circumvented in the current study, in which chimeric molecules are constructed by engrafting either the first or second or both extracellular domains of claudin-4 onto claudin-2 and vice versa (7). This approach accomplishes two things. 1) It circumvents the potential problems inherent in site-directed mutagenesis, and 2) it provides a means to examine the functional contribution of specified segments of the claudin in question. Because these experiments were conducted in cells expressing endogenous tight junction proteins, the authors acknowledge that the expression system utilized may lead to altered expression and/or localization of endogenous proteins. These potential problems aside, the study clearly shows that expression of the first or both extracellular domains of claudin-4 on claudin-2 markedly elevates TER and decreases the permeability of Na+ relative to Cl-. By contrast, engrafting the first or both extracellular domains of claudin-2 onto claudin-4 has only a modest effect on TER and charge selectivity. Furthermore, although these studies indicate that the larger, first extracellular loop of claudins-2 and -4 appears to confer many of the charge-selective characteristics, the function of the shorter second loop is unclear, because chimeric expression of the second loop alone produced biologically inactive molecules. Perhaps one of the most novel observations in the present study is that expression of the first or both extracellular domains of claudin-4 on claudin-2 has a more marked effect on TER and charge selectivity than does native claudin-4. This suggests that other segments of the molecule, including the carboxy terminus, may regulate, possibly by interactions with cytoskeletal proteins, the barrier function of the tight junction. Clearly, further studies addressing these questions will yield important new insights into the molecular structure and function of the tight junction.


    FOOTNOTES

Address for reprint requests and other correspondence: E. E. Schneeberger, Molecular Pathology Unit, Rm. 7147, 149 13th St., Charlestown, MA 02129 (E-mail: eschneeberger{at}partners.org).

10.1152/ajpcell.00037.2003

Received 23 January 2003; accepted in final form 27 January 2003.


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Am J Physiol Cell Physiol 284(6):C1331-C1333
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