• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of iaiPermissionsJournals.ASM.orgJournalIAI ArticleJournal InfoAuthorsReviewers
Infect Immun. Mar 2005; 73(3): 1275–1283.
PMCID: PMC1064919

Vibrio parahaemolyticus Disruption of Epithelial Cell Tight Junctions Occurs Independently of Toxin Production

Abstract

Vibrio parahaemolyticus is a leading cause of seafood-borne gastroenteritis worldwide. Virulence is commonly associated with the production of two toxins, thermostable direct hemolysin (TDH) and TDH-related hemolysin (TRH). Although the majority of clinical isolates produce TDH and/or TRH, clinical samples lacking toxin genes have been identified. In the present study, we investigated the effects of V. parahaemolyticus on transepithelial resistance (TER) and paracellular permeability in Caco-2 cultured epithelial cells. We found that V. parahaemolyticus profoundly disrupts epithelial barrier function in Caco-2 cells and that this disruption occurs independently of toxin production. Clinical isolates with different toxin genotypes all led to a significant decrease in TER, which was accompanied by an increased flux of fluorescent dextran across the Caco-2 monolayer, and profound disruption of actin and the tight junction-associated proteins zonula occludin protein 1 and occludin. Purified TDH, even at concentrations eightfold higher than those produced by the bacteria, had no effect on either TER or paracellular permeability. We used lactate dehydrogenase release as a measure of cytotoxicity and found that this parameter did not correlate with the ability to disrupt tight junctions. As the effect on barrier function occurs independently of toxin production, we used PCR to determine the toxin genotypes of V. parahaemolyticus isolates obtained from both clinical and environmental sources, and we found that 5.6% of the clinical isolates were toxin negative. These data strongly indicate that the effect on tight junctions is not due to TDH and suggest that there are other virulence factors.

Vibrio parahaemolyticus is a leading cause of seafood-borne gastroenteritis worldwide, which is often caused by ingestion of raw or improperly cooked shellfish (2, 11, 20, 33). The infection is usually self-limiting and of moderate severity. The symptoms include watery diarrhea, abdominal cramps, nausea, vomiting, and headache (2, 25). V. parahaemolyticus is routinely cultured from estuarine environments during the warmer months of the year. Although outbreaks commonly occur in Japan and other countries in Southeast Asia (10, 20), there were two recent major oyster-associated outbreaks of V. parahaemolyticus gastroenteritis in North America (in 1997 in British Columbia and Washington and in 1998 in Texas, New York, and Washington) involving over 200 cases (7, 11, 22, 29). Interestingly, there was a difference between the strains isolated in the Pacific Northwest and the strains isolated in the Gulf and eastern coasts with respect to serotype and toxin genotype (7, 11). The majority of clinical isolates from the Pacific Northwest belonged to serotype O4:K12 and produced both the thermostable direct hemolysin (TDH) and the TDH-related hemolysin (TRH) (21, 29, 35). However, the clinical isolates from the outbreak on the Gulf and eastern coasts were serotype O3:K6 and contained only the TDH gene (12, 22, 36). The emergence of this O3:K6 clone in 1996 has recently been shown to be associated with a rise in V. parahaemolyticus outbreaks in Japan and Taiwan (1, 36).

Most of the studies examining the effects of V. parahaemolyticus on the host have focused on TDH and TRH. TDH causes a number of cytotoxic effects, including erythrocyte lysis, disruption of the microtubule cytoskeleton, and ion influx into cultured cells (13, 17, 39). Less is known about the targets of TRH, although studies with purified toxin have shown that it too can lyse erythrocytes and produce fluid accumulation in the rabbit ileal loop model (23, 54). The results of recent studies suggest that TDH-induced ion influx may cause cell rounding and subsequent disruption of epithelial barrier function (39). The loss of barrier function may contribute to the diarrhea associated with V. parahaemolyticus infections, as well as infections caused by other enterovirulent bacteria (34, 42, 44).

Barrier function is maintained by tight junctions, which are dynamic cell-cell adhesions that form a continuous seal around the cell and control the permeability of solutes and fluids across the paracellular pathway (55). Tight junctions are comprised of transmembrane proteins, such as claudins and occludin, as well as cytosolic proteins which are recruited to the apicolateral membrane, including zonula occludin protein 1 (ZO-1), ZO-2, ZO-3, cingulin, and 7H6 (3, 51). Cytoskeletal components are anchored to tight junction structures through the cytosolic proteins, forming the perijunctional actin-myosin ring which gives the junctions architectural support, and they may also be involved in shuttling signaling molecules to and from the junctions (26, 27).

Previously published evidence suggests that there is a correlation between disease and TDH and/or TRH production; however, studies in which the presence of toxin genes in clinical isolates has been examined have revealed a small, unexplained number of strains that lack the genes encoding both toxins (19, 23, 45, 48). These findings suggest that V. parahaemolyticus may harbor other virulence factors in addition to TDH and TRH. In the present study, we examined the effects of naturally occurring V. parahaemolyticus strains, isolated from clinical and environmental sources and having different toxin genotypes, on epithelial barrier structure and function. We examined the effects on both transepithelial resistance (TER) and paracellular permeability across Caco-2 cell monolayers. Interestingly, we found that some V. parahaemolyticus strains, regardless of the site of isolation or the toxin genotype, had profound effects on tight junction structure and function.

MATERIALS AND METHODS

Culture of V. parahaemolyticus strains.

Strains of V. parahaemolyticus were received from the Southern Alberta Provincial Laboratory in Calgary, Alberta, Canada (four clinical isolates) and the Canadian Food Inspection Agency (CFIA) in Burnaby, British Columbia, Canada (73 clinical and 156 environmental isolates). Strains classified as clinical were cultured from patient samples or food associated with disease. The environmental strains included isolates from seawater or oysters collected from harvest sites predominantly in southern British Columbia, Canada. We also included the type strain (ATCC 17802) in our studies (14). For all experiments, cultures were grown at 37°C for 18 h with shaking (225 rpm) in nutrient broth (Difco, Sparks, Md.) supplemented with 3% NaCl (NBS) unless indicated otherwise.

Detection of TDH production.

TDH production was measured by using a Kanagawa phenomenon-reverse passive latex agglutination kit (Denka Seiken Co. Ltd., Tokyo, Japan), in which latex beads coated with anti-TDH immunoglobulin G are used. Strains of interest were grown either in mannitol-peptone broth (2% Bacto Peptone [Difco], 0.5% d-mannitol, 5% NaCl; pH 7.8) to induce TDH production or in Dulbecco modified Eagle medium (DMEM) containing 10% fetal bovine serum and 10 mM sodium pyruvate. In both cases, bacteria were grown for 18 h at 37°C with shaking. In addition, TDH titers were determined for the culture fluids obtained from Caco-2 cells infected for 3 h with tdh+ V. parahaemolyticus isolates to determine whether the presence of Caco-2 cells affected TDH production. Twofold dilutions of a cell-free culture supernatant were examined by using the manufacturer's instructions. We included purified TDH toxin (1 hemolytic unit [HU]/ml) as a positive control and nonsensitized latex beads as a negative control. The reciprocal of the highest dilution that gave a positive reaction was defined as the TDH titer.

TER of Caco-2 monolayers.

Transwells (diameter, 12 mm; pore size, 0.4 μm; Corning Inc., Corning, N.Y.) were treated with sterile collagen (type I; Sigma-Aldrich, St. Louis, Mo.) and UV sterilized before Caco-2 cells (ATCC HTB-37) were added to the apical compartment of each well. The cells were seeded at an initial density of 4 × 104 cells/cm2 in DMEM (Gibco, Grand Island, N.Y.) supplemented with 10% fetal bovine serum and 1 mM sodium pyruvate; 1.5 ml of the same medium was added to the basolateral compartment. The cells were incubated at 37°C in a 5% CO2 atmosphere for approximately 1 week, and the medium was changed every second day. TER was measured by using an epithelial voltohmmeter (World Precision Instruments Inc., Sarasota, Fla.); only wells displaying baseline resistance readings greater than 350 Ω/cm2 were used for the experiments. V. parahaemolyticus (0.5 μl) was added to the upper chamber of each well, and TER measurements were obtained hourly. Data are reported below as percentages of the preinfection TER remaining across the monolayer and are the means ± standard errors of the means for at least three independent experiments. Statistical differences between groups were determined by analysis of variance (ANOVA), followed by the Student-Newmann-Keuls multiple-range test (InStat; GraphPad Software).

In vitro permeability assays.

Paracellular permeability was studied by measuring the apical-to-basolateral flux of sulforhodamine dextran (Texas Red-dextran; molecular weight, 10,000; 100 μmol/liter; Molecular Probes, Eugene, Oreg.) as previously described (43). Transwells containing Caco-2 monolayers were infected with V. parahaemolyticus (0.5 μl) and washed with sterile Ringer's solution. The upper chambers were filled with 500 μl of Texas Red-dextran (100 nmol/liter), and 1.0 ml of Ringer's solution was added to the bottom chamber. After incubation for 3 h at 37°C in the presence of 5% CO2, two 300-μl samples were recovered from the bottom chamber and serially diluted on a 96-well plate, and the fluorescence was measured with a microplate fluorometer (Wallac Victor2 1420 multilabel counter; Perkin-Elmer Life Sciences, Boston, Mass.). The results obtained were expressed as the percentage of apical dextran that crossed the Transwell membrane per square centimeter per hour. The data given below are the means ± standard errors of the means for at least three independent experiments. Statistical differences between groups were determined by ANOVA, followed by the Student-Newmann-Keuls multiple-range test.

Immunofluorescence microscopy.

Caco-2 cells were seeded at an initial density of 4 × 104 cells/cm2 onto type I collagen-coated coverslips and incubated at 37°C in the presence of 5% CO2 for 3 days. The cells were infected with 1 μl of an overnight culture. To assess whether live bacteria were required to induce cytoskeletal and junctional rearrangements, approximately 2 × 108 bacteria (1 ml of an overnight culture of BCC34) were killed by UV irradiation (900,000 μJ; three pulses; UV Stratalinker) and added to Caco-2 cells. Killing was assessed by culturing the bacteria on nutrient broth agar. Cells were infected for 3 h and fixed with 2.5% paraformaldehyde in phosphate-buffered saline (PBS) for 10 min at 37°C. Next, the cells were rinsed with PBS supplemented with 0.5% Triton X-100 three times before they were blocked with 1% normal goat serum in PBS supplemented with 0.1% Triton X-100. The cells were double labeled either with mouse anti-occludin (1:100; BD Biosciences, Mississauga, Ontario, Canada) and Alexa488 phalloidin or with rabbit anti-ZO-1 (1:200; Zymed, San Francisco, Calif.) and Alexa488 phalloidin (1:200; Molecular Probes). Samples were visualized by using a Leica DMIERB2 inverted microscope equipped with a ×63 oil immersion objective and an Orca ER cooled charge-couple device camera. Images were acquired by using OpenLab (Improvision), and between 18 and 30 0.2-μm z sections were collected from each image. Projected images and xz and yz images were constructed from the z-section series by using Volocity 3.0 (Improvision), and figures were prepared by using Adobe Photoshop. The figures show representative images from at least three independent experiments.

DNA extraction for PCR.

V. parahaemolyticus isolates were inoculated into 5 ml of NBS from frozen stock and grown overnight at 37°C with shaking. One milliliter of an overnight culture was microcentrifuged at 5,400 × g for 10 min at room temperature, and the resulting pellet was resuspended in double-distilled water; this was followed by heating at 95°C for 10 min. The suspension was placed on ice for 10 min, treated with RNase A (10 μg/ml), and microcentrifuged at 15,700 × g for 30 min at 4°C. The supernatant was used as the template for a toxin profiling PCR.

PCR profiling for toxin genes.

The primer sequences used for PCR amplification of the genes encoding TRH (trh), TDH (tdh), and thermolabile hemolysin (TLH) (tlh) are listed in Table Table1.1. The TDH and TRH primers are identical to those currently used by the CFIA for surveillance for V. parahaemolyticus in oysters harvested off the west coast of British Columbia. We used the TLH (tlh) toxin for species identification to confirm the results of the automated bacterial identification system (VITEK; bioMerieux, Durham, N.C.) and R72H (30) PCR results provided by the CFIA. The tdh and tlh PCR mixtures (final volume, 25 μl) contained 10× PCR buffer (50 mM KCl, 10 mM Tris [pH 8.5], 0.1% gelatin, 1.5% MgCl2), each deoxynucleoside triphosphate (Invitrogen Life Technologies, Burlington, Ontario, Canada) at a concentration of 0.25 mM, Taq polymerase (1.25 U; New England Biolabs Ltd., Mississauga, Ontario, Canada), and nuclease-free water (Gibco). We used touchdown PCR to amplify the TDH and TLH genes. The amplification conditions were as follows: one cycle of 95°C for 3 min, followed by five cycles of denaturation at 95°C for 1 min, annealing at 58°C for 1 min, and extension at 72°C for 2 min; then in the subsequent 30 cycles the annealing temperature was decreased by 0.5°C every five cycles for a total of 35 cycles (final annealing temperature, 55°C). The conditions for amplifying the trh gene were identical to those used for tdh and tlh, except that the MgCl2 concentration was 2.0 mM. A similar touchdown program was used to amplify trh with annealing temperatures ranging from 53 to 50°C.

TABLE 1.
Primer sequences used for PCR toxin profiling

Southern hybridization.

PCR profiling was validated by using Southern hybridization to ensure specificity and sensitivity. The following four strains, representing different toxin profiles, were used: ATCC 17802 (Δtdh/trh+/tlh+), NY477 (tdh+trh/tlh+), DAL1094 (Δtdhtrh/tlh+), and WA1029 (tdh+/trh+/tlh+). The ATCC 17802 strain profile was based on our PCR results, and the other three profiles were determined previously by the CFIA.

Cultures (20 ml) were grown overnight at 37°C with shaking (225 rpm) in NBS. The cultures were microcentrifuged (5,400 × g, 5 min), and the cells were resuspended in 15 ml of a solution containing 10 mM NaCl, 20 mM Tris-HCl (pH 8.0), 1 mM EDTA, 100 μg of proteinase K per ml, and 0.5% (wt/vol) sodium dodecyl sulfate and incubated at 50°C overnight. Genomic DNA was extracted by using phenol-chloroform (30, 32). The DNA from each strain was digested with four different restriction enzymes (EcoRV, HincII, HindIII, and EcoRI; New England Biolabs Ltd.) and resolved on 0.8% agarose gels prior to immobilization onto Hybond-N membranes (Amersham Pharmacia Biotechnology Inc., Piscataway, N.J.). Segments of the toxin genes (tdh, trh, and tlh) were amplified by using primers shown in Table Table11 and were used to prepare labeled probes. DNA was labeled with [32 P]dCTP by using Ready to Go DNA labeling beads (Amersham Pharmacia) according to the manufacturer's instructions. Membranes were moistened with rapid-Hyb buffer (Amersham-Pharmacia) prior to addition of a denatured DNA fragment probe (7 × 106 cpm) and salmon sperm (Sigma). Hybridization was performed overnight at 65°C in a VWR model 5400 hybridization oven with constant rotation. The membranes were washed twice in 2× SSC at room temperature for 5 min, twice in 2× SSC-1% sodium dodecyl sulfate at 65°C for 30 min, and twice in 0.1× SSC at room temperature for 30 min (1× SSC is 0.15 M NaCl plus 0.015 sodium citrate). The membranes were air dried and exposed to Kodak BioMax film at −70°C overnight.

Cytotoxicity assay.

We measured release of lactate dehydrogenase (LDH) (Cytotox 96; Promega) in order to examine V. parahaemolyticus cytotoxicity. Caco-2 cells were seeded into 24-well plates at an initial density of 5 × 104 cells/ml in DMEM supplemented with 10% fetal bovine serum and incubated at 37°C in the presence of 5% CO2 overnight. The next day, the medium was replaced by fresh serum-free DMEM lacking phenol red (0.5 ml/well), and the cells were infected with overnight cultures of V. parahaemolyticus (0.5 μl/well). After 3 h, the medium was removed, microcentrifuged for 2 min at 15,700 × g to remove debris, and used to measure the LDH released during the course of infection. The remaining cells were incubated in phenol red-free DMEM containing 1% Triton X-100 at 37°C for 45 min, and the supernatants were collected, microcentrifuged for 5 min at 15,700 × g, and used to measure the total cellular LDH content. The LDH content was measured by following the manufacturer's instructions. The percentage of the LDH released was calculated as follows: 100 × (optical density at 490 nm of the culture supernatant/optical density at 490 nm of the lysed cells). The data below are the means ± standard deviations for at least three independent experiments, and statistical differences between groups were determined by ANOVA, followed by the Student-Newmann-Keuls multiple-range test.

RESULTS

V. parahaemolyticus disrupts epithelial barrier function.

We examined the effects of V. parahaemolyticus infection on epithelial permeability by measuring the TER across Caco-2 monolayers. Monolayers were infected with V. parahaemolyticus clinical isolates previously shown to have genes for TDH (BCC36), TRH (ATCC 17802), or both (WA10290) or were treated with purified TDH (3 HU/ml). All three strains were isolated from patients with confirmed V. parahaemolyticus gastroenteritis. We measured the levels of TDH produced by these strains under three different culture conditions using a latex agglutination assay as described in Materials and Methods, and we compared the results to those obtained with purified toxin. Under all conditions, the amount of TDH produced by the bacteria was at least eight times less than the lowest concentration of purified toxin commonly used (1 HU) (24, 39, 40). We detected purified TDH at levels up to a titer of 1,024, whereas the titer for BCC36, which produced the highest concentration of TDH, was only 128 when the organism was cultured overnight in mannitol-peptone broth (Table (Table2).2). Additionally, TDH was not detected in culture supernatants obtained from Caco-2 cells infected for 3 h with either WA10290 or BCC36. TER measurements were taken hourly over the 3-h course of infection and were expressed as percentages of the original TER remaining across the Caco-2 cell monolayers at each time. All three isolates caused a significant decrease in TER after 2 h of infection (Fig. (Fig.1A).1A). The TER measurements for the clinical isolates at 2 and 3 h postinfection were not significantly different from each other despite differences in the presence of toxin genes. At 3 h postinfection the values were as follows (P > 0.05): ATCC 17802, 30.53% ± 3.06%; NY477, 29.27% ± 6.95%; BCC36, 17.27% ± 2.57%; and WA10290, 16.30% ± 3.60%. In contrast, purified TDH (3 HU/ml) had no effect on TER, since at all times the resistance measurements were not significantly different from those obtained for uninfected control monolayers (P > 0.05) (Fig. (Fig.1A).1A). Addition of WA10290 to the basolateral compartment of a Transwell had no effect; at 3 h the value was 98.03% ± 1.98% (P > 0.05).

FIG. 1.
V. parahaemolyticus infection decreases the transepithelial resistance across Caco-2 monolayers: TER measurements after infection with toxigenic (A) or nontoxigenic (B) V. parahaemoltyicus isolates. The toxin genotypes of the strains in panel A are as ...
TABLE 2.
Strain genotypes and reverse latex agglutination test for TDH production

V. parahaemolyticus isolates lacking tdh and trh disrupt epithelial barrier function.

As our results suggested that the disruption of the epithelial barrier function is not solely due to TDH or TRH production, we examined the effects of four different Δtdhtrh V. parahaemolyticus strains on the TER across Caco-2 monolayers. BCC34 is a clinical isolate, DAL1094 was isolated from an oyster implicated in a case of gastroenteritis, and BCE306 and BCE515 are environmental isolates obtained from the coastal waters of southern British Columbia. The lack of TDH production by these strains was confirmed by the latex agglutination assay (Table (Table22 and data not shown). Addition of both of the disease-associated isolates, BCC34 and DAL1094, led to a significant decrease in TER by 2 h postinfection (Fig. (Fig.1B)1B) (BCC34, 13.47% ± 0.814%; DAL1094, 22.87% ± 6.53%). Surprisingly, one environmental isolate, BCE306, also decreased TER (20.27% ± 0.57%). In contrast, environmental isolate BCE515 had no effect on TER (90.80% ± 1.43%). The decrease in TER observed in experiments performed with the three Δtdh/Δtrh strains was not significantly different from the decease observed with the toxin-positive isolates shown in Fig. Fig.1A1A (P > 0.05). Similar to what was observed with the toxigenic isolates, addition of BCC34 basolaterally was ineffective (at 3 h the value was 99.61% ± 4.74% [P > 0.05]). Taken together, our data suggest that V. parahaemolyticus disrupts TER and that this disruption occurs independently of toxin production.

The decrease in TER is associated with an increase in paracellular permeability.

The observed decrease in TER could have been caused by either an increase in paracellular permeability or a change in ion flux across an intact monolayer. To distinguish between these possibilities, we measured the apical-to-basolateral flux of fluorescently labeled dextran (molecular weight, 10,000) across Caco-2 monolayers. Cells were infected with V. parahaemolyticus or treated with TDH (3 HU/ml) for 1.5 h prior to addition of dextran to the apical side of the washed monolayers. After a 3-h incubation, samples were removed from the basolateral side of the membrane, the fluorescence was measured, and the dextran flux was calculated as described in Materials and Methods. Infection with all three toxin-positive isolates led to significant enhancement of paracellular permeability, although there were differences in the magnitude of the effect (Fig. (Fig.2).2). WA10290 (tdh+/trh+) caused the greatest increase in paracellular permeability, with 43.25% ± 1.48% (P < 0.001) of the dextran crossing the monolayer. BCC36 (tdh+/Δtrh) had an intermediate effect, whereas type strain ATCC 17802 disrupted permeability the least (5.0% ± 0.6%; P < 0.05). We next examined the effect of the Δtdh/Δtrh strains. All three strains that caused a significant decrease in TER also enhanced paracellular permeability, albeit to different degrees (Fig. (Fig.2).2). The environmental isolate BCE306 had the strongest effect, with 37.38% ± 1.09% (P < 0.001) of the fluorescent dextran crossing the Caco-2 monolayer. BCC34 had an intermediate effect (22.88% ± 0.92%; P < 0.001), whereas DAL1094 was the least potent (5.97% ± 1.08%; P < 0.05). Collectively, our data suggest that the decrease in TER as a result of infection with either toxin-positive or toxin-negative strains is most likely due to enhancement of paracellular permeability.

FIG. 2.
The decrease in TER is accompanied by an increase in paracellular permeability. The apical-to-basolateral dextran flux in response to 1.5 h of infection with V. parahaemolyticus is shown. The data are the percentages of the apical sulforhodamine dextran ...

Tight junction disruption is accompanied by a dramatic reorganization of junction-associated proteins.

Alterations in TER and paracellular permeability are often accompanied by changes in the localization of tight junction proteins. The effect of V. parahaemolyticus infection on the localization of occludin (a transmembrane protein), ZO-1 (a peripheral membrane protein), and filamentous actin was examined by using immunofluorescence microscopy, as described in Materials and Methods (Fig. (Fig.3).3). In uninfected cells, occludin and actin (Fig. (Fig.3,3, left panels) and ZO-1 and actin (right panels) were located at the periphery of the apical side of the monolayer. Infection with WA10290 (tdh+/trh+), BCC34 (Δtdh/Δtrh), or BCE306 (Δtdh/Δtrh), all of which significantly altered epithelial barrier function, resulted in profound disruption of occludin, ZO-1, and actin. All three proteins appeared to be redistributed throughout the Caco-2 cell, as shown in the xz and yz images, and formed intensely staining aggregates. This effect required live bacteria and was not observed after addition of bacterial culture supernatant or killed whole bacteria (data not shown). In contrast, addition of TDH and BCE515, which had no effect on either TER or paracellular permeability, led to only minor alterations in the localization of actin and occludin, which were more diffusely labeled at the cell periphery compared to what we observed in uninfected cells. Additionally, there was no effect on the cells when the inoculum of strain BCE515 was doubled (data not shown), which eliminated the possibility that the disruptive effects were due to artifacts created by bacterial cell number.

FIG. 3.
V. parahaemolyticus infection causes a profound disruption of the actin cytoskeleton: immunofluorescence micrographs of Caco-2 cells exposed to V. parahemolyticus or TDH (3 HU/ml) for 3 h. Samples were double labeled for actin (green) and occludin (red) ...

Cytoskeletal disruption does not correlate with cytotoxicity.

We examined whether disruption of the actin cytoskeleton was accompanied by cytotoxicity by measuring LDH release from Caco-2 cells infected with each of seven V. parahaemolyticus isolates. As shown in Fig. Fig.4,4, there was no statistically significant increase in LDH release after infection for 3 h with any of the V. parahaemolyticus isolates shown to cause epithelial barrier function disruption (Fig. (Fig.4).4). We examined cytotoxicity after 6 h of infection with either WA10290 (tdh+/trh+), BCC34 (Δtdh/Δtrh), or BCE306 (Δtdh/Δtrh) and observed a small, statistically insignificant increase in the percentage of LDH released into the culture medium (Fig. (Fig.4).4). At this time, the monolayers were severely disrupted, and there was evidence of cell loss (data not shown). As expected, infection with BCE515 (Δtdh/Δtrh), which did not affect barrier function, was not cytotoxic at either time. In contrast, exposure to purified TDH (3 HU/ml) for either 3 or 6 h caused significant cytotoxicity without disrupting epithelial barrier function. Collectively, these results suggest that the disruption of epithelial barrier function does not correlate with Caco-2 cell cytotoxicity.

FIG. 4.
Cytotoxicity of V. parahaemolyticus isolates: percentage of LDH released from Caco-2 cells infected with V. parahaemolyticus isolates for 3 h (open bars) or 6 h (solid bars). The data are means ± standard deviations for at least three independent ...

Prevalence of Δtdhtrh clinical isolates.

Our data suggest that some Δtdhtrh V. parahaemolyticus isolates cause profound disruption of epithelial barrier function; therefore, we examined the prevalence of toxin-negative strains isolated from patients or food sources associated with confirmed V. parahaemolyticus disease. We used PCR to amplify genes for TDH, TRH, and TLH, which was used for species identification. The tdh and trh genes are 68% identical (9, 11), so to ensure that there was no cross-reactivity with the primers, we examined four representative strains by both PCR and Southern hybridization. As shown in Fig. Fig.5A,5A, we obtained identical results with both methods. All isolates were positive for the species marker, TLH (Fig. (Fig.4A).4A). NY477 was confirmed to be tdh+/Δtrh, ATCC 17802 was confirmed to be Δtdh/trh+, WA10290 was confirmed to contain both the tdh and trh genes, and DAL1094 was confirmed to have neither gene (Fig. 4B and C). Based on these results, we used PCR to determine the toxin genotypes of 72 clinical isolates and, as a control, 87 environmental isolates. As shown in Table Table3,3, although the majority of the clinical isolates contained both the tdh and trh genes, 5.5% were doubly toxin negative. In this population, 9.7% of the strains had the tdh gene alone, whereas 6.9% of the strains were Δtdh/trh+. Although environmental isolates that were collected from seawater or oysters from southern British Columbia were predominantly toxin negative (94.2%), 5.8% were positive for the trh gene, a prevalence similar to that observed for the clinical isolates.

FIG. 5.
Comparison of PCR and Southern hybridization results for four representative V. parahaemolyticus strains. (A) tdh detection; (B) trh detection; (C) tlh detection. Genes were detected by PCR (left panels) and Southern hybridization (right panels). Genomic ...
TABLE 3.
Summary of PCR toxin profiling results

DISCUSSION

Virulence in V. parahaemolyticus has historically been correlated with the production of the hemolytic toxins TDH and TRH. In this study, we found that V. parahaemolyticus can profoundly disrupt epithelial barrier function and that this disruption occurs independently of toxin production. Infection with V. parahaemolyticus led to a significant decrease in TER, and there was no difference in the magnitude of this effect between toxin-positive and toxin-negative isolates. The decrease in TER was accompanied by an increase in paracellular permeability, which was associated with a dramatic disruption of the enterocyte actin cytoskeleton, along with abnormalities in occludin and ZO-1 localization. These effects were not observed in response to purified TDH and were not restricted to clinical isolates as one toxin-negative environmental isolate was as potent at disrupting barrier function. Collectively, these data indicate that TDH and TRH do not play an essential role in the disruption of the epithelial barrier function and suggest that there are other, as-yet-unidentified V. parahaemolyticus virulence determinants.

Most of the studies examining the contribution of TDH and TRH to V. parahaemolyticus virulence have focused on the effects of exposure to purified toxins. Results from these studies suggest that in addition to red blood cell lysis (17), TDH has a number of potent effects on intestinal epithelial cells. Exposure to TDH has been documented to lead to an increase in short circuit current (Isc), to enhancement of Ca2+ entry from the extracellular medium, and to Ca2+-dependent chloride secretion (12, 16, 37). Although less well studied, TRH has also been demonstrated to elevate the intracellular Ca2+ concentration and to enhance chloride secretion (39, 40, 49). Raimondi and colleagues found that at high concentrations (>5 HU/ml) TDH caused both a decrease in TER across Caco-2 monolayers and enhanced cytotoxicity as measured by LDH release (39). Similar to our findings, lower concentrations (<5 HU/ml) had no effect on TER. In a second study the same group examined the effect of purified TDH on cytoskeletal organization. In this study, rat IEC6 intestinal cells were exposed to 2.5 HU of TDH per ml for 18 h, which resulted in microtubule-dependent reorganization of the cell cytoskeleton (13). The effects reported in these studies occurred at higher toxin concentrations than we and others have observed in bacterial culture supernatants (Table (Table2)2) (37) and with a longer exposure. Raimondi and colleagues suggested that high concentrations of TDH may occur in a stagnant intestinal loop with little luminal clearance, but they did not rule out an effect of other virulence determinants (39). Our results do not exclude a role for TDH and/or TRH; these toxins may exert their effects at later stages of infection. Instead, our results suggest that the effect on epithelial barrier function, which appears early (1.5 to 3 h postinfection), occurs independently of toxin production.

The role of TDH in infection with tdh+ V. parahaemolyticus strains is less clear. The effect of deleting the tdh gene was examined in two independent studies which produced opposite results. Nishibuchi and colleagues deleted both the tdh1 and tdh2 genes from strain AQ3815T and found that the mutant was completely attenuated in the ability to increase Isc and to cause fluid accumulation in ligated rabbit ileal loops (31). In contrast, deletion of tdh1 and tdh2 from the pandemic strain RIMD2210633, which lacks TRH, did not decrease cytotoxicity, as measured by trypan blue uptake, and resulted in only partial inhibition of fluid accumulation in the ligated rabbit ileal loop model, suggesting the presence of other virulence determinants (38). The possible explanations for these discrepancies include differences in the experimental systems used and the endpoints studied by the two groups and differences in the genetic backgrounds of the strains examined. There are over 300 serotypes of V. parahaemolyticus, and 75 serotypes have been identified as pathogenic (6), suggesting that there is genetic diversity within the species. This suggests that virulence in V. parahaemolyticus is complex and may involve both strain-specific and common virulence determinants. Our studies, which demonstrated that V. parahaemolyticus isolates with different toxin genotypes can have similar effects on epithelial barrier function, may reveal a common toxin-independent mechanism.

In this study, we examined the prevalence of toxin-negative strains in clinical isolates collected in North America between 1999 and 2001. The prevalence of toxin-negative isolates in our collection (5.6%) was consistent with the prevalence observed in samples collected from other populations. For example, in studies examining the occurrence of the tdh and trh genes in patient samples obtained in France (41), Japan (45), and the United States (4) the workers found that between 9 and 18.5% of the strains were toxin negative. In all three studies, the majority of the strains contained either tdh or trh or both, although there were site-specific differences in the prevalence of each toxin genotype. A compelling question is whether the toxin-negative clinical strains identified in these studies disrupt epithelial barrier function. Our studies suggested that tdh and/or trh is not required for disruption but is present in the majority of clinical strains. It is tempting to speculate that TDH may also contribute to virulence by promoting survival within the host gastrointestinal tract. This hypothesis is supported by the observation that TDH production is enhanced in the presence of bile salts and at pH 5.5 to 6.5, conditions that V. parahaemolyticus may encounter in the gastrointestinal tract (8, 37).

Our results indicate that disruption of the epithelial barrier function does not correlate with cytotoxicity, as measured by LDH release. Only TDH (3 HU/ml) (Fig. (Fig.4),4), which had no effect on barrier function, resulted in a significant increase in LDH release from Caco-2 cells. These results are in agreement with results of Raimondi and colleagues, who observed significant levels of LDH in the culture media of Caco-2 cells exposed to high concentrations (>25 HU/ml) of TDH (39). LDH release measures the release of a cytosolic enzyme resulting from cell lysis and does not detect cells beginning to undergo apoptosis. Although purified TDH can induce apoptosis in Rat1 cells (30), it is not known whether V. parahaemolyticus infection has the same effect.

The disruption of epithelial barrier function mediated by V. parahaemolyticus is associated with a profound reorganization of the enterocyte actin cytoskeleton. Infection leads to the redistribution and aggregation of actin, occludin, and ZO-1 within the Caco-2 cell. This effect appears to be similar morphologically to the effect observed in cells exposed to either enterotoxin A or B from Clostridium difficile or to the RTX toxin and hemagglutinin/protease expressed by Vibrio cholerae. These toxins cause reorganization of the perijunctional actin ring, leading to barrier function disruption. C. difficile toxins act to glucosylate the Rho family GTPases Rho, Rac, and Cdc42, which are important regulators of cytoskeletal dynamics (5). Exposure to the V. cholerae RTX toxin results in cell rounding due to covalent cross-linking of cellular actin (15). Exposure to V. cholerae hemagglutinin/protease results in disruption of tight junctions and the actin cytoskeleton, due in part to occludin proteolysis (52, 53). Although the V. parahaemolyticus genome encodes several proteases (28), infection in the presence of a broad-spectrum protease inhibitor cocktail does not inhibit epithelial barrier function disruption (unpublished observations). These mechanisms are in contrast to the mechanisms of pathogens that selectively target tight junction components. For example, tight junction disruption by enteropathogenic Escherichia coli involves the phosphorylation of myosin light chain kinase, occludin dephosphorylation, and activation of the membrane cytoskeletal linker ezrin (9, 43, 46, 47, 56). Barrier function disruption by the protozoan parasite Giardia lamblia can be reversed by addition of inhibitors of myosin light chain kinase and caspase-3 (9, 43). Preliminary data from our laboratory suggest that the cytoskeletal disruption observed in response to V. parahaemolyticus is not due to actin cross-linking and is not reversible upon addition of myosin light chain kinase, caspase-3, or protease inhibitors (unpublished observations). We are actively investigating the mechanism used by V. parahaemolyticus to disrupt tight junctions and the cytoskeleton.

Our data demonstrating that both toxigenic and nontoxigenic V. parahaemolyticus isolates can disrupt epithelial barrier function suggest the presence of additional, as-yet-unidentified virulence factors. The description of the genome sequence of one V. parahaemolyticus isolate, the pandemic strain RIMD2210633, revealed the presence of two putative type III secretion systems (TTSS) (28). One system, located on chromosome 1, is ubiquitous in both clinical and environmental isolates and exhibits homology with the TTSS from Yersinia species. A second system, in a putative pathogenicity island on chromosome II, appears to be restricted to clinical isolates and is homologous to the enteropathogenic E. coli TTSS. Additionally, homologues of virulence determinants such as E. coli cytotoxic necrotizing factor, Yersinia YopP, and Pseudomonas ExoT are encoded in the same region on chromosome II (28). Delivery of bacterial virulence factors into the host cell by the TTSS is contact dependent (18). Data from our laboratory suggest that the effect on barrier function does not occur in response to culture supernatants and is dependent on direct bacterial contact with epithelial cells (unpublished observations). We are currently constructing mutants with mutations in the two TTSS in both toxigenic and nontoxigenic V. parahaemolyticus strains to examine their roles in cytoskeletal and barrier function disruption.

In summary, our data demonstrate that V. parahaemolyticus potently disrupts epithelial barrier function independently of TDH or TRH production and suggest the presence of other virulence determinants. Our results extend our knowledge of V. parahaemolyticus virulence and may ultimately lead to the discovery of novel virulence factors, which could in turn enhance our ability to identify pathogenic V. parahaemolyticus strains.

Acknowledgments

We acknowledge Jennifer Liu at the Canadian Food Inspection Agency and the Alberta Provincial Laboratories for providing strains and data. We thank Emma Allen-Vercoe, Michael G. Surette, Wendy Hutchins, and Glen Armstrong for critical discussions and helpful suggestions.

Work in R.D.'s laboratory is supported by grants from the National Science and Engineering Research Council (NSERC) and the Canadian Bacterial Diseases Network. R.D. is an Alberta Heritage Foundation for Medical Research Scholar. Research in A.G.B.'s laboratory is supported by NSERC.

Notes

Editor: F. C. Fang

REFERENCES

1. Bag, P. K., S. Nandi, R. K. Bhadra, T. Ramamurthy, S. K. Bhattacharya, M. Nishibuchi, T. Hamabata, S. Yamasaki, Y. Takeda, and G. B. Nair. 1999. Clonal diversity among recently emerged strains of Vibrio parahaemolyticus O3:K6 associated with pandemic spread. J. Clin. Microbiol. 37:2354-2357. [PMC free article] [PubMed]
2. Barker, W. H., Jr., and E. J. Gangarosa. 1974. Food poisoning due to Vibrio parahaemolyticus. Annu. Rev. Med. 25:75-81. [PubMed]
3. Baumgart, D. C., and A. U. Dignass. 2002. Intestinal barrier function. Curr. Opin. Clin. Nutr. Metab. Care 5:685-694. [PubMed]
4. Bej, A. K., D. P. Patterson, C. W. Brasher, M. C. Vickery, D. D. Jones, and C. A. Kaysner. 1999. Detection of total and hemolysin-producing Vibrio parahaemolyticus in shellfish using multiplex PCR amplification of tl, tdh and trh. J. Microbiol. Methods 36:215-225. [PubMed]
5. Berkes, J., V. K. Viswanathan, S. D. Savkovic, and G. Hecht. 2003. Intestinal epithelial responses to enteric pathogens: effects on the tight junction barrier, ion transport, and inflammation. Gut 52:439-451. [PMC free article] [PubMed]
6. Bhuiyan, N. A., M. Ansaruzzaman, M. Kamruzzaman, K. Alam, N. R. Chowdhury, M. Nishibuchi, S. M. Faruque, D. A. Sack, Y. Takeda, and G. B. Nair. 2002. Prevalence of the pandemic genotype of Vibrio parahaemolyticus in Dhaka, Bangladesh, and significance of its distribution across different serotypes. J. Clin. Microbiol. 40:284-286. [PMC free article] [PubMed]
7. Centers for Disease Control and Prevention. 1998. Outbreak of Vibrio parahaemolyticus infections associated with eating raw oysters—Pacific Northwest, 1977. JAMA 280:126-127. [PubMed]
8. Cherwonogrodzky, J. W., and A. G. Clark. 1981. Effect of pH on the production of the Kanagawa hemolysin by Vibrio parahaemolyticus. Infect. Immun. 34:115-119. [PMC free article] [PubMed]
9. Chin, A. C., D. A. Teoh, K. G. Scott, J. B. Meddings, W. K. Macnaughton, and A. G. Buret. 2002. Strain-dependent induction of enterocyte apoptosis by Giardia lamblia disrupts epithelial barrier function in a caspase-3-dependent manner. Infect. Immun. 70:3673-3680. [PMC free article] [PubMed]
10. DePaola, A., L. H. Hopkins, J. T. Peeler, B. Wentz, and R. M. McPhearson. 1990. Incidence of Vibrio parahaemolyticus in U.S. coastal waters and oysters. Appl. Environ. Microbiol. 56:2299-2302. [PMC free article] [PubMed]
11. DePaola, A., C. A. Kaysner, J. Bowers, and D. W. Cook. 2000. Environmental investigations of Vibrio parahaemolyticus in oysters after outbreaks in Washington, Texas, and New York (1997 and 1998). Appl. Environ. Microbiol. 66:4649-4654. [PMC free article] [PubMed]
12. DePaola, A., J. Ulaszek, C. A. Kaysner, B. J. Tenge, J. L. Nordstrom, J. Wells, N. Puhr, and S. M. Gendel. 2003. Molecular, serological, and virulence characteristics of Vibrio parahaemolyticus isolated from environmental, food, and clinical sources in North America and Asia. Appl. Environ. Microbiol. 69:3999-4005. [PMC free article] [PubMed]
13. Fabbri, A., L. Falzano, C. Frank, G. Donelli, P. Matarrese, F. Raimondi, A. Fasano, and C. Fiorentini. 1999. Vibrio parahaemolyticus thermostable direct hemolysin modulates cytoskeletal organization and calcium homeostasis in intestinal cultured cells. Infect. Immun. 67:1139-1148. [PMC free article] [PubMed]
14. Fugino, T., R. Sakazaki, and K. Tamura. 1974. Designation of the type strain of Vibrio parahaemolyticus and description of 200 strains of the species. Int. J. Syst. Bacteriol. 24:447-449.
15. Fullner, K. J., and J. J. Mekalanos. 2000. In vivo covalent cross-linking of cellular actin by the Vibrio cholerae RTX toxin. EMBO J. 19:5315-5323. [PMC free article] [PubMed]
16. Honda, T., M. A. Abad-Lapuebla, Y. X. Ni, K. Yamamoto, and T. Miwatani. 1991. Characterization of a new thermostable direct haemolysin produced by a Kanagawa-phenomenon-negative clinical isolate of Vibrio parahaemolyticus. J. Gen. Microbiol. 137:253-259. [PubMed]
17. Honda, T., Y. Ni, T. Miwatani, T. Adachi, and J. Kim. 1992. The thermostable direct hemolysin of Vibrio parahaemolyticus is a pore-forming toxin. Can. J. Microbiol. 38:1175-1180. [PubMed]
18. Hueck, C. J. 1998. Type III protein secretion systems in bacterial pathogens of animals and plants. Microbiol. Mol. Biol. Rev. 62:379-433. [PMC free article] [PubMed]
19. Iida, T., K. S. Park, O. Suthienkul, J. Kozawa, Y. Yamaichi, K. Yamamoto, and T. Honda. 1998. Close proximity of the tdh, trh and ure genes on the chromosome of Vibrio parahaemolyticus. Microbiology 144:2517-2523. [PubMed]
20. Janda, J. M., C. Powers, R. G. Bryant, and S. L. Abbott. 1988. Current perspectives on the epidemiology and pathogenesis of clinically significant Vibrio spp. Clin. Microbiol. Rev. 1:245-267. [PMC free article] [PubMed]
21. Kaufman, G. E., M. L. Myers, C. L. Pass, A. K. Bej, and C. A. Kaysner. 2002. Molecular analysis of Vibrio parahaemolyticus isolated from human patients and shellfish during US Pacific north-west outbreaks. Lett. Appl. Microbiol. 34:155-161. [PubMed]
22. Khan, A. A., S. McCarthy, R. F. Wang, and C. E. Cerniglia. 2002. Characterization of United States outbreak isolates of Vibrio parahaemolyticus using enterobacterial repetitive intergenic consensus (ERIC) PCR and development of a rapid PCR method for detection of O3:K6 isolates. FEMS Microbiol. Lett. 206:209-214. [PubMed]
23. Kishishita, M., N. Matsuoka, K. Kumagai, S. Yamasaki, Y. Takeda, and M. Nishibuchi. 1992. Sequence variation in the thermostable direct hemolysin-related hemolysin (trh) gene of Vibrio parahaemolyticus. Appl. Environ. Microbiol. 58:2449-2457. [PMC free article] [PubMed]
24. Lang, P. A., S. Kaiser, S. Myssina, C. Birka, C. Weinstock, H. Northoff, T. Wieder, F. Lang, and S. M. Huber. 2004. Effect of Vibrio parahaemolyticus haemolysin on human erythrocytes. Cell. Microbiol. 6:391-400. [PubMed]
25. Levine, W. C., and P. M. Griffin. 1993. Vibrio infections on the Gulf Coast: results of first year of regional surveillance. Gulf Coast Vibrio Working Group. J. Infect. Dis. 167:479-483. [PubMed]
26. Ma, T. Y., D. Hollander, L. T. Tran, D. Nguyen, N. Hoa, and D. Bhalla. 1995. Cytoskeletal regulation of Caco-2 intestinal monolayer paracellular permeability. J. Cell Physiol. 164:533-545. [PubMed]
27. Madara, J. L., J. Stafford, D. Barenberg, and S. Carlson. 1988. Functional coupling of tight junctions and microfilaments in T84 monolayers. Am. J. Physiol. 254:G416-G423. [PubMed]
28. Makino, K., K. Oshima, K. Kurokawa, K. Yokoyama, T. Uda, K. Tagomori, Y. Iijima, M. Najima, M. Nakano, A. Yamashita, Y. Kubota, S. Kimura, T. Yasunaga, T. Honda, H. Shinagawa, M. Hattori, and T. Iida. 2003. Genome sequence of Vibrio parahaemolyticus: a pathogenic mechanism distinct from that of V. cholerae. Lancet 361:743-749. [PubMed]
29. Morbidity and Mortality Weekly Report. 1998. Outbreak of Vibrio parahaemolyticus infections associated with eating raw oysters—Pacific Northwest, 1997. Morb. Mortal. Wkly. Rep. 47:457-462. [PubMed]
30. Naim, R., I. Yanagihara, T. Iida, and T. Honda. 2001. Vibrio parahaemolyticus thermostable direct hemolysin can induce an apoptotic cell death in Rat-1 cells from inside and outside of the cells. FEMS Microbiol. Lett. 195:237-244. [PubMed]
31. Nishibuchi, M., A. Fasano, R. G. Russell, and J. B. Kaper. 1992. Enterotoxigenicity of Vibrio parahaemolyticus with and without genes encoding thermostable direct hemolysin. Infect. Immun. 60:3539-3545. [PMC free article] [PubMed]
32. Nishibuchi, M., and J. B. Kaper. 1985. Nucleotide sequence of the thermostable direct hemolysin gene of Vibrio parahaemolyticus. J. Bacteriol. 162:558-564. [PMC free article] [PubMed]
33. Nolan, C. M., J. Ballard, C. A. Kaysner, J. L. Lilja, L. P. Williams, Jr., and F. C. Tenover. 1984. Vibrio parahaemolyticus gastroenteritis. An outbreak associated with raw oysters in the Pacific Northwest. Diagn. Microbiol. Infect. Dis. 2:119-128. [PubMed]
34. Nusrat, A., C. von Eichel-Streiber, J. R. Turner, P. Verkade, J. L. Madara, and C. A. Parkos. 2001. Clostridium difficile toxins disrupt epithelial barrier function by altering membrane microdomain localization of tight junction proteins. Infect. Immun. 69:1329-1336. [PMC free article] [PubMed]
35. Okuda, J., M. Ishibashi, S. L. Abbott, J. M. Janda, and M. Nishibuchi. 1997. Analysis of the thermostable direct hemolysin (tdh) gene and the tdh-related hemolysin (trh) genes in urease-positive strains of Vibrio parahaemolyticus isolated on the West Coast of the United States. J. Clin. Microbiol. 35:1965-1971. [PMC free article] [PubMed]
36. Okuda, J., M. Ishibashi, E. Hayakawa, T. Nishino, Y. Takeda, A. K. Mukhopadhyay, S. Garg, S. K. Bhattacharya, G. B. Nair, and M. Nishibuchi. 1997. Emergence of a unique O3:K6 clone of Vibrio parahaemolyticus in Calcutta, India, and isolation of strains from the same clonal group from Southeast Asian travelers arriving in Japan. J. Clin. Microbiol. 35:3150-3155. [PMC free article] [PubMed]
37. Osawa, R., and S. Yamai. 1996. Production of thermostable direct hemolysin by Vibrio parahaemolyticus enhanced by conjugated bile acids. Appl. Environ. Microbiol. 62:3023-3025. [PMC free article] [PubMed]
38. Park, K. S., T. Ono, M. Rokuda, M. H. Jang, T. Iida, and T. Honda. 2004. Cytotoxicity and enterotoxicity of the thermostable direct hemolysin-deletion mutants of Vibrio parahaemolyticus. Microbiol. Immunol. 48:313-318. [PubMed]
39. Raimondi, F., J. P. Kao, C. Fiorentini, A. Fabbri, G. Donelli, N. Gasparini, A. Rubino, and A. Fasano. 2000. Enterotoxicity and cytotoxicity of Vibrio parahaemolyticus thermostable direct hemolysin in in vitro systems. Infect. Immun. 68:3180-3185. [PMC free article] [PubMed]
40. Raimondi, F., J. P. Kao, J. B. Kaper, S. Guandalini, and A. Fasano. 1995. Calcium-dependent intestinal chloride secretion by Vibrio parahaemolyticus thermostable direct hemolysin in a rabbit model. Gastroenterology 109:381-386. [PubMed]
41. Robert-Pillot, A., A. Guenole, J. Lesne, R. Delesmont, J. M. Fournier, and M. L. Quilici. 2004. Occurrence of the tdh and trh genes in Vibrio parahaemolyticus isolates from waters and raw shellfish collected in two French coastal areas and from seafood imported into France. Int. J. Food Microbiol. 91:319-325. [PubMed]
42. Sakaguchi, T., H. Kohler, X. Gu, B. A. McCormick, and H. C. Reinecker. 2002. Shigella flexneri regulates tight junction-associated proteins in human intestinal epithelial cells. Cell. Microbiol. 4:367-381. [PubMed]
43. Scott, K. G., J. B. Meddings, D. R. Kirk, S. P. Lees-Miller, and A. G. Buret. 2002. Intestinal infection with Giardia spp. reduces epithelial barrier function in a myosin light chain kinase-dependent fashion. Gastroenterology 123:1179-1190. [PubMed]
44. Sears, C. L. 2000. Molecular physiology and pathophysiology of tight junctions. V. Assault of the tight junction by enteric pathogens. Am. J. Physiol. Gastrointest. Liver Physiol. 279:G1129-G1134. [PubMed]
45. Shirai, H., H. Ito, T. Hirayama, Y. Nakamoto, N. Nakabayashi, K. Kumagai, Y. Takeda, and M. Nishibuchi. 1990. Molecular epidemiologic evidence for association of thermostable direct hemolysin (TDH) and TDH-related hemolysin of Vibrio parahaemolyticus with gastroenteritis. Infect. Immun. 58:3568-3573. [PMC free article] [PubMed]
46. Simonovic, I., M. Arpin, A. Koutsouris, H. J. Falk-Krzesinski, and G. Hecht. 2001. Enteropathogenic Escherichia coli activates ezrin, which participates in disruption of tight junction barrier function. Infect. Immun. 69:5679-5688. [PMC free article] [PubMed]
47. Simonovic, I., J. Rosenberg, A. Koutsouris, and G. Hecht. 2000. Enteropathogenic Escherichia coli dephosphorylates and dissociates occludin from intestinal epithelial tight junctions. Cell. Microbiol. 2:305-315. [PubMed]
48. Suthienkul, O., P. Aiumlaor, K. Siripanichgon, B. Eampokalap, S. Likhanonsakul, F. Utrarachkij, and Y. Rakue. 2001. Bacterial causes of AIDS-associated diarrhea in Thailand. Southeast Asian J. Trop. Med. Public Health 32:158-170. [PubMed]
49. Takahashi, A., Y. Sato, Y. Shiomi, V. V. Cantarelli, T. Iida, M. Lee, and T. Honda. 2000. Mechanisms of chloride secretion induced by thermostable direct haemolysin of Vibrio parahaemolyticus in human colonic tissue and a human intestinal epithelial cell line. J. Med. Microbiol. 49:801-810. [PubMed]
50. Taniguchi, H., H. Ohta, M. Ogawa, and Y. Mizuguchi. 1985. Cloning and expression in Escherichia coli of Vibrio parahaemolyticus thermostable direct hemolysin and thermolabile hemolysin genes. J. Bacteriol. 162:510-515. [PMC free article] [PubMed]
51. Tsukita, S., M. Furuse, and M. Itoh. 1999. Structural and signalling molecules come together at tight junctions. Curr. Opin. Cell Biol. 11:628-633. [PubMed]
52. Wu, Z., D. Milton, P. Nybom, A. Sjo, and K. E. Magnusson. 1996. Vibrio cholerae hemagglutinin/protease (HA/protease) causes morphological changes in cultured epithelial cells and perturbs their paracellular barrier function. Microb. Pathog. 21:111-123. [PubMed]
53. Wu, Z., P. Nybom, and K. E. Magnusson. 2000. Distinct effects of Vibrio cholerae haemagglutinin/protease on the structure and localization of the tight junction-associated proteins occludin and ZO-1. Cell. Microbiol. 2:11-17. [PubMed]
54. Xu, M., K. Yamamoto, T. Honda, and X. Ming. 1994. Construction and characterization of an isogenic mutant of Vibrio parahaemolyticus having a deletion in the thermostable direct hemolysin-related hemolysin gene (trh). J. Bacteriol. 176:4757-4760. [PMC free article] [PubMed]
55. Yap, A. S., J. M. Mullin, and B. R. Stevenson. 1998. Molecular analyses of tight junction physiology: insights and paradoxes. J. Membr. Biol. 163:159-167. [PubMed]
56. Yuhan, R., A. Koutsouris, S. Savkovic, and G. Hecht. 1998. Enteropathogenic Escherichia coli-induced myosin light chain phosphorylation alters intestinal epithelial permeability. Gastroenterology 113:1873-1882. [PubMed]

Articles from Infection and Immunity are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...