Ovarian Tumor Cell Expression of Claudin-4 Reduces Apoptotic Response to Paclitaxel Free

Because of the lack of adequate early detection screening, the majority (>85%) of the patients with epithelial ovarian cancer (EOC) are diagnosed at advanced stages of disease. Delayed detection means that ovarian tumor cells have disseminated beyond their site of origin (fallopian tube or ovary) into the peritoneal cavity. The standard of care for these patients is surgical debulking of the tumor, followed by adjuvant platinum- and taxane-based chemotherapies (cisplatin/carboplatin and paclitaxel/docetaxel; ref. 1). One third of patients with ovarian cancer have tumors that do not respond to initial chemotherapy and of the patients that do respond, approximately 75% of these patients will have disease recurrence (2). Patients with recurrent EOC succumb to the disease following the development of chemoresistance (3).

There are several proposed mechanisms responsible for the reduced therapeutic response that include both de novo pathways that inherently make tumor cells resistant and the acquired pathways that are upregulated in response to repeated chemotherapeutic insult. These pathways include, but are not limited to, drug transporters (i.e. P-glycoprotein) that either inhibit drug uptake or enhance drug efflux (4–6), epigenetic changes (i.e. MLH1 and Tap73 methylation, miR-214, and miR-376c upregulation) that prevent expression of antitumorigenic genes (7–10), expression or mutations in proapoptotic proteins [i.e., p53, Bcl-2 family proteins, PTEN (11–14)], and cytoskeletal organization that prevents binding of microtubule-targeting agents (15). Despite the increased understanding of chemoresistance mechanisms, it is unlikely that a single pathway can be targeted to restore tumor sensitivity to all cytotoxic drugs. However, a deeper understanding of patients’ genetic and proteomic characteristics, which can predict response and inform optimal therapeutic strategies, is a significant step forward.

Claudin-4 is a member of a large family of transmembrane proteins, of which there are 27 different subtypes (16). Claudins have been most studied for their canonical role in tight junctions, where they mediate the specific paracellular barrier properties of an epithelium. Although claudin-4 has been shown to alter the permeability of epithelial monolayers in cell culture (17–19), it is rarely localized to tight junctions in human and mouse tissue and is generally found along basolateral membranes and throughout the cytosol (20–22). The findings that claudin-4 knockout (KO) mice are phenotypically normal with only a subclinical increase in permeability to small molecules in lung epithelium, but have a reduced capacity for wound healing, further suggest a noncanonical role for claudin-4 (23). Furthermore, it has been well documented that claudin-4 is significantly upregulated in multiple epithelia-derived cancer cells that lack traditional tight junction structures. High claudin-4 expression has, in fact, been associated with a phenotypically aggressive ovarian cancer cell [chemoresistant, highly mobile, and stem-like (24–26)]. The mechanism by which claudin-4 exerts these effects is currently unknown.

To investigate the noncanonical role of claudin-4 in both normal and tumor cell biology, we previously designed a small claudin mimic peptide (CMP) that interferes with the DFYNP sequence in the second extracellular loop of claudin-4 that is thought to be important in claudin–claudin interactions (27). We have recently shown that disrupting claudin-4 activity with CMP increased tumor cell response to the potent apoptotic inducer, staurosporine, and inhibited cell migration (28). In addition, treatment of mice bearing EOC cell line–derived xenograft tumors with CMP significantly reduced the tumor burden. These observations suggest that claudin-4 expression in ovarian cancer cells plays a functional role in tumor progression.

To determine whether claudin-4 plays a role in the apoptotic response of ovarian tumors to standard chemotherapeutics, we examined whether the presence of the protein altered the apoptotic response of EOC cell lines to cisplatin and paclitaxel in the presence and absence of CMP. We found that claudin-4 expression led to reduced tumor cell sensitivity to paclitaxel-induced apoptosis. Tumor cell sensitivity to paclitaxel was restored by silencing claudin-4 expression or co-treatment with CMP. We also show claudin-4 involvement in cell-cycle progression and an interaction of claudin-4 with microtubules, providing a possible mechanism by which claudin-4 drives resistance to paclitaxel.

Human-derived OVCAR3, OVCAR4, OVCAR5, OVCAR8, PEO4, OV429, and DOV13 ovarian tumor cells were obtained from the laboratory of Monique A. Spillman (University of Colorado Denver, Aurora, CO) and cultured in RPMI1640 medium (Gibco, Thermo Fisher Scientific) plus 10% heat-inactivated FBS (Access Biologicals) and 1% penicillin/streptomycin (Gibco, Thermo Fisher Scientific) at 37°C and 5% CO₂. Cells were trypsinized (0.25% trypsin, EDTA, Mediatech) and plated 1:3 every 3–4 days. All cell lines were authenticated at the beginning of this study by short tandem repeat profiling, as described previously (29). Cells were passaged approximately 10 times before thawing out a fresh vial of authenticated cells. Authentication was then repeated at the end of the study. Cells were plated at 1 × 10⁴ cells/well into type I collagen-coated 8-well chamber slides (Lab-Tek, NUNC) for treatment and analysis. Upon 80%–90% confluence, cells were treated with either 400 μmol/L control peptide [(NH2-GDGYNPG-OH, D-amino acid conformation, University of Colorado Protein and Peptide Chemistry Core (27)], 400 μmol/L claudin mimic peptide (CMP, NH2-GDFYNPG-OH, D-amino acid conformation, University of Colorado Protein and Peptide Core), 10 nmol/L paclitaxel (Sigma-Aldrich), and/or 10 μmol/L cisplatin (Sigma-Aldrich) for 24 hours. To examine the apoptotic mechanism, cells were also treated with 50 μmol/L etoposide (Sigma-Aldrich).

De-identified patient tumor samples were collected from the University of Colorado Gynecologic Tissue and Fluid Bank under an Institutional Review Board–approved protocol (COMIRB 05-1081). Each tumor sample was flash frozen in liquid nitrogen and stored at −80°C until processed for protein. Frozen tissue was placed in lysis buffer [30 mmol/L Tris HCl pH7.4, 150 mmol/L NaCl, 1% Triton X-100, 10% glycerol, 2 mmol/L EDTA, 0.57 mmol/L phenylmethylsulfonylfluoride (PMSF), 1× cOmplete Protease Inhibitor Cocktail] and homogenized using a Polytron tissue homogenizer (Brinkmann Instruments, Thermo Fisher Scientific). Lysates were then spun at 13,000 rpm for 10 minutes and the supernatant collected for protein analysis.

To determine levels of claudin-4 protein expression, tumor cells were scraped from culture plates in the presence of lysis buffer (30 mmol/L Tris HCl pH 7.4, 150 mmol/L NaCl, 1% Triton X-100, 10% glycerol, 2 mmol/L EDTA, 0.57 mmol/L PMSF, 1× cOmplete Protease Inhibitor Cocktail), placed on a shaker for 10 minutes and spun at 13,000 rpm for 10 minutes. Supernatant was collected and 20 μg of total protein was denatured, resolved on 10% SDS-PAGE, and transferred to a polyvinylidene difluoride membrane (Bio-Rad). Membranes were blocked with 5% nonfat dry milk in Tris-buffered saline with 0.1% Tween-20 (TBST) for 1 hour at room temperature before treatment with either rabbit anti-human claudin-4 (1:500, Invitrogen), mouse anti-human claudin-4 (1:500; Invitrogen), or rabbit antihuman GAPDH (1:10,000; Sigma) overnight at 4°C. Membranes were then washed four times with TBST for 15 minutes before treatment with horseradish peroxidase (HRP)-conjugated goat anti-rabbit (1:10,000; GE Healthcare) or goat anti-mouse (1:10,000; Jackson ImmunoResearch Laboratories) antibodies for 1 hour at room temperature. Membranes were washed with TBST as described above and then visualized using an ECL Prime Western Blotting Detection Reagent (GE Healthcare) and X-ray film (CL-XPosure Film, Thermo Fisher Scientific).

Apoptosis was measured by determining the number of cells exhibiting activated caspase-3. After treatment, cells were washed with PBS (Gibco, Thermo Fisher Scientific) and fixed with 10% phosphate-buffered formalin (Thermo Fisher Scientific) at room temperature for 15 minutes. Cells were washed twice with PBS before cell membrane permeablization with 0.5% Triton X-100 (IBI Scientific) for 5 minutes and washed again with PBS. Cells were treated with blocking buffer (2% BSA; Sigma-Aldrich) for 1 hour before application of primary antibody directed to cleaved caspase-3 (1:400; rabbit anti-human cleaved caspase-3, Cell Signaling Technology) overnight at 4°C. Cells were then washed with PBS five times before application of secondary antibody conjugated to CY3 (1:100; donkey anti-rabbit, Jackson ImmunoResearch Laboratories) and 5 μg/mL 4′,6-diamidino-2-phenylindole (DAPI; Sigma) for 45 minutes at room temperature followed by washing with PBS five times. After removal of PBS, o-phenylenediamine dihydrochloride (20 mg/mL; OPDA) in 1 mol/L Tris (pH 8.5) was added to the slides to preserve fluorescence and coverslipped. The Olympus FV1000/RICS confocal microscope (University of Colorado AMC Light Microscopy Core) was used to image fluorescence. The percent of cells positive for active caspase was determined using SlideBook software (Intelligent Imaging Innovations, Inc.).

OVCAR3 and PEO4 cells were plated 3.2 × 10⁴ in a 96-well plate and incubated at 37°C for 24 hours. When cells reached 70% confluence, 10 μL of claudin-4 short hairpin RNA (shRNA; TRC#: TRCN0000116627, TRCN0000116628, or TRCN0000116631) or control shRNA (SHC001, pLK0.1-puro Empty Vector) lentiviral suspension (Sigma-Aldrich MISSION shRNA, University of Colorado Functional Genomics Facility) was added to the cells and incubated overnight at 37°C. Fresh medium was added to remove lentivirus and cells were allowed to recover for 24 hours before being treated with 0.5 μg/mL puromycin for selection and expansion of transduced cells. Western blot analysis (as described above) confirmed loss of claudin-4 expression.

GFP-tagged claudin-4 and a GFP-only control (pLenti-C-mGFP vector; OriGene) were transduced into OVCAR8 and OVCAR4 cells via lentiviral particles in the presence of 10 μg/mL hexadimethrine bromide. GFP-positive cells were flow-sorted and expanded.

To examine the number of cells in mitosis, we examined the images of DAPI fluorescence in OVCAR3 cells cultured in 8-well chamber slides. Cells that had clearly condensed chromatin that aligned along a single plane were counted as mitotic cells and calculated as the percentage of total number of nuclei per field of view (at least 12 fields of view were analyzed for each condition). In addition, a Mitotic Assay Kit (Active Motif) was used as per manufacturer's instructions. Briefly, cells were cultured in 96-well plates, fixed, and treated with a phospho-histone H3 (Ser 28) monoclonal primary antibody followed by an HRP-conjugated secondary antibody. Absorbance at 450 nm was read after 20-minute incubation with the Developing Solution. Crystal violet was used to normalize for total cell number.

A total of 10,000 cells were plated in each well of 12-well culture plates. Cells were trypsinized (0.25% trypsin, EDTA) and counted by a Countess Automated Cell Counter (Invitrogen) after 24, 48, and 72 hours in culture.

Cells were plated in 100-mm culture dishes at 5 × 10⁵ cells per dish. After 24 hours in culture, media were changed to serum-free RPMI1640 for 48 hours. Cells were then exposed to media containing serum and harvested 6 hours later. Cells were placed in Krishan stain (0.224 g sodium citrate 2H₂O, 9.22 mg propidium iodide, 2.0 mL 1% NP40 in H₂O, 2.0 mL 1 mg/mL RNase in 200 mL H₂O) overnight at 4°C before being sent to the University of Colorado Flow Cytometry Core Facility, Aurora, CO). Cells were analyzed by flow cytometry (Beckman Coulter Gallios, University of Colorado Flow Cytometry Core Facility, Aurora, CO), measuring fluorescence emission at >575 nm (FL3). FlowJo software was used to determine number of cells in G₂–M phase.

Claudin-4 interaction with tubulin was examined using the DuoLink Proximity Ligation Assay Kit (Sigma), following the manufacturer's protocol. Briefly, cells were plated in 8-well chamber slides and cultured to 70% confluence before being fixed, permeablized, and blocked as described above for capase-3 immunofluorescence. Cells were then treated with antibodies directed to claudin-4 (mouse anti-human claudin-4; 1:200; Invitrogen) and α-tubulin (rabbit anti-human α-tubulin; 1:100; Abcam) or β-tubulin (rabbit anti-human β-tubulin; 1:100; Abcam) overnight at 4°C. After washing with PBS, cells were treated with PLA probes (1:5; anti-mouse PLUS, anti-rabbit MINUS) for 1 hour at 37°C in a preheated humidity chamber. After washing, Ligation Solution was added for 30 minutes at 37°C followed by washing and addition of amplification polymerase solution for 100 minutes at 37°C in a humidity chamber. Slides were washed and dried before adding mounting medium and placing coverslip. Fluorescence was imaged using an Olympus FV1000/RICS confocal microscope (University of Colorado AMC Light Microscopy Core).

Claudin-4 (and interacting proteins) was pulled from cell lysates with an antibody directed to claudin-4 (mouse anti-human claudin-4, Invitrogen) bound to Protein A/G Dynabeads Magnetic beads, using the Pierce Crosslink Immunoprecipitation (IP) Kit (Thermo Fisher Scientific). Beads were incubated with lysate (∼500 μg protein) for 1 hour at room temperature and then collected using a Magnetic Separation Rack (New England BioLabs). Proteins were eluted from beads per manufacturer's protocol. Eluted proteins were run on 10% SDS-PAGE and Western blot analysis was performed as described above, using antibodies directed to claudin-4 (rabbit anti-human claudin-4; 1:500) and tubulin (rabbit antihuman α-tubulin or β-tubulin; 1:1,000).

Immunofluorescence was performed, as described above, for caspase-3 activation as well as to visualize claudin-4 and tubulin in tumor cells. Primary antibodies used in these assays were the same (including dilutions) as those described above for the proximity ligation assays.

The Microtubules/Tubulin In Vivo Assay Kit (Cytoskeleton Inc.) was used to determine changes in tubulin polymerization with changes in claudin-4 expression. Cells were grown to 80% confluence before being washed with PBS and scraped from 100-cm² plates in tubulin stabilization buffer provided by the manufacturer of the kit and homogenized by pipetting. Cell suspension was centrifuged at 1,000 × g (37°C) for 5 minutes. Cell pellets and supernatant were collected separately and resolved on 10% SDS-PAGE gel. Primary tubulin antibody and HRP-conjugated secondary antibody provided by the manufacturer of the kit were used to determine polymerized (pellet fraction) and nonpolymerized (supernatant fraction) tubulin levels. Densitometry, using ImageJ, was performed to quantify changes in levels of polymerized to nonpolymerized tubulin in each cell line.

Data are presented as mean ± SEM. An unpaired Student t test was used for statistical comparison between control and treatment groups. A one-way ANOVA was used to determine variance among multiple groups, with a Bonferroni Multiple Comparison posttest to determine significance between individual groups. A P value of <0.05 was considered significant.

To identify changes in claudin-4 mRNA expression levels in human ovarian cancer patient tumors compared with normal human ovarian surface epithelium, we queried the Gene Expression Ominbus database. We analyzed expression in the GSE18521 (ID: 200018521) dataset containing 10 normal human ovarian surface epithelium (HOSE) and 53 high-grade serous epithelial ovarian cancer (HGSOC) samples. We found a significant (P = 0.0002) difference in claudin-4 (CLDN4) expression, with 1.8- to 18-fold higher expression in the HGSOC compared with HOSE (Fig. 1A). Next, we examined claudin-4 protein expression in the patient fallopian tube and tumor samples from the University of Colorado Gynecologic Tissue and Fluid Bank (Fig. 1B). The majority of the HGSOC tumor samples expressed claudin-4. In fact, the HGSOC samples expressed significantly (P = 0.0210) more claudin-4 protein than the normal fallopian tube tissue (see Supplementary Fig. S1 for representative Western blot analyses and list of tissue types analyzed). We then examined claudin-4 protein levels in several ovarian tumor cell lines previously identified as having origins from high-grade serous epithelial ovarian tumors (29, 30). In agreement with previous findings, we found that OVCAR3, PEO4, OV429, and OVCAR5 expressed high levels of claudin-4 protein and OVCAR8, OVCAR4, HEY, and DOV-13 cells expressed very low/no claudin-4 protein (Fig. 1C).

To determine whether the presence of claudin-4 in tumor cells alters the response to standard-of-care chemotherapies, we examined the activation of caspase-3 in response to cisplatin and paclitaxel. Claudin-4–expressing OVCAR3 cells exhibited an increase in apoptosis (as measured by percent cleaved caspase-3–positive cells) in response to the claudin-4–disrupting peptide (CMP), cisplatin, and paclitaxel, individually (Fig. 2A and B). Combined treatment with cisplatin and CMP did not enhance cisplatin response (P = 0.3602). However, apoptosis was enhanced nearly 3-fold when cells were cotreated with CMP and paclitaxel compared with paclitaxel alone (P = 0.0002). Similar responses to paclitaxel and CMP were seen in claudin-4–expressing PEO4, OV429, and OVCAR5 cells (Supplementary Fig. S2A–S2C). In contrast, the OVCAR8 (lacking claudin-4 expression) cells were more sensitive to both cisplatin (P = 0.0032) and paclitaxel (P = 0.0063) compared with OVCAR3 cells and CMP did not enhance the apoptotic response to either cisplatin or paclitaxel (Fig. 2C). Analysis of additional claudin-4–deficient EOC cell lines (OVCAR4 and DOV-13, Supplementary Fig. S2D and S2E) consistently showed no enhancement of the apoptotic response to paclitaxel with CMP cotreatment. As a secondary assay of apoptotic response, we used flow cytometry to measure number of OVCAR3 cells exhibiting Annexin V binding and confirmed enhanced apoptosis (Annexin V binding) in response to paclitaxel when cotreated with CMP (Supplementary Fig. S3).

Using a small hairpin RNA (shRNA) specific to claudin-4, we knocked down claudin-4 expression in OVCAR3 cells (“shCLDN4_2” and shCLDN4_3; Fig. 3A). A control, empty vector, shRNA, and shCLDN4_1 did not impact expression of the claudin-4 protein (“shCTRL” and “shCLDN4_1”, respectively). Similar to parental OVCAR3 cells, shCTRL and shCLDN4_1 cells showed increased apoptosis with CMP or paclitaxel alone and a significantly enhanced apoptotic response to paclitaxel in the presence of CMP (P < 0.0001; Fig. 3B). CMP did not induce an apoptotic response in OVCAR3 shCLDN4_2 and shCLDN4_3 cells (cells that successfully lost claudin-4 expression) and CMP cotreatment did not enhance paclitaxel-induced apoptosis (P = 0.8890 and 0.8851, respectively, for shCLDN4_2; P = 0.5024 and 0.7734, respectively, for shCLDN4_3). However, the response to paclitaxel alone was significantly (P < 0.0001) higher in the shCLDN4_2 and shCLDN4_3 cells compared with shCTRL and shCLDN4_1 cells, indicating that disrupting claudin-4 augments the apoptotic response to paclitaxel. Similar results were seen in the claudin-4–expressing PEO4 cell line (Supplementary Figs. S4 and S5).

We next evaluated the impact of overexpressing claudin-4 in OVCAR8 and OVCAR4 cells, which endogenously express no/low claudin-4 protein (Fig. 1). We transfected both cell lines with GFP-tagged claudin-4 or a control GFP vector. Western blot analysis showed strong expression of the claudin-4-GFP (Fig. 4A). GFP control cells did not respond to CMP treatment; apoptosis was not induced with CMP and paclitaxel apoptotic response was not enhanced with CMP cotreatment. However, cells with claudin-4-GFP overexpression had a decreased apoptotic response to paclitaxel and were responsive to CMP, with induction of apoptosis with CMP alone and enhanced apoptotic response to paclitaxel in the presence of CMP (Fig. 4B, representative images of nuclei and activated caspase-3 in claudin-4 nonexpressing control and claudin-4 overexpressing tumor cells are shown in Supplementary Fig. S6). Similar responses were seen in OVCAR4 cells (Supplementary Fig. S7).

We have shown in the experiments above that claudin-4 diminishes the apoptotic response to paclitaxel. We next wanted to determine a potential mechanism by which claudin-4 could affect cell biology to influence paclitaxel response. Upon examination of nuclei in the OVCAR3 cell line, we observed significantly more mitotic figures in shCLDN4_2 cells compared with the claudin-4–expressing shCTRL cells (P < 0.0001; Fig. 5A and B). To confirm this observation, we used a colorimetric mitotic assay kit that utilizes a phospho-histone H3 (Ser28) mAb with HRP-conjugated secondary antibody to detect the level of mitosis. The loss of claudin-4 expression significantly increased the number of cells in the M-phase of mitosis (Fig. 5C, P < 0.0001). To determine whether this increased mitotic index resulted in an increase in cell proliferation, we measured cell number over time in the shCTRL and shCLDN4_2 cell lines. We found that the claudin-4 KO cells proliferated significantly more slowly than the control cells (P = 0.0453, Fig. 5D). Cell-cycle analysis, using flow cytometry (Fig. 5E and F), confirmed significantly (P < 0.0001) more tumor cells in the G₂–M phase at 6 hours postrelease from serum-free conditions in cells that did not express claudin-4 (shCLDN4_2) compared with the cells that did express claudin-4 (shCTRL). These findings suggest that the increase in mitotic figures in the cells lacking claudin-4 expression is not the result of increased proliferation, but reflects an inhibition of G₂–M progression.

Microtubules are known to play an important role in both paclitaxel response and cell-cycle progression. Therefore, we examined whether claudin-4 interacted with microtubules. Proximity ligation assays were performed on OVCAR3 cells with antibodies directed to claudin-4 and either α-tubulin or β-tubulin. Claudin-4 was found to be within interacting distance of both α-tubulin and β-tubulin (Fig. 6A). IP of claudin-4 confirmed the interaction with α-tubulin (Fig. 6B), but we were unable to confirm the interaction with β-tubulin by this method. However, immunofluorescence assays revealed that claudin-4 was colocalized with β-tubulin at the mitotic spindle in tumor cells undergoing mitosis (Fig. 6C). In addition, images of microtubules in claudin-4–expressing (shCTRL cells) and nonexpressing (shCLDN4_2) OVCAR3 cells revealed significant changes in microtubule structure when claudin-4 expression was present (Fig. 6D).

To identify a potential role for claudin-4 in altering microtubule stability, we examined the levels of tubulin within microtubules (polymerized tubulin) compared with free tubulin (nonpolymerized tubulin) in OVCAR3 cells expressing and not expressing claudin-4. Figure 7A shows a shift between polymerized (pellet: p) and nonpolymerized (supernatant: s) tubulin levels when claudin-4 expression is silenced. Densitometry analysis revealed that there was a significant (P = 0.0054) decrease in the percentage of total tubulin found within microtubules (polymerized) in the claudin-4 KO (shCLDN4_2) compared with claudin-4 expressing control (shCTRL) cells (Fig. 7B), suggesting that claudin-4 is enhancing the polymerization of microtubules.

In this study, we have shown that claudin-4 expression reduces the apoptotic response of ovarian tumor cells to paclitaxel. Disruption of claudin-4 expression (shRNA knockdown) or activity (CMP treatment) can enhance or restore the paclitaxel apoptotic response through activation of apoptosis. In addition, we demonstrated that loss of claudin-4 delayed progression through the cell cycle and reduced proliferation. The effect on both apoptosis and the proliferative response in tumor cells may be due the interaction of claudin-4 with microtubules and alteration of tubulin polymerization.

Previously, high claudin-4 expression was shown to be associated with ovarian tumor cells that are resistant to chemotherapeutic agents (24, 25, 31). For example, CD44⁺ ovarian tumor cells were shown to have significantly higher expression levels of claudin-4 compared with CD44⁻ populations isolated from the same tumor. The CD44⁺/high claudin-4–expressing cells were more resistant to carboplatin and paclitaxel treatment compared with the low/no expressing cells (25). In addition, Gao and colleagues have demonstrated that inhibiting claudin-3 and -4 expression with a fragment of the Clostridium perfringens enterotoxin increased the ovarian tumor cell response to both paclitaxel and carboplatin (31). Although our data shows that cells expressing high levels of claudin-4 (OVCAR3) are more resistant to cisplatin compared with the cells that do not express claudin-4 (OVCAR8), CMP was unable to enhance the apoptotic response to cisplatin in the claudin-4–expressing tumor cells. These results suggest the possibility that claudin-4 may regulate the apoptotic response to cisplatin through a different mechanism than paclitaxel, potentially through an interaction of claudin-4 that does not involve the second extracellular loop (which is targeted by CMP).

Previous studies investigating a role for claudin-4 in proliferation have not been consistent. Agarwal and colleagues demonstrated that overexpressing claudin-4 in normal HOSE cells enhanced survival, but did not alter proliferation (32). On the other hand, Ma and colleagues demonstrated that loss of claudin-4 expression inhibits proliferation in MCF7 breast cancer cells in vitro and in vivo (33). The discrepancies could be due to differences in the regulation (or loss of regulation) of claudin-4 in normal versus tumor cells. In addition, our results demonstrate that tumor cells still proliferate in the absence of claudin-4. This suggests that claudin-4 is not required for proliferation but does enhance proliferation when it is present. The observation that loss of claudin-4 leads to increased number of cells in mitosis indicates that claudin-4 likely enhances proliferation by facilitating progression through mitosis.

A proteomic analysis of proteins within interacting distance of claudin-4 in MDCK cells was performed by Fredricksson and colleagues and β-tubulin was identified as a potential interacting partner of claudin-4 (34). Our results confirm an interaction of claudin-4 with both α-tubulin and β-tubulin in ovarian tumor cells and that this interaction changes during the cell cycle. We were able to capture an interaction of claudin-4 at sites at or near the plasma membrane as well as within the mitotic spindle. Further analysis of this interaction will be needed to fully understand how the interaction of claudin-4 with microtubules may be regulating tumor cell survival and growth. Our results do suggest that interaction of claudin-4 with tubulin may alter the polymerization of tubulin. McGrail and colleagues have shown that ovarian tumor cells that are resistant to paclitaxel-induced death have altered microtubule stability (35). Therefore, it is possible that claudin-4 alters microtubule dynamics in a way that makes tumor cells more resistant to paclitaxel-induced death.

In conclusion, we have demonstrated that claudin-4 plays an important role in tumor cell resistance to paclitaxel. Therefore, expression of claudin-4 by tumor cells may be a biomarker for patient response to paclitaxel. In addition, the development of a claudin-4 therapeutic (e.g., CMP) could provide a novel way to improve the patient response to paclitaxel.

No potential conflicts of interest were disclosed.

Conception and design: C.A. Breed, B.G. Bitler, K. Behbakht, H.K. Baumgartner

Development of methodology: C.A. Breed, D.A. Hicks, B.G. Bitler, K. Behbakht, H.K. Baumgartner

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): C.A. Breed, D.A. Hicks, C.E. Galimanis, B.G. Bitler, K. Behbakht, H.K. Baumgartner

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): C.A. Breed, D.A. Hicks, K. Behbakht, H.K. Baumgartner

Writing, review, and/or revision of the manuscript: C.A. Breed, B.G. Bitler, K. Behbakht, H.K. Baumgartner

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): P.G. Webb, C.E. Galimanis, K. Behbakht, H.K. Baumgartner

Study supervision: K. Behbakht, H.K. Baumgartner

The authors wish to acknowledge Dziuleta Cepeniene of the Peptide and Protein Chemistry Core at the University of Colorado for synthesizing CMP. We would also like to acknowledge that imaging was made possible by the University of Colorado Advanced Light Microscopy Core, supported in part by NIH/NCRR CCTSI grant UL1 RR025780. Flow cytometry was performed by the University of Colorado Cancer Center Flow Cytometry Shared Resource (supported by NCI Cancer Center Support Grant P30CA046934). Funding was also provided by the Department of Obstetrics and Gynecology, University of Colorado Denver.

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.