Ochratoxin A increases permeability through tight junctions by removal of specific claudin isoforms

John McLaughlin,1 Philip J. Padfield,1 Julian P. H. Burt,2 and Catherine A. O'Neill1

1Section of Gastrointestinal Sciences, Faculty of Medicine, University of Manchester, Salford M6 5HD; and 2School of Informatics, University of Wales, Bangor LL57 1UT, United Kingdom

Submitted 7 January 2004 ; accepted in final form 28 June 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
On interaction with the intestine, the mycotoxin ochratoxin A is know to cause rapid inflammation, diarrhea, and increased bacterial translocation. All these effects are consistent with a decrease in epithelial barrier function. However, this has not been shown directly. We determined that ochratoxin A is able to reduce the barrier properties of the model intestinal cell line Caco-2. Over 24 h, ochratoxin A reduces the transepithelial electrical resistance of Caco-2 monolayers growing on Transwell filters by ~40%. At the same time, the permeability of the monolayer is increased with respect to 4- and 10-kDa FITC dextrans, but not to 20- or 40-kDa dextrans. Immunoblotting and immuofluorescence reveal that the decrease in barrier properties is concomitant with disappearance of claudins 3 and 4, but not claudin 1 from Caco-2 cell membranes. These results suggest that ochratoxin A is able to modulate the barrier function of Caco-2 cells by removal of specific claudin isoforms.

Caco-2; intestinal permeability


THE INTESTINAL EPITHELIUM has two conflicting roles: It plays a major function in the absorption of nutrients while at the same time it maintains a barrier between the internal and external environments. This "gut barrier" is a primary means of defense against infection, preventing the translocation of potentially harmful toxins, bacteria and viruses from the gut lumen to the circulation. Where there is poor gut barrier integrity there is increased risk of systemic infection and nutritional compromise (2, 13).

The gut barrier is formed to a large extent by tight junctions (TJ). These are multiprotein complexes that link adjacent epithelial cells near their apical border. Proteins present in the TJ complexes include ZO-1, occludin, and one or more claudin isoforms. TJ seal the luminal end of the intercellular space and limit transport by this paracellular route to relatively small hydrophilic molecules.

Recent data from several laboratories have begun to elucidate the roles of individual TJ proteins. ZO-1, a member of the MAGUK family of kinases, acts as a scaffold to organize transmembrane TJ proteins and recruits various signaling molecules to the complex (1, 11). Occludin binds to ZO-1 and the actin cytoskeleton and appears to have a role in regulating permeability through the TJ (8, 24). However, numerous studies have pointed to the claudin family of TJ proteins as a key determinant of paracellular characteristics. These proteins appear to form the backbone of the TJ and to provide it with selectivity to ions (9, 10, 26, 27).

In vivo the TJ is a dynamic and highly regulated structure. Its permeability changes in response to various physiological and pathophysiological stimuli, including disease and trauma (4). It has also become clear that the integrity of the gut barrier is dependant on the presence of micronutrients within the gut. Parenteral nutrition is not sufficient to maintain the barrier, and this form of feeding can lead to bacterial translocation and increased risk of infection (7, 17, 20, 21). By contrast, enteral feeding preserves TJ integrity and maintains the gut barrier. This suggests that the gut can respond to signals generated by the presence of food within it. The particular dietary constituents required to maintain the gut barrier are as yet unidentified, and the mechanisms by which changes in junctional permeability occur are not generally understood. In addition, to date little attention has been paid to the non-nutritive constituents of food that may have adverse effects on the gut barrier.

In this study, we have examined the effects of ochratoxin A on epithelial barrier function. Ochratoxin A is a small organic toxin that is produced by many of the common fungal molds, such as Aspergillus and Penicillium (19, 22). The toxin has been found in foods such as cereals, coffee, grapes, and meat and poses a serious threat to both human and animal health. Ochratoxin A has been shown to be nephrotoxic, immunotoxic, and carcinogenic to a variety of animals (5, 12, 15). Its principal site of action is the kidneys, where it has been implicated in at least three diseases. However, upon ingestion, its primary interaction is with the gut epithelium. A single previous study by others described the effects of the toxin on transepithelial electrical resistance in cultured intestinal cell lines (18). However, it is not clear from that study whether these effects are due to increase in paracellular permeability or to membrane effects. In this study, using the intestinal cell line Caco-2, we have demonstrated that ochratoxin A alters the permeability of the epithelium by a mechanism involving removal of specific claudin isoforms from the TJ.


    MATERIAL AND METHODS
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 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. Caco-2 cells, obtained from the European Collection of Cell Cultures (ECACC no. 86010202; Salisbury, UK) were maintained in Dulbecco's modified Eagle's medium (DMEM) containing high glucose (4.5 g/l), 10% fetal calf serum, 2 mM glutamine, 1% nonessential amino acids, and 50 IU/ml penicillin-50 µg/ml streptomycin (Invitrogen, Paisley, UK). These cells (passages 41–50) were cultured in 75-cm2 T flasks (Fisher, Loughborough, UK) at 37°C in a 5% CO2 constant humidity environment with medium replaced three times weekly. Monolayers were split when they reached ~80% confluence at a 1:10 split ratio using 0.05% trypsin-0.02% EDTA.

Immunofluorescence microscopy. Caco-2 cells cultured on chamber slides (Nalge Nunc, Naperville, IL) were fixed with methanol for 20 min at –20°C. Cells were then washed three times in Tris-buffered saline (TBS), treated with 0.5% Triton-X-100 in TBS for 5 min, washed in TBS three times, and then soaked in 3% normal serum for 1 h at room temperature. Samples were incubated with the primary antibodies overnight at 4°C. These were rabbit polyclonal anti-claudin-1 (Jay.8), claudin 2 (MH44; Zymed Laboratories, South San Francisco, CA), or goat anti-claudin 3 (C-20) or 4 (C-18) (Santa Cruz Biotechnology, Santa Cruz, CA). FITC-conjugated goat anti-rabbit and rabbit anti-goat IgG (Stratech Laboratories, Cambridge, UK) were used as the secondary antibodies. Cells were then washed three times in TBS. Images were captured using confocal laser scanning microscopy (Bio-Rad MRC1024 MP confocal scanning system mounted on a Nikon Eclipse TE300 fluorescence microscope). A gallery of 30 optical sections (1 µM) through the z-plane was obtained, and composite images were processed using Confocal Assistant version 4.02 (Bio-Rad, Hercules, CA).

Measurement of transepithelial electrical resistance. Cells were seeded on Transwell polycarbonate cell culture inserts with a mean pore size of 0.4 µM (Costar) at 3 x 105 cells/cm2 and were grown for 21 days before experiment or until the transepithelial electrical resistance (TEER) had become stable. TEER was monitored using an Evometer (World Precision Instruments, Stevenage, UK) fitted with Chopstick electrodes. TEER was normalized by the area of the monolayer, and the background TEER of blank filters was subtracted from the TEER of the cell monolayer.

Treatment of cells with ochratoxin A. Cell medium was replaced with phenol red-free, fetal calf serum-free medium 24 h before the start of the experiment. Ochratoxin A (Sigma-Aldrich, Dorset, UK) was dissolved in 100 mM sodium bicarbonate (pH 7.4) The toxin was added routinely at a final concentration of 100 µM to either the apical or the basolateral side of cells growing in Transwell chambers.

MTT assay. 3-[4,5-Dimethylthiazol-2-yl]diphenyltetrazolium bromide (MTT; Sigma-Aldrich, Poole, UK) was prepared as a stock solution of 5 mg/ml in phosphate-buffered saline. Caco-2 cells were grown to 100% confluence in 96-well plates and then treated with ochratoxin A for 24 h as described above. Treatment medium was then replaced by medium containing 10% (vol/vol) MTT stock solution. Plates were incubated for 4 h at 37°C, after which the medium was replaced with dimethyl sulfoxide. Plates were shaken to dissolve the purple formazan producer, and the absorbance of each well at 570 nm was read on a Bio-Tek EL340 plate reader (Bio-Tek Instruments, Winooski, VT). Cell viability was expressed as follows: cell viability = treated wells A570/untreated wells A570.

Paracellular tracer flux assay. For paracellular tracer flux assays, FITC-labeled dextran with a molecular mass of 4, 10, 20 or 40 kDa (Sigma, Dorset, UK) was dissolved in medium at a concentration of 2 mg/ml. FITC-dextran was added at a final concentration of 0.2 mg/ml to Caco-2 monolayers growing in Transwell chambers that had been pretreated with ochratoxin A for 24 h. To evaluate the permeability of the monolayers, basal compartment media were collected after 4-h incubation with the FITC-dextran, and the amount of fluorescence in the media was measured using a Wallac 1420 Victor2 fluorimeter (Wallac, Finland). The excitation and emission wavelengths were 485 and 535 nm, respectively.

Cell extraction, SDS-PAGE, and immunoblotting. Cells were extracted by scraping into 200 µl of buffer containing NaCl (120 mM) and HEPES, pH 7.5 (25 mM), Triton X-100 (1%), EDTA (2 mM), NaF (25 mM), NaVO4 (1 mM), SDS (0.2%), aprotinin (10 µg/ml), leupeptin (10 µg/ml), and pepstatin A (10 µg/ml). The samples were incubated on ice for 30 min and then centrifuged for 15 min at full speed in a microcentrifuge. The supernatant was removed for further analysis.

SDS-PAGE was performed according to the method of Laemmli (16), and proteins were electrophoretically transferred from 12% gels onto PVDF membranes. The membranes were blocked in 5% skimmed milk and then incubated with the primary antibodies (1:1,000 dilution of rabbit anti-claudin 1, 3, or 4, mouse anti-ZO-1, or mouse anti-occludin; Zymed Laboratories). After being washed, the membranes were incubated with the secondary antibodies horseradish peroxidase-conjugated rabbit or mouse IgG as appropriate at a dilution of 1:5,000. The blots were developed using enhanced chemiluminescence (Amersham, Little Chalfont, UK) and were quantified densitometrically. Gels were scanned at 300 dpi in transmission mode using a high-resolution flat bed scanner (Epson Expression 1600) and stored in 16-bit grayscale bitmap format using a nondestructive compression algorithm tagged image file format (TIFF) to preserve image integrity. Stored bitmaps were subsequently analyzed using a program written for the MATLAB technical computing environment (Mathworks, Natick, MA).

The optical density of individual pixels within a gel image scanned in transmission or transparency mode was derived from an adaptation (Eq. 1) of the standard optical density formula

(1)
where I is the intensity of a given pixel in the scanned gel image and n is the bit resolution of the scanned grayscale image (e.g., n = 16 for a 16-bit image). Band densities were found by first selecting a region of the gel that represented a typical background intensity distribution and calculating the mean optical density of this region. Next, a rectangular region around each band was selected. For dark bands, the precise band area was extracted from this rectangular region by selecting pixels with optical densities greater than the upper 99% confidence limit of the previously measured background region. The mean background intensity was then subtracted from the selected band pixels to allow the band total optical density and optical volume of the band to be calculated by summing all band pixels and, for optical volume, calibrating this with the physical size of an individual pixel. The accuracy of this technique was verification by a comparison between the analysis of a gel using the above algorithm and a commercial densitometry system (Bio-Rad Multi-Analyst). Optical volume measurements from both methods were within 1% of each other.

Statistical analysis. Data are expressed as means ± SD. Statistical analysis was performed using the nonparametric Mann-Whitney U test, with P < 0.05 considered statistically significant. All experiments were conducted more than five times.


    RESULTS
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Ochratoxin A causes a reduction in transepithelial resistance. In this study, to examine the possibility that ochratoxin A affects the barrier function of TJ, we analyzed the effects of the toxin on cultured Caco-2 cells. First, we assessed the effects of ochratoxin A on the TEER of Caco-2 cells growing on Transwell filters. Ochratoxin A was added to either the apical or the basolateral side of the Transwell chamber at 100 µM concentration, and the TEER of the monolayer was measured over a 24-h period (Fig. 1A). When the mycotoxin was added apically, a decrease in TEER was observed after 4 h (82 ± 6% of control, n = 10; triplicate points within each experiment, P < 0.03; Fig. 1A). The reduction in TEER continued in a time-dependent manner such that by 24 h, the TEER of the treated monolayers was reduced to ~60% (±7%; P < 0.04) of the control (Fig. 1A). Very similar effects on TEER were obtained when ochratoxin A was added to the basolateral side of the chamber (Fig. 1A). Furthermore, the ochratoxin A-induced reduction in TEER was dose dependent, with maximum effect occurring at 100 µM when the toxin was added apically (Fig. 1B). An almost identical dose response was obtained when the toxin was added basolaterally (data not shown). Because the toxin produced very similar effects whether added apically or basolaterally, toxin at 100 µM was applied apically in all subsequent experiments.



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Fig. 1. A: effect of ochratoxin A on the transepithelial electrical resistance (TEER) of Caco-2 cells. Caco-2 cells were plated at confluent density on 24-mm filters. Ochratoxin A (100 µM) was added to either the apical ({circ}) or the basolateral chamber ({blacktriangledown}), and the resistance was measured over time. Significant differences in TEER between treated and control cells ({bullet}) were noted after 4 h (n = 6; P < 0.03). After 24 h, the resistance of the monolayer was reduced by ~40% (±7%, n = 6; P < 0.04). B: ochratoxin A induces a dose-dependent decrease in the TEER of Caco-2 cells. Caco-2 cells were plated at confluent density on 24-mm filters. Ochratoxin A was added to either the apical or the basolateral chamber at the concentrations indicated. TEER was measured 24 h after addition of the toxin and expressed as a percentage of the initial resistance of the monolayer. The mycotoxin has its maximum effect at 100 µM concentration. C: ochratoxin A increases the permeability of Caco-2 monolayers to FITC-labeled dextrans. Caco-2 monolayers were treated with 100 µM ochratoxin A for 24 h. After this time, 4-, 10-, 20-, or 40-kDa FITC-labeled dextran was added to the apical side of the chamber, and flux to the basolateral chamber was measured after 4 h. Ochratoxin A caused an ~3-fold increase in the paracellular transport of 4-kDa dextrans (P < 0.006, n = 6) in treated cells (shaded bar) vs. control (closed bar) cells. The paracellular pathway was also available to 10-kDa dextrans in treated but not control cells (P < 0.002). However, the paracellular pathway was not available to 20- and 40-kDa dextrans in either control or treated cells (data not shown).

 
To eliminate the possibility that cell death was causing the observed effects on TEER, the effect of 100 µM ochratoxin A on the viability of confluent Caco-2 cells was examined using the MTT assay. However, this concentration of toxin had no effect on cell viability up to 24 h after treatment (cell viability = 99.8% of untreated cells, n = 6; data not shown). However, longer exposure to the toxin (>48 h) resulted in an ~10% decline in cell viability as judged by MTT assay. By 72 h, the majority of the cells had lifted from the filters (data not shown).

We next investigated whether ochratoxin A-induced effects on TEER were reversible by washing the cells three times in PBS at the end of the experiment. However, recovery of the TEER was not possible after 24 h of exposure. Interestingly, recovery of the TEER was possible only after very short (≤2 h) exposure to the toxin. Under these conditions, cells would subsequently recover 85% (±8%, n = 5) of the control TEER within 12 h of removal of the toxin (data not shown).

Ochratoxin A increases the permeability of Caco-2 cells to FITC-labeled dextrans. Changes in TEER are not always indicative of alterations in epithelial barrier function and can often be explained as alterations in the transcellular permeability of ions. To eliminate this possibility, we studied the flux of the membrane-impermeant paracellular tracer FITC-labeled dextran across Caco-2 monolayers. As shown in Fig. 1C, ochratoxin A added to the apical side of the cells caused an approximately threefold increase in the paracellular flux of 4-kDa dextran (n = 6 with duplicates within each experiment; P < 0.006). Untreated Caco-2 monolayers are not readily permeable to 10-kDa dextrans; however, upon treatment with the mycotoxin, the monolayer became significantly more permeable to this species (Fig. 1C, n = 6; P < 0.002). The paracellular pathway in Caco-2 cells was not found to be available to FITC-labeled dextrans of either 20 or 40 kDa in either the presence or absence of ochratoxin A (data not shown). These findings demonstrate that ochratoxin A is able to modulate the paracellular pathway in Caco-2 cells.

Ochratoxin A removes specific claudin isoforms from the tight junction. We next examined whether the decrease in TEER and increase in permeability observed were due to effects of ochratoxin A on specific TJ proteins. First, we examined which claudin isoforms are expressed in Caco-2 cells. A total cell lysate was prepared from Caco-2 cells grown to confluence on a Transwell filter, and the lysate was subjected to SDS-PAGE followed by immunoblotting using antibodies specific to claudin 1, 2, 3, or 4. As shown in Fig. 2A, in the total cell lysate of Caco-2 cells, claudins 1, 3, and 4 but not claudin 2 were detected by immunoblotting as bands of the expected molecular mass (~22 kDa). To confirm the immunoblotting results and also to analyze the cellular distribution, we obtained a z series of confocal images of Caco-2 cells stained for claudin 1, 2, 3, or 4. Claudins 1, 3, and 4 were appropriately localized in a characteristic chicken wire pattern (11) consistent with their distribution in TJ (Fig. 2B). However, claudin 2 could not be detected in Caco-2 cells (Fig. 2B). Therefore, to control for the antibody, Madin-Darby canine kidney (MDCK) type II cells were stained for claudin 2. In agreement with previous work by others (25), claudin 2 produced the typical chicken wire pattern of staining in MDCK type II cells (Fig. 2C).



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Fig. 2. Expression of claudins by Caco-2 cells. A: Caco-2 cells growing on Transwell filters were scraped into buffer containing detergent. This total cell lysate was subjected to Western blot analysis using antibodies for claudin isoforms 1 (Cld 1), 2 (Cld 2), 3 (Cld 3), and 4 (Cld 4). Caco-2 cells express isoforms 1, 3, and 4 but not isoform 2. B: immunofluorescence microscopy confluent cultures of Caco-2 cells were stained with antibodies for claudins 1 (Cld 1), 2 (Cld 2), 3 (Cld 3), and 4 (Cld 4). In agreement with the immunoblotting data, immunofluorescent staining of Caco-2 shows that these cells express claudins 1, 3, and 4 but not 2. C: claudin 2 expression can be detected in Madin-Darby canine kidney (MDCK) cells.

 
Next we examined the effects of ochratoxin A on the expression and localization of TJ proteins. Caco-2 cells growing on Transwell filters were treated with or without 100 µM ochratoxin A for 24 h, at which time cells were harvested. Equal amounts of protein from control and toxin-treated samples were analyzed by performing SDS-PAGE followed by immunoblotting (Fig. 3). When the signals obtained were compared, it was noted that that there was a reduction in the signal for claudin 3 (Fig. 3) and claudin 4 in treated cells compared with untreated cells. In contrast, no reduction in claudin 1 was observed in the toxin-treated cells. (Fig. 3). We also examined the influence of ochratoxin A on the expression of ZO-1 and occludin. The level of both proteins was found to be unchanged by treatment of the cells for 24 h (data not shown). Densitometric analysis of the immunoblots revealed that the signal for claudin 3 was reduced by ~87% (±10%, n = 7; P < 0.04) in ochratoxin-treated vs. control cells. The signal for claudin 4 was also reduced, but to a lesser extent, and was ~72% (±8%; P < 0.05) lower in treated than in untreated cells.



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Fig. 3. Ochratoxin A removes specific claudins from the membrane. Caco-2 monolayers were treated with 100 µM ochratoxin A for 24 h. At this time, the cells were scraped into buffer containing detergent, and the cell lysate was analyzed using SDS-PAGE followed by immunoblotting with antibodies for claudins 1, 3, and 4. The signal for claudin 1 (cld-1) remained unchanged after treatment with the toxin. However, the signal for claudin 3 (cld-3) was reduced by ~87% (±10%; P < 0.04) in treated (T) vs. control cells (C). The signal obtained for claudin 4 (cld-4) was also reduced by treatment with the toxin and was ~72% (±8%; P < 0.05) lower in treated cells than in control cells.

 
When ochratoxin-treated Caco-2 cells were analyzed by confocal microscopy, a large reduction in the intensity of the staining for claudins 3 and 4 was observed (Fig. 4). Z scans of the cells did not reveal the appearance of the signal in other cellular areas, suggesting that claudins 3 and 4 had disappeared from the cell rather than being relocalized. In agreement with the immunoblotting results, claudin 1 staining was unchanged by treatment of the cells with ochratoxin. In addition, the overall morphology of toxin-treated cells was unchanged (Fig. 4).



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Fig. 4. Confocal laser scanning microscopy. Caco-2 cells grown to confluence on chamber slides were treated for 24 h with 100 µM ochratoxin A. The cells were then fixed and stained with antibodies for claudin 1, 2, 3, or 4. In treated cells (A), the signals for claudin 3 (Cld3) and claudin 4 (Cld4) are very much reduced in intensity compared with control cells (B). The signal for claudin 1 (Cld1) is unchanged in treated and control cells.

 
To determine whether reduction in specific claudin isoform expression was concomitant with decreases in TEER, we treated cells with 100 µM ochratoxin A for 4, 8, 12, 16, or 24 h. At these time points, the cells were harvested and subjected to immunoblotting with antibodies specific for claudin 3 or 4, and the immunoblots were quantified by densitometry. Interestingly, these results demonstrated temporal differences in the way that each claudin species disappeared from the cells. Immunoblotting revealed that even after only 4-h exposure to the toxin, much of the signal for claudin 3 had disappeared; it was measured as ~29% of the control value (±9%, n = 7; P < 0.05; Fig. 5). Levels then continued to decrease very slowly over the remaining time, such that by 24 h, only 13% (±10%, P < 0.05) of the original signal remained (Fig. 5). By contrast, the signal for claudin 4 declined much more slowly with time. After 4 h posttreatment, the claudin 4 signal was still ~80% (±7%; P < 0.05) of the control, with a gradual further reduction of the signal to ~28% (±8%) of control levels (Fig. 5).



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Fig. 5. Claudin 3 is lost from cells within 4 h of exposure to ochratoxin A. Caco-2 monolayers growing on Transwell filters were treated with ochratoxin A for 4, 8, 12, 16, or 24 h. At each time point, the cells were scraped into buffer containing detergent and the lysate was analyzed by Western blotting using antibodies specific for claudin 3 or 4. The signal for claudin 4 ({bullet}) underwent a gradual decrease with time. By contrast, the signal for claudin 3 ({circ}) underwent a more rapid decrease to ~29% (±9%; P < 0.05) within 4 h of exposure to the toxin.

 

    DISCUSSION
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 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
Ochratoxin A is a mycotoxin that is a common contaminant of cereals and animal feed. In vivo, the effects of ochratoxin A on the intestine have been demonstrated to include rapid inflammation, acute diarrhea, and increased bacterial translocation (6, 14, 23). All of these effects are consistent with increased permeability of the intestine.

Previously, it was shown by others that ochratoxin A decreased the TEER of Caco-2 cells (18). However, the mechanism underlying this observation was not explored. In particular, it was not clear whether reduction in TEER was due to effects on the TJ barrier properties or to plasma membrane effects such as differences in transcellular ion transport. The main finding from this study is that ochratoxin A is able to reduce the barrier function of Caco-2 monolayers. The reduction in barrier properties is evident on the basis of two measures. In addition to a time-dependent reduction in TEER, ochratoxin A is able to increase the permeability of Caco-2 cells to paracellular tracers of 4 and 10 kDa. However, the barrier is still preserved with respect to dextrans with greater molecular mass (20 and 40 kDa). This indicates that the toxin does not simply destroy the TJ complex but can effect a "loosening" of the TJ that facilitates the passage of smaller molecules through the paracellular pathway.

The mechanism by which ochratoxin A reduces the barrier properties of Caco-2 cells appears to be via removal of two specific claudin isoforms, 3 and 4, from the TJ. This is clear from our observation that reduction in TEER follows a time scale comparable to that of the disappearance of claudins 3 and 4 from the cells. The levels of other TJ proteins, ZO-1, occludin and claudin 1 stay constant over this time period, which suggests that claudins 3 and 4 may have a central role in TJ barrier function in Caco-2 cells.

The mechanism by which ochratoxin A is able to remove specific claudins from the TJ is at present unclear. The toxin is known to be an inhibitor of protein synthesis in other systems (18). However, the observation that levels of claudins 3 and 4 are depressed, while the levels of other TJ components remain constant, would tend not to favor a general inhibition of protein or RNA synthesis by the toxin. Similarly, it is difficult to explain the effects in terms of general cell damage, because even after 24-h exposure to the toxin, the barrier is still ~60% intact, as judged by TEER, and is still able to discriminate small from large paracellular probes. Interestingly, similar results regarding the barrier have been observed with enterotoxin A from Clostridium perfringens (25). The COOH-terminal half of this peptide toxin is able to remove claudins 3 and 4 from TJ strands in MDCK cells, resulting in decreased TEER and increased permeability. The underlying mechanism appears to be direct binding of the toxin to claudin isoforms. However, in these studies, the C. perfringens toxin did not induce temporal differences between removal of claudin 3 and 4, suggesting a different mechanism of action. When we examined a z series of cells treated with ochratoxin and stained with anticlaudin antibodies, we noted a large reduction in the intensity of the junctional staining for claudins 3 and 4 but not for claudin 1. We did not see relocalization of the fluorescent signal to other cellular areas, suggesting that claudins 3 and 4 had disappeared from the cell. This observation is in keeping with the immunoblotting results, which demonstrate a reduction in the levels of claudins 3 and 4 but not claudin 1. The overall morphology of the cells was unchanged by ochratoxin treatment, suggesting specific effects on claudins rather than gross changes to the cells. However, why only claudins 3 and 4 are affected is still a subject for debate.

Ochratoxin A displays similar kinetics, regardless of to which side of the membrane it is added. This indicates that both apical and basolateral membranes are equally susceptible to the effects of the toxin. This is most likely explained by the observation that ochratoxin A is able to diffuse through the Caco-2 cell membrane by virtue of its physicochemical characteristics and is accumulated within the cells (3). Certainly, in our experiments, it was impossible to reverse the effects of the toxin by washing if the cells had been exposed for >2 h. Shorter exposures followed by washing of the cells resulted in the reestablishment of the TEER within ~12 h (data not shown). This would tend to suggest that the toxin is accumulated within the cells. At the end point of our studies (24 h), the cells were still viable as judged by MTT assay, and only minor changes in morphology were observed. However, very prolonged (>72 h) exposure to the mycotoxin resulted in the cells lifting from the filter, presumably due to other toxic effects (data not shown).

Overall, our data demonstrate for the first time that ochratoxin A is able to induce a decrease in the barrier function of Caco-2 cells and is associated with the removal of specific claudins from the TJ. This observation may help to explain, at a molecular level, some of the in vivo effects of the toxin on the intestine. This study is also a further demonstration of the importance of the claudin family of proteins in the maintenance of epithelial barrier properties.


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 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
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This work was supported by The Wellcome Trust Grant 060834/Z/00/Z and The Digestive Disorders Foundation Grant RA/3, North Manchester Fund.


    ACKNOWLEDGMENTS
 
We are grateful to all members of our department for helpful discussions. Thanks are also due to Dr. R. Watson (Department of Dermatology, University of Manchester, Manchester, UK) for expert help with immunostaining studies and to Robert Fernandez of the University of Manchester Confocal Microscopy Unit, who helped with image capture.


    FOOTNOTES
 

Address for reprint requests and other correspondence: C. A. O'Neill, Section of Gastrointestinal Sciences, Univ. of Manchester Faculty of Medicine, Clinical Sciences Bldg., Hope Hospital, Salford M6 5HD, United Kingdom (E-mail: coneill{at}fs1.ho.man.ac.uk)

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


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