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Originally published In Press as doi:10.1074/jbc.M502003200 on May 19, 2005

J. Biol. Chem., Vol. 280, Issue 28, 26233-26240, July 15, 2005
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Phosphorylation of Claudin-3 at Threonine 192 by cAMP-dependent Protein Kinase Regulates Tight Junction Barrier Function in Ovarian Cancer Cells*

Theresa D'Souza{ddagger}, Rachana Agarwal{ddagger}, and Patrice J. Morin{ddagger}§

From the {ddagger}Laboratory of Cellular and Molecular Biology, Gerontology Research Center, NIA, National Institutes of Health, Baltimore, Maryland 21224 and §Department of Pathology, The Johns Hopkins Medical Institutions, Baltimore, Maryland 21287

Received for publication, February 22, 2005 , and in revised form, April 28, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Claudins are integral membrane proteins essential in the formation and function of tight junctions (TJs). Disruption of TJs, which have essential roles in cell permeability and polarity, is thought to contribute to epithelial tumorigenesis. Claudin-3 and -4 are frequently overexpressed in ovarian cancer, but the molecular pathways involved in the regulation of these proteins are unclear. Interestingly, several studies have demonstrated a role for phosphorylation in the regulation of TJ complexes, although evidence for claudin phosphorylation is scarce. Here, we showed that claudin-3 and -4 can be phosphorylated in ovarian cancer cells. In vitro phosphorylation assays using glutathione S-transferase fusion constructs demonstrated that the C terminus of claudin-3 is an excellent substrate for cAMP-dependent protein kinase (PKA). Using site-directed mutagenesis, we identified a PKA phosphorylation site at amino acid 192 in the C terminus of claudin-3. Overexpression of the protein containing a T192D mutation, mimicking the phosphorylated state, resulted in a decrease in TJ strength in ovarian cancer cell line OVCA433. Our results suggest that claudin-3 phosphorylation by PKA, a kinase frequently activated in ovarian cancer, may provide a mechanism for the disruption of TJs in this cancer. In addition, our findings may have general implications for the regulation of TJs in normal epithelial cells.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Claudin proteins are a large family of transmembrane proteins essential in the formation and maintenance of tight junctions (TJs).1 Tight junctions in epithelial and endothelial cells provide a selective barrier and establish cellular polarity (1–4). These structures are typically lost in cancer, and this loss may contribute to the invasive and metastatic phenotype of tumor cells (5–8).

Using serial analysis of gene expression, we have previously shown that claudin-3 and -4 are among the most highly up-regulated genes in ovarian cancer (9). The high expression of these claudins in ovarian cancer has been confirmed by our group and others using a variety of approaches such as microarrays, tissue arrays, and reverse transcription-PCR (10–14). Considering that TJs are typically lost in neoplasia, these findings are somewhat surprising, and the exact role of these proteins in ovarian cancer remains unclear. There is, however, some evidence that claudin overexpression may actually lead to increased cell survival and invasion.2 A better understanding of the regulation of claudin proteins in ovarian cancer cells may help clarify their roles in this disease.

Several lines of evidence suggest that TJs are involved in various cell signaling pathways, providing a possible link between cell polarity/cell contact and downstream signaling events (4, 15). Furthermore, the involvement of kinases in the biogenesis and regulation of several tight junction components has been established (16–20). However, only a few studies have clearly demonstrated phosphorylation of claudins. For example, phosphorylation of claudin-1 by mitogen-activated protein kinases (21) and protein kinase C (22), as well as phosphorylation of claudin-5 by cAMP-dependent protein kinase (PKA) (23, 24), has been reported. Also, WNK4 kinase has been shown to phosphorylate claudin-3 and -4 (25).

To address the issue of claudin regulation in ovarian cancer, we have investigated phosphorylation as a possible modulatory mechanism of claudin function. We first show that both claudin-3 and -4 are phosphorylated in ovarian cancer cells, although probably by different kinases. Indeed, whereas claudin-3 is a target of PKA, claudin-4 appears to be phosphorylated by protein kinase C. For this report, we have focused on PKA-mediated phosphorylation of claudin-3. We show that claudin-3 is phosphorylated at threonine 192 in the cytoplasmic C-terminal region. Mimicking the phosphorylated state by mutating this site to an aspartic acid decreases TJ strength in ovarian cancer cells. Our data provide the first evidence for the phosphorylation of claudin-3 and -4 in ovarian cancer cells and suggest that phosphorylation of claudin-3 by PKA, a kinase frequently activated in ovarian cancer, may provide a mechanism for the disruption of TJs in this cancer.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture—Ovarian cell lines BG-1, CAOV3, HEY, IGROV-1, UCI101, 2008, A2780, OVCA420, OVCA429, OVCA432, OVCA433, OVCAR-2, OVCAR-3, OVCAR-4, and OVCAR-5 were cultured in McCoy's 5A growth medium (Invitrogen) supplemented with 10% fetal bovine serum and antibiotics (100 units/ml penicillin and 100 µg/ml streptomycin). HOSE-B, an ovarian surface epithelial cell line immortalized with E6 and E7 (26), was maintained in RPMI 1640 medium supplemented with 10% fetal calf serum and 300 µg/ml Geneticin (Invitrogen). HEK293 cells were maintained in McCoy's growth medium with the supplements described above. To establish stable clones of OVCA433, plasmid vectors were transfected using Lipofectamine plus reagent according to the manufacturer's protocol (Invitrogen). Clones were selected and maintained in McCoy's growth medium containing 500 µg/ml Geneticin.

Antibodies—Rabbit polyclonal claudin-3 and mouse monoclonal claudin-4 were purchased from Zymed Laboratories Inc. Mouse monoclonal PKAc was part of a sampler kit obtained from BD Transduction Laboratories. Rabbit polyclonal PKAc {alpha} was obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Mouse monoclonal {beta}-actin and glyceraldehyde-3-phosphate dehydrogenase were purchased from Abcam (Cambridge, UK). Peroxidase-linked donkey anti-rabbit immunoglobulin and sheep anti-mouse IgG horseradish antibodies were obtained from Amersham Biosciences. Alexa fluor antibodies were purchased from Molecular Probes.

Immunohistochemistry—Expression of claudin-3 and PKAc {alpha} was assessed by immunohistochemical staining of normal ovarian tissue and an ovarian cancer tissue array constructed in our laboratory (10). Antigen-bound primary antibody was detected using streptavidin-biotin immunoperoxidase complex (Ultravision detection system; Labvision). For negative controls, absence of primary antibodies processed in parallel showed no positive staining. Images were acquired by Axiovision 3.1 software on a Zeiss Axiovert S100 microscope under x40 objective lens (Carl Zeiss, Thornwood, NY).

Immunoblotting—Total cell lysates were resolved by SDS-PAGE (Tris-glycine gels; Invitrogen) and transferred to polyvinylidene difluoride membranes (Millipore). Membranes were blocked with 5% nonfat dry milk in 10 mM Tris, pH 7.5, 100 mM NaCl, and 0.1% Tween 20 (v/v) and incubated overnight with primary antibody (claudin-3, 1:200; claudin-4, 1:250, PKAc, 1:1000; {beta}-actin, 1:2000; glyceraldehyde-3-phosphate dehydrogenase, 1:4000). Following the washes in Tris-buffered saline, membranes were probed with horseradish peroxidase-conjugated antibody (anti-mouse or anti-rabbit, 1:10,000). ECL Western blotting detection kit (Amersham Biosciences) was used for visualization of the positive bands.

In Vivo Phosphorylation and Immunoprecipitation—Ovarian cancer cell lines HEY, OVCA432, and OVCA433 were plated at 80–90% confluence in 60-mm dishes. Cells were serum-starved for 16 h and then labeled with [32P]orthophosphate (500 µCi/ml; MP Biomedicals, Irvine, CA) in phosphate-free, serum-free Dulbecco's modified Eagle's medium for 3 h at 37 °C. Cells were stimulated with forskolin (FSK; 30 µM; Sigma) and phorbol 12-myristate 13-acetate (PMA; 50 ng/ml; Sigma) for 20 min at 37 °C. When used, H89 (30 µM; Sigma) was added 30 min prior to the addition of FSK. HEK293 cells were plated at 70–80% confluence in 100-mm dishes and transfected with plasmid pCMV PKAc (Clontech), together with a plasmid encoding either human claudin-3 or the T192A mutant using FuGENE 6 transfection reagents. After 18 h of transfection, cells were labeled with [32P]orthophosphate (500 µCi/ml). Cells were washed in ice-cold phosphate-buffered saline (PBS) and incubated with 700 µl of modified radioimmune precipitation assay buffer (50 mM Tris-Cl, pH 7.4, 150 mM NaCl, 1% Nonidet P-40, 1.5 mM MgCl2, 10% glycerol, 1 mM EDTA, 25 mM NaF, 1 mM Na3VO4, CompleteTM EDTA-free protease inhibitor mixture (Roche Applied Science) for 10 min at 4 °C. The lysate was cleared by centrifugation at 10,000 x g for 20 min. For immunoprecipitation, the supernatant was incubated overnight with 2 µg of primary antibody and 40 µl of protein A-Sepharose (Amersham Biosciences). Immune complexes were washed three times with the lysis buffer without inhibitors, with the final elution in 2x sample buffer, boiled for 5 min, and separated on SDS-PAGE. The gels were either dried, exposed to film for autoradiography, or transferred onto polyvinylidene difluoride membranes and immunoblotted as described above.

Generation of Claudin-3 and -4 WT and Mutant Constructs—The coding sequence of human CLDN3 and CLDN4 was amplified by PCR from cDNA obtained from ovarian cancer cell line OVCAR5 and cloned into pCI-neo mammalian expression vector (Promega, Madison, WI) using the EcoRI and SalI restriction sites. DNA fragments of the GST fusion constructs were generated by PCR using the claudin plasmids described above. GST fusion proteins were constructed with the C-terminal areas of claudin-3 (amino acids 185–220) and claudin-4 (amino acids 181–209). The fragments were designed to contain EcoRI/SalI restriction sites allowing them to be subcloned in-frame with the GST fragment in pGEX-4T-2 vector (Amersham Biosciences). Point mutations of the serine or threonine residues in the consensus PKA site were introduced using the QuikChange site-directed mutagenesis kit (Stratagene). All constructs were confirmed by DNA sequencing.

Expression and Purification of GST-claudin Fusion Proteins— Recombinant constructs of pGEX-CLDN3C, pGEX-CLDN3C-T192A, pGEX-CLDN3C-S199A, and pGEX-CLDN4C were transformed in BL-21 Escherichia coli (Stratagene) for protein expression. Purification was performed according to the manufacturer's protocol with some modifications. Briefly, cultures were grown overnight, and protein expression was initiated with 0.1 mM isopropyl {beta}-D-thiogalactopyranoside at 30 °C for 3 h. Bacterial pellets obtained after centrifugation at 8000 rpm for 10 min at 4 °C were resuspended in cold PBS with CompleteTM EDTA-free protease inhibitor (Roche Applied Science), 1 mM phenylmethylsulfonyl fluoride, and 1 mg/ml lysozyme. After sonication, the lysate was cleared by centrifugation at 10,000 rpm for 20 min. Fusion proteins were purified by affinity chromatograpy using glutathione-Sepharose 4B (Amersham Biosciences). The Sepharose beads were washed three times with PBS, and the fusion protein was eluted with glutathione Elution Buffer (50 mM Tris-Cl/5 mM reduced glutathione).

In Vitro Phosphorylation—Purified GST, GST-fusion proteins, and immunoprecipitated claudin-3 (WT and T192A) proteins were incubated with kinase buffer containing 50 mM Tris-Cl, pH 7.5, 5.1 mM MgCl2, 100 µM ATP, and 1 µCi of [{gamma}-33P]ATP. Catalytic subunit of PKA (5U; Promega) was added to the reaction to a final volume of 20 µl and incubated at 30 °C for 30 min. The reaction was stopped with 5x sample buffer. The proteins was separated on SDS-PAGE (14% Tris-glycine gels), dried, and exposed for autoradiography. Coomassie Blue staining and immunoblot were used to confirm equal loading of proteins.

TER Measurements and Calcium Switch Assay—OVCA433 cells were seeded at a density of 5 x 105 on 12-mm polycarbonate transwell clear membranes (0.4-µm pore size; Costar, Cambridge, MA). TER was measured using a Millicell-ERS V-ohm meter (WPI, New Haven, CT). The values were normalized for the area of the filter and obtained after subtraction of blank values (filter and bath solution). For experiments with drugs, DMSO (vehicle) and FSK (30 µM) were added at time 0, with measurements taken every hour thereafter.

For the calcium switch assay, the calcium in the medium was chelated using EGTA at a final concentration of 4 mM (27). Experiments were performed 7 days after seeding, after the TER had reached an optimal value. Cells were washed with Hanks' balanced salt solution and replaced with normal medium or low calcium medium. TER readings were taken for 4–5 h followed by replacement of low calcium medium with normal medium, and measurements were taken the following day.

Permeability Assay—OVCA433 clones were plated at 5 x 105 cells/transwell and grown for 7 days to confluence and optimal TER development. Fluorescein isothiocyanate-dextran, with an average molecular mass of 4 kDa (Sigma-Aldrich), dissolved in McCoy's media to a concentration of 1 mg/ml was added to the upper chamber. The lower chamber was replaced with fresh media. At the 0, 4, and 24 h time points, 50-µl aliquots were collected from the lower chamber and assayed by luminescence spectrometer using excitation at 485 nm and emission at 530 nm (Cytofluor; PerSeptive Biosystems). The value for vector-transfected OVCA433 was used for normalization and set at 100%.

Immunofluorescence—OVCA433 clones were washed with PBS, followed by fixation with cold methanol:acetone (1:1) for 4 min. OVCA433 cells treated with DMSO (vehicle), FSK (30 µM), or FSK+H89 were fixed in a similar fashion. Fixed cells were washed three times with PBS and blocked in 5% bovine serum albumin, followed by incubation with claudin-3 (1:100) at room temperature for 1 h. After three washes with PBS, the cells were stained with Alexa 594 goat anti-rabbit antibody for 1 h.

Statistical Analysis—Data are expressed as means ± S.E. Statistical analysis was performed by using repeated measures analysis of variance and Tukey post hoc test (28) (using SAS software version 9), with p < 0.05 considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Claudin-3 and -4 Are Phosphorylated in Ovarian Cancer Cell Lines—We first sought to determine whether claudin-3 and -4 could be phosphorylated in ovarian cancer cells. We used in vivo phosphorylation assays in which we stimulated metabolically labeled ovarian cancer cell lines with FSK and PMA, a PKA and a protein kinase C activator, respectively (Fig. 1A). Compared with unstimulated cells, claudin-3 phosphorylation was significantly increased in both OVCA432 and OVCA433 cells. In addition, some of the claudin-3 protein seems to be constitutively phosphorylated in the unstimulated OVCA432 cells. HEY cells, which do not express endogenous claudin-3, were included as a negative control, and the phosphorylated bands observed in these cells were nonspecific, with molecular masses slightly different from those of the claudin bands. Similar results were obtained with claudin-4 (Fig. 1B). Indeed, treatment of cells with the agonists induced phosphorylation of claudin-4 in both OVCA432 and OVCA433 cell lines. The increase in claudin-4 phosphorylation is particularly apparent in OVCA433 cells, in which phosphorylation is essentially absent without activation but highly elevated after FSK/PMA treatment. HEY cells were again used as negative control because they lack endogenous claudin-4 protein as well.



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  		FIG. 1.
Claudin-3 and -4 are differentially phosphorylated in ovarian cancer cell lines. A, OVCA432, OVCA433, and HEY cells metabolically labeled with [32P]orthophosphate were treated with FSK (30 µM) and PMA (50 ng/ml) or vehicle (DMSO) and then immunoprecipitated with either claudin-3 or FLAG antibody. Arrow indicates the position of phosphorylated claudin-3 confirmed by immunoblot. B, OVCA432, OVCA433, and HEY cells were stimulated with FSK and PMA and immunoprecipitated with either claudin-4 or FLAG antibody. Phosphorylated claudin-3 and -4 were detected in OVCA432 and OVCA433 cell lines upon induction by FSK and PMA but were not detected in HEY, a cell line deficient in both claudin-3 and -4. The accompanying immunoblot shows equal amount of protein immunoprecipitated by the anti-claudin-4 antibody. C, stimulation of OVCA433 cells by FSK and PMA individually shows that claudin-3 is strongly phosphorylated following FSK treatment and that claudin-4 is phosphorylated following PMA treatment. D, the phosphorylation level of claudin-3 brought about by FSK is reduced in the presence of PKA inhibitor H89 (30 µM).

 



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  		FIG. 2.
Protein expression of PKA and claudin-3 in ovarian cancer cell lines and ovarian cancer tissues. A, expression of PKAc {alpha} and claudin-3 in normal HOSE-B and various ovarian cancer cell lines was assessed by immunoblotting. High levels of PKAc {alpha} were found in most of the cell lines examined, with the exception of HEY, which exhibited little or no PKAc {alpha} expression. Approximately 68% of the ovarian cancer cell lines expressed both claudin-3 and PKAc. B, a and d, PKAc {alpha} is present, but claudin-3 is absent in normal ovarian epithelium (arrow). B, b, c, e, and f, prominent presence of PKAc {alpha} and claudin-3 is shown in the representative ovarian serous carcinoma tumors by immunohistochemical staining. The insets of c and f show the marked areas of the serous carcinoma tissue at higher magnification. Scale bar = 10 µm.

 
To determine which of the agonists increased the phosphorylation of claudin-3 and -4 proteins, an in vivo phosphorylation assay was performed in OVCA433 cells in the presence of either FSK or PMA. Fig. 1C shows that claudin-3 was strongly phosphorylated following FSK treatment, whereas PMA increased the phosphorylation of claudin-4 in OVCA433 cells. Similar results were obtained in OVCA432 cells (data not shown). These results indicate that, in these cells, claudin-3 is phosphorylated by PKA, whereas claudin-4 is phosphorylated by protein kinase C. Moreover, FSK-dependent phosphorylation of claudin-3 was inhibited by H89, a PKA inhibitor, bringing it back to basal phosphorylation level (Fig. 1D). This experiment further strengthens our conclusion that claudin-3 is phosphorylated in vivo by PKA in these cells.

Expression of PKA Catalytic and Claudin-3 Proteins in Ovarian Cancer Cell Lines and Cancer Tissues—Because of the findings above suggesting that claudin-3 is a likely target for PKA in ovarian cancer cells, we decided to examine endogenous protein levels of PKAc {alpha} in a panel of ovarian cancer cells. PKAc {alpha} was prominently present in all cell lines examined except HEY, which also fails to express claudin-3 (Fig. 2A). Combined expression of the catalytic subunit with the various regulatory subunits can generate various physiological consequences in different cells (29). Most of the cell lines in Fig. 2A also expressed regulatory subunits (data not shown), indicating the existence of functional PKA complexes in these cells. This is in agreement with previous reports showing a frequent activation of PKA in ovarian cancer (30, 31). Claudin-3 expression was absent in non-malignant HOSE-B cells but found in most of the ovarian cancer lines. Overall, 11 of 16 cell lines expressed both claudin-3 and PKAc {alpha}. This is consistent with our earlier finding demonstrating expression of claudin-3 protein in the vast majority of ovarian cancers (10). Staining of normal ovarian tissue with PKAc {alpha} and claudin-3 indicated the presence of PKAc {alpha} in the epithelium and stroma of the tissue, but claudin-3 was clearly absent (Fig. 2B, a and d), similar to what we observed for the expression of these proteins in the HOSE-B non-maligant ovarian epithelial cells (Fig. 2A). Staining of our ovarian cancer tissue array with PKAc {alpha} antibody showed that this protein was present in all the ovarian cancer tissues and that the areas of PKAc staining overlapped with areas of claudin-3 staining (Fig. 2B, b, c, e, and f). These proteins were found both at the membrane and cytoplasm. The insets of Fig. 2B, c and f, which are shown at higher magnification, verify the localization of PKAc {alpha} and claudin-3. These results are consistent with our hypothesis that claudin-3 may be a target for PKA phosphorylation in ovarian cancer cells.



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  		FIG. 3.
Claudin-3 is phosphorylated by cAMP-dependent protein kinase in vitro. A, diagrammatic representation of the GST-claudin C terminus. The amino acids that were mutated (underlined) are shown within the consensus sequence of the C terminus of claudin-3. Purified recombinant C termini of claudin-3, claudin-4 (B) and mutants of claudin-3, T192A, and S199A (C) were incubated with PKAc in a kinase assay. Fusion proteins were resolved by SDS-PAGE, and the phosphorylated proteins were visualized by autoradiography. The autoradiograms shown are representative examples of four separate experiments. D, claudin-3 WT and T192A immunoprecipitates, from transfected HEY and HEK293 cells, were phosphorylated by PKAc in an in vitro kinase assay. The band intensities in the autoradiogram are shown as arbitrary levels of 33P. The gels were transferred and immunoblotted with anti-claudin-3 antibody to show the presence of the protein. E, HEK293 cells deficient in claudin-3 were transfected with claudin-3 WT or its mutant, T192A, alone or with PKAc, and metabolically labeled with [32P]orthophosphate. Claudin-3 was immunoprecipitated from cell lysates, resolved by SDS-PAGE, and detected by autoradiography. Protein presence is shown by the accompanying immunoblot. F, phosphorylated claudin-3 WT was present in the Nonidet P-40-soluble fraction, but not in the Nonidet P-40-insoluble fraction, even though the protein levels are approximately equal. The phosphorylated levels were determined from the autoradiogram intensities and are shown in the accompanying bar diagrams as arbitrary levels of 32P.

 
Recombinant Claudin-3 Protein Is Phosphorylated in Vitro by PKA—We next wished to determine the exact PKA phosphorylation site(s) in claudin-3. We performed an in vitro phosphorylation assay with GST fusion proteins containing the C terminus of claudin-3 and -4 (Fig. 3A). PKA strongly phosphorylated the claudin-3 C-terminal region, whereas claudin-4 was very weakly phosphorylated compared with claudin-3 (Fig. 3B). The GST protein without claudin sequences was not phosphorylated in these assays. These in vitro findings are consistent with the in vivo observations, showing that claudin-3 is strongly phosphorylated by a PKA activator compared with claudin-4 (Fig. 1). To determine which of the serine/threonine residues in the C-terminal region of claudin-3 is phosphorylated, we performed site-directed mutagenesis on putative sites that met the minimal consensus sequence for PKA (R/KXT/S) (Fig. 3A). Fig. 3C shows that a single threonine to alanine substitution at position 192 of the C-terminal region of claudin-3 abolished the phosphorylation signal. Mutation at another putative phosphorylation residue (Ser-199), although leading to a slight reduction in signal, did not prevent strong phosphorylation of the claudin-3 C terminus. In addition, various GST fusions with other regions of claudin-3 (N-terminal, loop, and transmembrane regions) were not phosphorylated in these in vitro assays (data not shown).

Claudin-3 Is Phosphorylated in Vitro and in Vivo by PKA at Thr-192—We next asked whether full-length claudin-3 could be phosphorylated in vitro and in vivo at Thr-192. Because a theoretical tryptic digest analysis of claudin-3 showed that the fragment containing Thr-192 would be out of the detection range for mass spectrometry, we resorted to genetic approaches to answer this question. Full-length claudin-3 WT and mutant (T192A) were overexpressed in two cell lines, HEY and HEK293, both of which were devoid of endogenous claudin-3. Immunoprecipitated proteins were incubated with PKAc along with radiolabeled ATP for a kinase assay. Immunoprecipitates of claudin-3 WT from both HEY and HEK293 exhibited phosphorylated claudin-3, whereas no phosphorylation was observed in cells expressing the mutant T192A (Fig. 3D). In addition, we co-expressed claudin-3 WT or its mutant T192A with PKAc in HEK293 cells. Fig. 3E shows that, in the presence of the active PKAc, claudin-3 WT was phosphorylated, whereas the mutant T192A was not. Interestingly, the phosphorylated claudin-3 was more prominent in the Nonidet P-40-soluble fraction than in the insoluble fraction (Fig. 3F).



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  		FIG. 4.
Overexpression of claudin-3 and its mutants affects barrier function. A, TER measurements of OVCA433 stable clones. TER values (expressed as means ± S.E.; n = 5 in triplicate) were taken for several days after seeding stable clones of OVCA433 overexpressing claudin-3 WT ({blacksquare}) and its mutants T192A ({triangleup}) and T192D ({circ}) and vector alone ({diamondsuit}) in transwells. TER of claudin-3 T192D significantly decreased compared with control, WT, and T192A (*, p < 0.0001). The immunoblot (inset) shows the overexpression of claudin-3 WT and mutants over endogenous claudin-3 in the stable clones of OVCA433. The same membrane was stripped and probed with glyceraldehyde-3-phosphate dehydrogenase (GAPDH) for loading. B, effect of claudin-3 WT and mutants on the permeability of OVCA433 cell monolayers. Fluorescein isothiocyanate-dextran (4 kDa, 1 mg/ml) was added to upper chamber, and the paracellular tracer was measured from the lower chamber at regular time intervals after incubation. Claudin-3 mutant T192D significantly increased the permeability (*, p < 0.005) compared with the vector alone. C, effect of calcium removal on Claudin-3 T192D. Monolayers of OVCA433 expressing claudin-3 WT and mutants were subjected to Ca2+ switch 7 days after seeding (filled arrow). TER values (n = 5; means ± S.E.) show that barrier function recovered to the original levels after replacement of calcium (normal medium, open arrow). Cells expressing the T192D mutant failed to reach high levels of TER. D, confluent OVCA433 cells growing on transwell membranes for several days were treated with FSK (30 µM), and TER was measured over a 5-h period. FSK treatment decreased TER in a time-dependent manner. Data are expressed as means ± S.E. (n = 3 in triplicates), with significant changes observed after 2 h (p < 0.05; for 3, 4, and 5 h, p < 0.001).

 
Claudin-3 Phosphorylation Affects Barrier Function—To evaluate the functional role of PKA phosphorylation in the barrier function of TJs, we overexpressed claudin-3 WT, mutant T192A (non-phosphorylatable), and mutant T192D (mimicking the phosphorylated state of the residue) in ovarian cancer cells. For this analysis, we chose the OVCA433 cell line because, among the ovarian cancer cell lines tested, these cells had the highest TJ function as evaluated by transepithelial electrical resistance (TER) measurements. Non-transfected cells, as well as cells expressing claudin-3 WT or the T192A mutant, developed relatively high TER over a period of ~7 days (Fig. 4A). However, expression of the T192D mutant, mimicking the phosphorylated state, led to a significant inhibition of TER with at least a 6- or 4-fold reduction compared with the values of vector or claudin-3 WT, respectively (F3,16 = 14, p < 0.0001) at day 7, suggesting that the TJs could not form properly. The data shown here are from one set of OVCA433 clones, but comparable results were obtained with another series of clones, as well as with pools of transfectants (data not shown). We carried out similar experiments with Madin-Darby canine kidney I cells, which have high TER and are commonly used in TJ studies. Similar to what we observed for OVCA433 cells, expression of the mutant T192D in Madin-Darby canine kidney I cells decreased the TER 4- or 2-fold compared with vector or claudin-3 WT, respectively (data not shown).

We also assessed permeability changes of OVCA433 stable clones using the paracellular tracer fluorescein isothiocyanate-dextran (4 kDa) across the established monolayers. Compared with vector-transfected cells, cells expressing claudin-3 T192D mutant had significantly increased paracellular flux of fluorescein isothiocyanate-dextran. Indeed, the transported dextran was increased by 30% after 4 h of incubation and by 150% after 24 h of incubation (Fig. 4B). The other two stable transformants expressing claudin-3 WT and mutant T192A exhibited permeability properties similar to the vector-transfected control.



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  		FIG. 5.
Localization of claudin-3. A, overexpression of claudin-3 WT and mutants T192A and T192D in OVCA433 cells was detected by immunostaining. T192D overexpression leads to a more diffused staining at the intercellular junctions (arrows), indicative of a disrupted state. B, the confluent cells grown for 5 days were subjected to FSK, FSK+H89 (PKA inhibitor), or DMSO (vehicle) treatment for 2 h followed by immunofluorescence to show the localization of claudin-3. FSK treatment led to diffuse staining of claudin-3 at the intercellular junctions (arrows).

 
Recovery of Barrier Function is Normal in Claudin-3-overexpressing Cells after Ca2+ Switch—Removal of extracellular calcium results in a PKA-dependent disruption of TJs (32). PKA has also been shown to affect the resealing of TJs following the replenishment of calcium, possibly via vasodilator-stimulated phosphoprotein, a direct substrate for PKA, implicated in TJ regulation (33). We therefore sought to investigate whether breaking down TJs by depletion of extracellular calcium would be affected by overexpression of claudin-3 and its phosphorylation mutants. The OVCA433 cells expressing claudin-3 WT or mutants were washed with Hanks' balanced salt solution before adding normal medium (control) or EGTA-containing medium (calcium chelation). As expected, the TER values plummeted in all the clones after calcium removal (Fig. 4C). Calcium was then replenished at the time indicated by the open arrow in Fig. 4C. The following day, all the previous TER values were regained, following the same pattern as before depletion of calcium. After TJ destruction in confluent cells, the TJ assembly could take place in cells expressing claudin-3 WT or the T192A mutant, but not in the T192D-expressing cells, again suggesting that phosphorylation of the Thr-192 residue led to a decrease in the ability of the cells to form TJs. In addition, consistent with these results, FSK stimulation of OVCA433 cells, which leads to claudin-3 phosphorylation (Fig. 1C), resulted in a decrease in TER in a time-dependent manner after 2 h of treatment (p < 0.05) (Fig. 4D).

Localization of Phosphorylated Claudin-3—Experiments described thus far have shown that claudin-3 can be phosphorylated by PKA and that a mutated form mimicking the phosphorylated state can decrease TJ function when overexpressed. To better understand the mechanisms leading to this weakening of TJs, we determined whether phosphorylation at Thr-192 might affect subcellular localization of claudin-3. Both claudin-3 WT and mutant T192A were found tightly localized at the intercellular junctions. However, whereas claudin-3 T192D was still found at the surface, the levels appeared to be decreased, and the staining appeared to be more diffused (Fig. 5A). These results show that phosphorylation may cause a partial redistribution of claudin-3 from the TJs to other membrane or cytoplasmic areas and may provide a mechanism for the decreased TJ strength observed above. In addition, the localization pattern obtained with FSK treatment was consistent with this hypothesis. Indeed, without treatment, claudin-3 was tightly localized to the surface of the cells (Fig. 5B). Upon exposure to FSK, however, claudin-3 appeared disrupted and was more diffuse at the intercellular junctions. Pretreatment of the cells with the PKA inhibitor H89, followed by addition of FSK in the presence of H89, prevented the effects of FSK.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
A growing body of work has confirmed the overexpression of claudin-3 and -4 in ovarian cancer (9–14), but the role and regulation of claudin proteins in this cancer have not been elucidated. As a first step in clarifying the regulation of claudins in ovarian cancer, we have focused on the effects of phosphorylation of these proteins. Using PKA and protein kinase C activators, we show that, in two different cancer cell lines, claudin-3 is target of PKA, whereas claudin-4 is phosphorylated by protein kinase C. The significance of the differential regulation of these two claudins is unclear at this point, but because the effects of PKA on TJs have remained elusive, we have focused on the phosphorylation of claudin-3 by PKA. We clearly demonstrate that treatment with PKA activator FSK of both OVCA432 and OVCA433 leads to increased claudin-3 phosphorylation. However, the basis for the difference observed in the endogenous phosphorylation level of claudin-3 between these cell lines (claudin is constitutively phosphorylated in OVCA432 compared with OVCA433) is unknown. It may be brought about by differences in the endogenous activity of PKA in these cell lines or may be due to other kinases. Either way, the fact remains that claudin-3 phosphorylation is significantly induced in these cells and that H89, a PKA inhibitor, abolishes this effect. We next identified Thr-192 as the main PKA phosphorylation site in claudin-3. This is demonstrated using in vitro phosphorylation assays of wild type and mutant proteins. We also extend these findings in vivo through transfection assays involving either wild type or mutant claudin-3 in cells lacking endogenous claudin-3.

To elucidate the possible phenotypic effects of claudin-3 phosphorylation in ovarian cancer cells, we have assessed the most widely studied function of claudin proteins, the ability to mediate TJ formation. OVCA433 was chosen for these functional studies because it gave measurable TER, a widely used index for the measurement of TJ strength. Interestingly, overexpression of a mutant claudin-3 (T192D) mimicking the phosphorylated form had a dominant negative effect compared with the WT and the non-phosphorylatable forms of the protein, suggesting that phosphorylation at threonine 192 may interfere with TJ formation. In addition, we showed that expression of the mutant T192D increased the permeability of OVCA433 cells using an immunofluorescent tracer. In light of both the TER and permeability assays, it appears that the TJ integrity may be hampered in the presence of overexpressed claudin-3 T192D. The exact mechanisms leading to a decrease in TJ function by phosphorylated claudin-3 are unclear, but our immunofluorescence data provide some insights (Fig. 5B). Indeed, compared with the WT claudin-3, expression of the T192D mutant appeared to lead to much more diffused staining at the intercellular junctions, suggesting that the phosphorylation may disrupt the ability of claudin-3 to participate in TJ formation. Moreover, stimulation of the cells with FSK caused a similar diffused pattern of staining for claudin-3 (Figs. 4D and 5B). In addition, we found that whereas total claudin-3 was partitioned in both Nonidet P-40-soluble and -insoluble fractions, the phosphorylated form was found mostly in the Nonidet P-40-soluble fraction (Fig. 3F). Although it is formally possible that phosphorylation affects the detergent solubility of the claudin protein, these results, in combination with our immunofluorescence study (Fig. 5), strongly suggest a shift in subcellular localization and molecular interactions following phosphorylation. This, in turn, likely affects the barrier function or signaling of claudin proteins. These results are in contrast with recent findings suggesting that cAMP signaling does not affect the detergent solubility of claudin-5 (24). However, it is certainly possible that different phosphorylation events may have different effects on claudin proteins. In addition, it is important to note that interaction of claudin-3 with other intercellular junctional and associated cytosolic proteins and their modulation by PKA may also affect TJs. Zonula Occludens-2, a TJ-associated protein, has been shown to be phosphorylated by PKA, affecting its function at the junctional complex (35). Additionally, recent work demonstrated that PKA negatively regulates adherens junction integrity (36). It will be interesting to evaluate the dynamic relationship between claudin and other TJ proteins and the effect of claudin phosphorylation on these interactions.

In this study, we show that, in ovarian cancer cells, claudin-3 and -4 can be phosphorylated by PKA and protein kinase C, respectively. This represents, to our knowledge, the first report of claudin phosphorylation by these kinases in any cell model. The consequences of the differential modulation brought about by these kinases on these claudins remain to be determined but may contribute to ovarian tumorigenesis. Recently, it was shown that claudin-3 and -4 are substrates for mutant WNK4 kinase (25), which is linked to pseudohypoaldosteronism type II, a disease associated with an increase in paracellular chloride permeability. Whereas the exact molecular mechanisms are still unclear, it was postulated that hyperphosphorylation of claudin could be causally related to the increase in chloride permeability. These data are consistent with our finding that claudin-3 can be phosphorylated in vivo and the observation that this phosphorylation interferes with TJ function. Studies have implicated protein kinase C in the regulation of TJs through phorbol ester stimulation (17, 34), but PKA-dependent regulation of TJs has been poorly documented and is more controversial (20). PKA activation has been suggested to have a role in TJ disruption (32), and claudin-3 phosphorylation may represent one of the targets to achieve this physiological outcome. Interestingly, PKA is often activated in ovarian cancer (30, 31). In addition, our unpublished work2 suggests that claudin overexpression in ovarian cancer cells may contribute to an increase in invasion and survival of these cells, mediated at least in part through the breakdown of TJs. PKA, through its role in the phosphorylation of claudin-3, may therefore contribute to the decrease in polarity and increase the invasive properties of ovarian cancer cells. In this context, this PKA-claudin-3 pathway may represent an interesting target for cancer therapy, and our finding would provide yet an additional incentive to develop PKA inhibitors for antitumor therapy (37). Our current finding of claudin phosphorylation provides a new mechanism for the regulation of claudin-3 in ovarian cancer. In addition, we are currently investigating the mechanisms of transcriptional regulation of this protein in ovarian cancer. It remains to be seen whether the regulatory mechanism described in this report is a general property of epithelial cells or is restricted to the pathophysiological state of ovarian cancer.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Laboratory of Cellular and Molecular Biology, Gerontology Research Center, NIA, National Institutes of Health, 5600 Nathan Shock Dr., Baltimore, MD 21224. Tel.: 410-558-8506; Fax: 410-558-8386; E-mail: morinp{at}grc.nia.nih.gov.

1 The abbreviations used are: TJ, tight junction; FSK, forskolin; GST, glutathione S-transferase; PKA, cAMP-dependent protein kinase; PMA, phorbol 12-myristate 13-acetate; TER, transepithelial electrical resistance; PBS, phosphate-buffered saline; WT, wild type; DMSO, dimethyl sulfoxide; PKAc, PKA catalytic subunit. Back

2 R. Agarwal, T. D'Souza, and P. J. Morin, unpublished observations. Back


   ACKNOWLEDGMENTS
 
We thank Cheryl Sherman-Baust and Drs. Hiroshi Honda, Jianghong Li, Roman Wernyj, and Ashani Weeraratna for helpful comments on the manuscript. We thank Dr. Jeff Leips for the assistance in statistical analysis and comments on the manuscript. We thank Cheryl Sherman-Baust for immunohistochemistry. The A2780 cell line was obtained from Dr. Vilhelm Bohr (Baltimore, MD); BG-1 was obtained from Dr. Carl Barrett (Durham, NC); UCI 101 was obtained from Dr. Michael Birrer (Rockville, MD); OVCA420, OVCA429, OVCA432, OVCA433, and HEY were obtained from Dr. Robert Bast (Houston, TX); and OVCAR2, OVCAR4, and OVCAR5 were obtained from Dr. Thomas C. Hamilton (Philadelphia, PA).



   REFERENCES
TOP
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 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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