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1 Department of Internal Medicine and Cell Biology
2 Department of Molecular Biophysics and Biochemistry, Yale University School of Medicine, New Haven, Connecticut 06520, USA
3 Department of Cell and Molecular Physiology, University of North Carolina-Chapel Hill, Chapel Hill, North Carolina 27599, USA
Reprint requests to: James Melvin Anderson, Department of Cell and Molecular Physiology, University of North Carolina-Chapel Hill, 266 Medical Sciences Research, Building CB# 7545, Chapel Hill, NC 27599-7545, USA; e-mail: jandersn{at}med.unc.edu; fax: (919) 966-6413.
(RECEIVED July 2, 2002; FINAL REVISION October 18, 2002; ACCEPTED October 31, 2002)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.0233903.
4 Present address: Department of Biochemistry and Biophysics, University of California, San Francisco, 513 Parnassus Ave., San Francisco, CA 94143-0448, USA 
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Keywords: Claudin; tight junction; perfluoro-octanoic acid (PFO); oligomer
Abbreviations: cldn-4, claudin-4 • PFO, perfluoro-octanoic acid • DDM, dodecyl maltoside • OG, ß-octylglucoside • DHPC, diheptanoyl-glycerol-3-phosphocholine • DOC, sodium deoxcholate • ZW-16, zwittergent-16 • LLS, laser light scattering
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Introduction |
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23 kD) and occludin (65 kD) have cytoplasmic N- and C-termini with two extracellular loops (Furuse et al. 1993; Furuse et al. 1998a), and extracellular adhesive loop contacts presumably create the tight junction seal. The physical structure of these contacts is unknown, although it is expected to be highly unique since the resultant pore allows selective passage of ions through the paracellular space between cells without generating a channel in the plasma membrane. Whether claudin physically interacts with occludin and whether claudins can form heteromeric assemblies is also unknown. Since endogenous claudins are in low abundance in vertebrate epithelial cells, we overexpressed cldn-4 in a heterologous cell system to make available the amount necessary for biophysical characterization. Functionally, the tight junction is a discriminating barrier, exhibiting ionic and size selectivity (for review, see Reuss 2001). Claudins are directly responsible for the charge selective movement of ions in the paracellular space (Van Itallie et al. 2001; Colegio et al. 2002). Claudin-4 overexpressed in MDCK cells creates monolayers with decreased permeability to sodium (Van Itallie et al. 2001), suggesting claudin-4 selectively restricts passage of sodium in the paracellular space. Reversing the charge of a single residue in the first extracellular loop of claudin-4 from positive to negative reverses the sodium permeability so that it is increased, suggesting charge selectivity is conferred by individual residues in the extracellular loops. Tight junctions of different organs and cell types exhibit widely varying junctional characteristics, presumably created by differing combinations of claudins. Expression profiles support this hypothesis, with distinct localization of claudins in the rat digestive tract (Rahner et al. 2001), nephron (Simon et al. 1999; Enck et al. 2001; Kiuchi-Saishin et al. 2002), testis (Gow et al. 1999), and ear (Wilcox et al. 2001). How claudins assemble in three dimensions to create this selective pore is unknown.
Comparison with other intramembrane protein particles of known composition suggest tight junction particles are composed of claudin multimers. Each particle is approximately 10 nm in diameter. 10-nm particles are also observed in freeze fracture replicas of gap junctions (Goodenough 1976) and Xenopus oocytes overexpressing a neuronal glutamate receptor (Eskandari et al. 2000). In both cases the 10-nm particle corresponds to an oligomer of membrane proteins; six connexins and five glutamate receptors, respectively. Claudins are smaller proteins (23 kD), suggesting that 10-nm particles at the tight junction are composed of multimers of claudins. In the present study, we overexpressed claudin-4 in insect cells using the baculovirus expression system and characterized its detergent solubility and oligomerization state. Claudin-4’s behavior in SDS and perfluoro-octanoic acid suggests it forms multimers. However, stable oligomeric forms were not observed in other detergents, suggesting the supramolecular organization of claudins in tight junctions varies significantly from the rigid organization of tetrapan oligomers at other types of junctions such as gap junctions.
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23 kD (Fig. 1
), the expected molecular mass of claudin-4. This band was also visible in Coomassie-stained gels of membrane preparations (Fig. 1), demonstrating robust overexpression of this small membrane protein.
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Claudin-4 is variably solubilized by multiple detergents
To determine the solubility of claudin-4 expressed in insect cells, we exposed crude Sf9 membranes from infected cells to a panel of 39 detergents. Our choice of detergents was motivated by differences in head groups and hydrophobic regions—the 39 used cover the main classes of detergents. Identical amounts of membrane were used to compare the effectiveness of each detergent. To ensure complete solubilization, the amount of detergent used was equivalent to the mass of the dried membrane pellet plus 1 x the critical micelle concentration (CMC). Samples were agitated for 1 h at room temperature, centrifuged for 30 min at 100,000g, and the resultant supernatant was mixed with SDS sample buffer and loaded on a 13% SDS-PAGE gel. As visible in Figure 2, cldn-4 is variably soluble in 33 of 39 detergents. These results are quantitated in Table 1. In general, ionic and phospholipid-like detergents were most effective at liberating cldn-4 from the membrane. However, nonionic detergents solubilized only a small fraction or no percentage of cldn-4. These results identified a number of candidate detergents for further characterization and purification of cldn-4.
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). Evaluation of the same sample on an 8% gel did not reveal additional high molecular weight adducts, confirming six as the total number of visible adducts (data not shown). SDS-resistant adducts are routinely seen with other membrane proteins, such as connexins. These results suggest PFO permits maintenance of oligomeric claudin-4 species and SDS is unable to completely disrupt a complex previously exposed to PFO.
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50 to >140 kD, with most immunoreactivity concentrated at 120 kD, which is consistent with a hexameric configuration (Fig. 4A). Although bands are typically diffuse in this system because of the lack of a focusing step, the broadness of this band suggested multiple oligomeric species exist in PFO lysates. Mass was approximated based on comparison with the linear relationship between the distance of migration and the logarithm of molecular weight of soluble protein standards (Fig. 4B).
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). In this experiment, membranes are exposed solely to PFO before loading on the PFO-containing sucrose gradient and mixed with SDS sample buffer only after fractionation. Thus, the HMW species are not artifacts from exposure to SDS, since they migrate as discrete species before being exposed to SDS.
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, samples increase in apparent molecular mass across the gradient. A PFO sucrose gradient was fractionated as in Figure 5
, and samples were mixed with PFO sample buffer (instead of SDS as above) and electrophoresed on a 4%–20% PFO-PAGE gel. Estimated sizes range from 50 kD in fraction 7 to 120 kD in fraction 13. These data confirm PFO solubilization produces a range of oligomeric species that can be resolved by velocity gradient centrifugation.
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, depending on the detergent, we observed a peak near the top or middle of the gradient. Protein solubilized in ß-octylglucoside (OG) or diheptanoyl-glycerol-3-phosphocholine (DHPC) sedimented with a peak centered at fraction 4 (Fig. 7). Similar results were obtained with Triton X-100, sodium deoxycholate (DOC), and zwittergent-16 (ZW-16) (data not shown). Protein solubilized by CHAPS sedimented with a peak in fraction 5 (data not shown). Based on sedimentation of soluble protein markers, the peak at fractions 4–5 likely represents a claudin monomer. In contrast, solubilization in dodecyl maltoside (DDM) produced a peak centered at fraction 8, consistent with a larger oligomeric species (Fig. 7). These sedimentation profiles suggested that DDM maintained a larger oligomeric species, while OG, DHPC, DOC, ZW-16, and CHAPS disrupted the oligomer during solubilization. However, an estimate of the approximate molecular mass of the DDM species by comparison with sedimentation of soluble protein standards is complicated by the contribution of an unknown amount of bound detergent. Therefore, to obtain the molecular mass of the DDM species, we performed high-pressure liquid chromatography together with laser light scattering–a technique that is insensitive to the bound detergent (Hayashi et al. 1989).
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), we expected LLS would estimate the molecular mass of monomeric cldn-4 when solubilized in OG and oligomeric cldn-4 when solubilized in DDM. Protein was loaded onto a detergent equilibrated Superdex 200 gel filtration column, and UV, light scattering, and refractive index detectors analyzed eluate. Molecular mass was estimated using the three-detector method of Hayashi et al (1989), which is insensitive to the amount of bound detergent or lipid (Hayashi et al. 1989). Surprisingly, the estimated molecular mass of protein solubilized in either DDM or OG was 23 kD, the size of monomeric claudin-4. These data suggest the shifted sedimentation profile of DDM-solubilized claudin-4 seen by sucrose velocity centrifugation represented an anomalous shift not related to oligomeric state, perhaps produced by a highly compacted conformation that contributed to a larger sedimentation value. |
Discussion |
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We expressed claudin in insect cells because traditional biochemical and biophysical studies of endogenous claudin-4 in epithelial cells are complicated by its interactions with multiple endogenous cytoplasmic binding partners. Claudins interact with the PDZ-containing proteins ZO-1, ZO-2, ZO-3 (Itoh et al. 1999), MUPP1 (Hamazaki et al. 2002), and PATJ (Roh et al. 2002). Further, it is not known whether claudin interacts with occludin, another fibril-forming transmembrane protein. A detergent that maintains claudin’s oligomeric interactions would also likely maintain claudin’s interactions with other proteins. The contributions of these additional proteins to claudin’s sedimentation in sucrose velocity gradients or migration in native gels would complicate any possible interpretation. Since insect cells do not make tight junctions, expression in these cells should obviate these concerns.
However, the relevance of our observations in this heterologous cell system to claudin-4’s assembly under native conditions in vertebrate epithelial cells depends on the assumption that claudin-4’s oligomerization state in Sf9 cells mimics its native state. It is possible our inability to find a conventional detergent that maintains an oligomer reflects a limitation in the expression system. For instance, incorrect lipid composition, a missing unknown stabilization cofactor, or a requirement for transcellular contact may bias our results towards the monomer. Thus, it is possible that claudin-4 may readily assume a multimeric configuration stable in a variety of detergents when in its native epithelial cell environment. However, other oligomeric membrane proteins, such as connexin 32 (Stauffer et al. 1991) and the Shaker voltage-gated K+ channel (Li et al. 1994), adopt their native quaternary structures when expressed in Sf9 cells. This suggests our results are not a general failure of membrane proteins to form stable oligomers in these cells. Moreover, our PFO results argue that an oligomeric claudin state does exist in Sf9 cells. Specifically, the migration of claudin-4 centered at 120 kD on a 4%–20% PFO gel is consistent with a hexameric claudin confirmation. In addition, six SDS-resistant claudin-4 bands are visible on SDS gels of PFO-solubilized membranes, which is also consistent with a hexamer. There is no evidence that PFO artifactually creates oligomers (Ramjeesingh et al. 1999), suggesting these data represent in vivo oligomers.
The ability of all conventional detergents tested to disrupt the claudin oligomer suggests a highly dynamic and unstable configuration. This is in contrast to other oligomers, such as connexons, which require harsh conditions to disrupt. Why would a claudin oligomer be so labile in the membrane? Evidence exists suggesting this might reflect unique in vivo properties of claudins. First, rapid massive assembly of fibrils occurs under various conditions that induce cellular stress, presumably as a protective mechanism (Kachar and Pinto da Silva 1981; Lynch et al. 1995). Kachar and Da Silva (1981) reported the emergence of tight junction strands along the entire length of lateral membranes from excised rat prostate tissue incubated at 37°C at the earliest time point tested (3 min). Similarly, Lynch et al. (1995) reported the appearance of aberrant fibrils extending laterally from tight junctions within 5 min after basolateral exposure of monolayers to protease. These examples of rapid fibril proliferation support a model where a preexisting pool of fibril components can rapidly reorganize to create functional tight junction fibrils. These data indicate the fibrils themselves are dynamic structures, and allow the speculation that their components may in turn be equally dynamic.
Additional support for dynamic oligomers comes from the paradoxical relationship between transepithelial electrical resistance (TER) and flux. Experimental manipulations that increase TER, an instantaneous measurement of ion movement through tight junctions, could be expected to decrease flux, which measures the passage of larger molecules through the junction over a longer time course (min). In other words, as the junction becomes electrically tighter, the movement of larger uncharged molecules should be retarded. However, the opposite is seen. Overexpression of occludin (Balda et al. 1996; McCarthy et al. 1996) and claudin (McCarthy et al. 2000; Van Itallie et al. 2001) increases fibril number and TER while reducing or not affecting flux. Movement of ions through claudin pores is directly dependent on the charged residues in the extracellular loops of claudins (Colegio et al. 2002). It is difficult to envision an aqueous pore with ion selectivity that can also accommodate large uncharged molecules. Thus, it has been proposed that uncharged molecules pass though transient breaks in fibrils (for review, see Mitic and Anderson 1998). Dynamic claudin oligomers may promote this proposed strand breakage and reformation, thereby enabling the uncoupling of TER and flux.
Our studies describe the robust expression of claudin-4 in insect cells and its biochemical characteristics, which will be helpful in future structural studies. Determining the structure of the claudin pore is of fundamental importance in understanding epithelial barrier function. These data provide a useful starting point for future structural approaches that will ultimately provide invaluable insight into the paracellular pathway.
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Materials and methods |
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Recombinant baculovirus construction and infection
The human claudin-4 cDNA (Van Itallie et al. 2001) was PCR-amplified and decahistidine tagged using primers 5'-CGGGATCCCTGACAATGGCCTCCATGGGGCTAC-3' (sense) and 5'- GCTTAATGATGATGATGATGATGATGATGATGATGCTTG TCTGTGCGGGGTGGACA-3'(antisense). The resultant amplicon was shuttled through pCR2.1-Topo (Invitrogen) and subcloned into pFastBac1 (Gibco BRL). Generation of recombinant virus using the Bac-to-Bac baculovirus expression system (Gibco BRL) was according to manufacturer’s instructions. First-generation virus was amplified, clarified by low-speed centrifugation, and stored at 4°C. Sf9 or High-Five cells were infected in suspension at a MOI of 7–10, incubated for 52–56 h and harvested by centrifugation.
Insect cell membrane preparations
An infected cell pellet from a 3 L culture (25 mL) was washed 2x with 500 mL ice-cold PBS, resuspended in 75 mL ice-cold 20 mM Hepes, pH 7.4, 5 mM EDTA, 1 mM PMSF plus a 10 µg/mL protease inhibitor cocktail (leupeptin, antipain, aprotinin, and trypsin-chymotrypsin inhibitor in 10 mM Hepes, pH 7.4, Sigma) and incubated on ice for 10 min. Cells were sonicated on ice 5x for 15 sec each at power level 6, duty cycle 3 (Branson Sonifier 250). Light microscopy revealed 100% of cells were disrupted under these conditions. Cell membranes were pelleted by centrifugation for 90 min at 39,000 rpm at 4°C in a SW41 rotor (Beckmann). Peripheral membrane proteins were extracted by resuspending this pellet in 75 mL of ice-cold high-salt wash (20 mM Hepes, pH 7.4, 5 mM EDTA, 1 mM PMSF plus protease inhibitor cocktail and 300 mM NaCl), followed by sonication and centrifugation as before. Pelleted membranes were resuspended in 50 mL of either 20 mM Hepes, pH 7.4, 10 mM NaCl for detergent extraction studies, or Ni-NTA lysis buffer (50 mM NaH2PO4, 300 mM NaCl, and 10 mM imidazole, pH 8.0) for protein purification. Membrane droplets were snap frozen directly in liquid N2, collected into 50 mL tubes and stored at -80°C.
Detergent extraction
100 µL aliquots of infected membranes were mixed 1:1 (v/v) with freshly prepared detergent equivalent to the mass of 100 µL dried membranes plus 1x the critical micelle concentration. Detergents were obtained from Anatrace or Avanti Polar Lipids. Extraction was performed in 20 mM Hepes, pH 7.4 containing either 10 mM NaCl ("low salt" extraction) or 210 mM NaCl ("high salt" extraction). Samples were stirred using a magnetic "flea" stir bar for 1 h at room temperature, spun 30 min at 100,000g at room temperature in a Beckman TLA-100 rotor, and the supernatant containing solubilized claudin-4 was removed. Supernatant was mixed with 5x SDS loading buffer and loaded onto 13% SDS-PAGE gels.
PFO-PAGE
PFO-PAGE was performed using the method described by Ramjeesingh et al. (1999). Briefly, 4%–20% native gradient gels (Jule Biotechnologies or Bio-Rad) were preelectrophoresed in PFO-running buffer (25 mM Tris base, 192 mM glycine, 0.5% (w/v) PFO (Aldrich), pH of buffer 8.0) (Ramjeesingh et al. 1999) for 45 min at 50 volts. Samples in PFO sample buffer (100 mM Tris base, 8% (w/v) PFO, 20% (v/v) glycerol, 0.005% bromophenol blue) were loaded on preelectrophoresed gels and electrophoresed at room temperature at 60–100 volts until the dye front had run off the gel. The migration of 25 µg of each protein standard (thyroglobulin, catalase, apo-transferrin, urease, and albumin, all from Sigma) was analyzed by Coomassie Brilliant Blue staining. The migration of cldn-4 was analyzed by immunoblotting. Gels were transferred at 4°C in buffer containing 25 mM Tris, 192 mM glycine and 20% (v/v) methanol.
Sucrose velocity gradient centrifugation
Sucrose velocity centrifugation was performed using the method described by Blount and Merlie (Blount and Merlie 1988). 0.2 mL high speed supernatant from solubilzation was loaded on a 5%–20% linear sucrose gradient prepared in 20 mM Hepes and 150 mM NaCl plus 0.1% of the appropriate detergent. Gradients were formed using a Gradient Master (BioComp Instruments). Non-PFO samples were centrifuged in a Beckman SW41 rotor at 39,000 rpm for 22 h at 4°C. PFO samples were centrifuged in a Beckman SW41 rotor at 39,000 rpm for 8 h at 23°C. 0.5 mL fractions were collected by hand from the top. A portion of each fraction was mixed with sample buffer and analyzed by SDS-PAGE.
Purification of cldn-4 in OG and DDM
Sf9 membranes (5 mL per purification, from a 50 mL membrane stock obtained from a 3 L culture) were diluted 1:5 (v/v) with Ni-NTA lysis buffer (50 mM NaH2PO4, 300 mM NaCl, and 10 mM immidazole, pH 8.0) plus 4% detergent (w/v) and solubilized with stirring for 1 h at room temperature. Insoluble material was pelleted by 100,000g centrifugation for 90 min in a Beckmann SW41 rotor. The resultant high-speed supernatant was mixed with washed Ni-NTA Agarose or Superflow resin (QIAGEN) for 1 h at RT. Cldn-4 bound resin was transferred to a column and washed by gravity flow with 5 column volumes of wash buffer (50 mM NaH2PO4, 300 mM NaCl, 1% detergent (w/v) and 20 mM immidazole, pH 8.0). Protein was eluted by gravity flow across eight 1 mL fractions using 50 mM NaH2PO4, 300 mM NaCl, 1% detergent (w/v), and 250 mM immidazole, pH 8.0. An aliquot of each fraction was electrophoresed on a 13% SDS-PAGE gel and analyzed by Coomassie Brilliant Blue staining and immunoblotting. Purified protein was stored at 4°C overnight until use.
Purification of Cldn-4 in PFO
Protein was purified as above except using the following buffers: lysis buffer, pH 8.0 (25 mM NaH2PO4, 200 mM NaCl, 4% (w/v) PFO), wash buffer, pH 7.8 (20 mM NaH2PO4, 200 mM NaCl, 2% (w/v) PFO), and elution buffer, pH 6.5 (20 mM NaH2PO4, 150 mM NaCl, 42 mM PFO). In addition, solubilization and all purification steps were performed at 30°C to maintain PFO in solution. Centrifugation was performed at room temperature. Purified protein was stored at room temperature overnight until use.
Size exclusion chromatography and laser light scattering
Purified claudin-4 was dialyzed overnight against column buffer containing the appropriate detergent. Approximately 100 µg of sample was loaded onto a detergent-equilibrated Superdex 200 gel filtration column, and UV, light scattering, and refractive index were recorded as the sample came off the column. Estimated molecular mass was calculated using the method of Hayashi et al. (1989).
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Acknowledgments |
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The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.
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Y. Yang, Y. Liu, Z. Dong, J. Xu, H. Peng, Z. Liu, and J.-T. Zhang Regulation of Function by Dimerization through the Amino-terminal Membrane-spanning Domain of Human ABCC1/MRP1 J. Biol. Chem., March 23, 2007; 282(12): 8821 - 8830. [Abstract] [Full Text] [PDF]
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J. Hou, A. S. Gomes, D. L. Paul, and D. A. Goodenough Study of Claudin Function by RNA Interference J. Biol. Chem., November 24, 2006; 281(47): 36117 - 36123. [Abstract] [Full Text] [PDF]
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J. Zheng, G.-G. Du, C. T. Anderson, J. P. Keller, A. Orem, P. Dallos, and M. Cheatham Analysis of the Oligomeric Structure of the Motor Protein Prestin J. Biol. Chem., July 21, 2006; 281(29): 19916 - 19924. [Abstract] [Full Text] [PDF]
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 S. P. Chen, B. Zhou, B. C. Willis, A. J. Sandoval, J. M. Liebler, K.-J. Kim, D. K. Ann, E. D. Crandall, and Z. Borok Effects of transdifferentiation and EGF on claudin isoform expression in alveolar epithelial cells J Appl Physiol, January 1, 2005; 98(1): 322 - 328. [Abstract] [Full Text] [PDF]
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J. Xu, Y. Liu, Y. Yang, S. Bates, and J.-T. Zhang Characterization of Oligomeric Human Half-ABC Transporter ATP-binding Cassette G2 J. Biol. Chem., May 7, 2004; 279(19): 19781 - 19789. [Abstract] [Full Text] [PDF]
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 C. B. Coyne, T. M. Gambling, R. C. Boucher, J. L. Carson, and L. G. Johnson Role of claudin interactions in airway tight junctional permeability Am J Physiol Lung Cell Mol Physiol, November 1, 2003; 285(5): L1166 - L1178. [Abstract] [Full Text] [PDF]
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