Protective role of HSP72 against Clostridium difficile toxin A-induced intestinal epithelial cell dysfunction

Tom S. Liu1, Mark W. Musch1, Kazunori Sugi1, Margaret M. Walsh-Reitz1, Mark J. Ropeleski1, Barbara A. Hendrickson3, Charalabos Pothoulakis2, J. Thomas Lamont2, and Eugene B. Chang1

1 The Martin Boyer Research Laboratories of the Inflammatory Bowel Disease Research Center, Department of Medicine, The University of Chicago, Chicago, Illinois 60637; 2 Division of Gastroenterology and Department of Surgery, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, Massachusetts 02215; and 3 Department of Pediatrics, The University of Chicago, Chicago, Illinois 60637


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We determined whether the cytoprotective heat shock protein HSP72 protects against the injurious effects of Clostridium difficile toxin A (TxA) on intestinal epithelial cells. Colonic epithelial Caco-2/bbe (C2) cells were stably transfected with HSP72 antisense (C2AS) or vector only (C2VC), resulting in low and high HSP72 expression, respectively. Measurements of epithelial barrier integrity, mitochondrial function, and apoptosis activation were assessed after TxA exposure. HSP72 and RhoA interactions were evaluated with immunoprecipitations. In C2AS cells, TxA was associated with a greater decrease in transepithelial resistance (TER), an increase in [3H]mannitol flux, and increased dissociation of perijunctional actin. Although HSP72 binds RhoA, it failed to prevent RhoA glucosylation. TxA caused a more rapid decrease in ATP, release of cytochrome c, and activation of caspase-9 in C2AS cells. To determine whether ATP depletion decreases TER, we treated cells with antimycin A, which caused a decline in TER. We conclude that HSP72 may protect intestinal epithelial cells from TxA-mediated damage through several mechanisms, including actin stabilization, mitochondrial protection, and inhibition of apoptosis activation, but not by prevention of RhoA glucosylation.

bacterial toxin; caspase; cytochrome c; heat shock proteins; transepithelial electrical resistance


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

AN ESSENTIAL FUNCTION of the intestinal epithelium is to maintain a selective barrier through which nutrients and electrolytes can permeate while excluding potentially harmful agents. The maintenance and regulation of barrier function is critically dependent on the actin cytoskeleton and an intricate cascade of intracellular mediators. The actin cytoskeleton is found in two major structures: basal stress filaments serving as scaffolding and cortical actin providing lateral support at the perijunctional rings (1). Destabilization of the actin cytoskeleton results in compromised barrier function reflected by decreased transepithelial resistance (TER) and increased paracellular flux of water and electrolytes (26).

To breach the intestinal epithelial barrier, pathogenic bacteria have evolved with potent toxins that specifically target critical elements that have an important role in maintaining barrier function. As an example, Clostridium difficile, an anaerobic pathogen, secretes two toxins, A and B, capable of causing epithelial damage and cell death (11). Toxins A and B monoglucosylate and inactivate a group of small molecular weight GTP-binding proteins called Rho (ras homologous proteins) (17). Rho proteins are believed to play an important role in regulating and maintaining the perijunctional actin ring of polarized epithelial cells (2, 3, 13, 23, 32). However, Rho inactivation may not be the exclusive mechanism of C. difficile toxins. Toxin A damages mitochondria, activates p38 MAP kinase pathway, and causes increased expression of the proinflammatory cytokine IL-8 in a Rho-independent fashion (14, 15, 42).

To counteract potentially harmful or toxic effects, cells possess intrinsic mechanisms to protect cellular functions and enhance survival. For instance, the induction of heat shock proteins (HSPs), a highly conserved family of molecular chaperones, appears to be an important response to enhance cell survival in a hostile environment (9, 24). HSP72 is a major inducible heat shock protein of epithelial cells that protects against a variety of injurious agents such as thermal injury, monochloramine (oxidant injury), heavy metals, and ischemia (9, 23, 29, 36, 44). This effect can be mitigated by stabilization of cell proteins, protection of certain cytoskeletal elements, and inhibition of the apoptotic cascade.

Because toxin A has well-characterized effects on the epithelial processes and functions, we examined whether the expression of HSP72 in intestinal epithelial cells would counteract the effects of toxin A on glucosylation of RhoA, damage to the mitochondria, loss of epithelial barrier integrity, and initiation of proapoptotic proteins. Human colonic Caco-2/bbe (C2) cells were studied because they form polarized and differentiated epithelial monolayers when grown on semipermeable supports (34). Unlike nontransformed intestinal cells such as IEC-18, C2 cells express high levels of HSP72 under basal conditions (29). Therefore, the effects of toxin A on intestinal epithelial barrier function were determined in C2 cells stably transfected with HSP72 antisense (C2AS) or vector-only cDNA (C2VC) exhibiting negligible and high levels of HSP72, respectively.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Toxin A purification and cell culture. Toxin A was purified to homogeneity as previously described (35). The human colonic epithelial Caco-2/bbe subclone (C2) (34) was grown in Dulbecco's modified Eagle's medium (high glucose, 4.5 g/l) with 10% (vol/vol) fetal bovine serum, 10 µg/ml transferrin, 50 µg/ml streptomycin, and 50 U/ml penicillin. Cells were used between passages 50 and 65.

Construction of HSP72 antisense cells. C2AS and C2VC cells have been previously described (31). The inhibition of HSP72 production was confirmed in all experiments by Western blotting, as previously described, using the C92 monoclonal anti-HSP72 antibody (Stressgen, Victoria, BC, Canada) specific for the inducible form. The expression of HSP72 was checked routinely in the cells, and in all cases the amount of HSP72 expressed by the antisense cells was <7% of the HSP72 expressed by the vector-transfected cells.

Transepithelial resistance and paracellular flux. For measurements of TER or paracellular mannitol fluxes, cells were grown on collagen-coated Transwell filters (6.5-mm diameter, polycarbonate membrane, 0.4-µm pore size; Costar, Corning, NY). To ensure stable resistance measurements, an EVOM epithelial voltohmmeter (Millipore, Bedford, MA) was used to measure TER 1 h before and immediately prior to addition of toxin A to the apical compartment. Only those cells that exhibited >200 Omega  · cm2 were used. We performed [3H]mannitol fluxes from the apical to the basolateral compartment as a specific demonstration of paracellular permeability. [3H]mannitol was added to the apical side of the Transwell (1 µCi/ml) with fluorescein-labeled dextran (3 kDa) as a marker of monolayer integrity. Cells were incubated with toxin A (30 ng/ml), with addition of the radioactive marker after 5.5 h. After 6 h, 10 µl of the apical and 20 µl of the basolateral compartment bathing media were removed. Samples were taken from the basolateral compartment every 15 min (up to 1 h) with an additional sample (apical) taken with the last set to ensure that the specific activities of the markers had not changed. Samples were first analyzed in a Hitachi F-2000 spectrofluorometer to determine fluxes of dextran, and then scintillation fluid was added. [3H]mannitol in the identical samples was then analyzed by liquid scintillation spectroscopy.

Immunofluorescence of actin cytoskeleton. C2 cells grown to confluence on glass coverslips were treated with toxin A (30 ng/ml) for 6 h and fixed in paraformaldehyde, using a pH shift method to preserve three-dimensional structure of the cells (5, 31). Briefly, cells were washed with K-PIPES buffer (80 mM K-PIPES, pH 6.5, 5 mM EDTA, and 2 mM MgCl2) at 37°C. Cells were fixed for 5 min in 3.75% (vol/vol) ethanol-free formaldehyde in K-PIPES buffer and then fixed for a second time with 3.75% formaldehyde in 100 mM NaB4O7, pH 11.0. They were subsequently permeabilized with 0.1% (vol/vol) Triton X-100 and then incubated in a 1:50 dilution of a 1 mg/ml stock of Alexa 488-phalloidin for 60 min at 37°C. After staining, slides were washed in saline and coverslips were mounted on the slides using a 1:1 solution of saline and DABCO (10 mg/ml). Slides were imaged by using an Olympus confocal microscopy system with Fluoview 2.1 software (Digestive Disease Research Core Center, The University of Chicago, Chicago, IL).

HSP72 and RhoA immunoprecipitation. Twenty-four hour confluent cells were grown on plastic culture dishes. Cells were treated with or without toxin A for up to 10 h or were heat shocked (42°C, 1 h) and harvested 60 min later. After removal of media, cells were washed with ice-cold PBS (1×) and scraped. All procedures were performed at 4°C or with samples in ice. Cell suspensions were centrifuged at 5,000 g for 10 s. Cell pellets were resuspended in 200 µl of buffer A [1% (vol/vol) Triton X-100, 1% (wt/vol) BSA, 0.01 M Tris, pH 8.0, 0.14 M NaCl, 2 mM PMSF, and 10 µg/ml aprotinin]. Samples were rotated for 1 h and spun at 100,000 g for 5 min. Supernatants were removed and placed in new tubes with 50 µl of Pansorbin cells [50% (vol/vol) slurry; Calbiochem, La Jolla, CA]. Samples were rotated for 1 h and spun at 10,000 g for 30 s. The supernatants were removed and added to 1 µg of polyclonal RhoA antibody (Santa Cruz Biotechnology, Santa Cruz, CA), 1 µg of anti-HSP72 (C92, described above for Western analysis), or 1 µg of polyclonal anti-pan ras antiserum (Upstate Biotechnology, Lake Placid, NY) coupled to protein A-Sepharose beads using the Seize-X kit (Pierce, Rockford, IL) and rotated overnight. Samples were spun at 5,000 g for 15 s and washed three times with buffer B (buffer A with Triton reduced to 0.1% and BSA reduced to 0.1%). Pellets were resuspended in Laemmli stop buffer containing 2-mercaptoethanol. Samples were resolved on 10 or 12.5% SDS-PAGE and transferred to a polyvinylidene difluoride membrane in 1× Towbin buffer. Blots were blocked in 5% (wt/vol) nonfat dry milk in Tris-buffered saline (TBS: 10 mM Tris, pH 7.4, 140 mM NaCl, and 5 mM KCl) with 0.5% (vol/vol) Tween 20 (TBST). Blots were incubated with the monoclonal anti-HSP72 antibody, the polyclonal anti-RhoA, or a monoclonal anti-pan ras (clone F132; Santa Cruz Biotechnology) overnight. Blots were washed five times in TBST, incubated with peroxidase-conjugated secondary antibody in TBST, and then washed five times in TBST and once in TBS. Detection was performed using an enhanced chemiluminescent system (Supersignal; Pierce). Additional Western blots using the same RhoA antibody confirmed the immunoprecipitation of RhoA.

ADP-ribosylation assay. To assay the effect of toxin A on RhoA, we used a Clostridium botulinum C3 exoenzyme ADP-ribosylation assay (10, 17). Treatment with toxin A causes a glucosylation of threonine-37 of RhoA, which prevents subsequent ADP-ribosylation of asparagine-41 of RhoA by C3 exoenzyme (17). Cells (C2AS and C2VC) were plated onto dishes and treated with toxin A (30 ng/ml) after 24-h confluence. At varying time intervals, cells were scraped off the dish, pelleted, and resuspended in 100 µl of lysis buffer [20 mM Tris, pH 7.4, 3 mM MgCl2, and 1 mM EGTA, with the Complete protease inhibitor cocktail (Roche Molecular, Indianapolis, IN) and 1% (vol/vol) Triton X-100]. Cells were allowed to lyse on ice for 10 min. Protein concentrations in the samples (sonicated to promote solution) were measured by the bicinchoninic acid (BCA) procedure (38). Samples (10 µg) were diluted into assay buffer, and C3 exoenzyme and [32P]NAD (5 µCi per reaction) were added to a final volume of 25 µl. Reactions were allowed to proceed 1 h at 37°C. The reaction was stopped by addition of 12 µl of 3 × Laemmli stop solution while heating to 65°C for 10 min. Proteins were separated on 12.5% SDS-PAGE, dried, and processed for autoradiography to identify 32P-labeled ADP- ribosylated RhoA.

ATP assay. ATP levels in the C2 cells were measured using a luminescent technique (7). The levels of ATP were measured by using a reagent mixture of firefly luciferase and luciferin, formulated to provide a time-independent light output (Roche Molecular). Cell samples were harvested from plates of cell cultures (C2AS and C2VC) after incubation with toxin A (30 ng/ml) or antimycin A (1 or 10 µM) for various times. Cells were scraped while in PBS, washed twice with ice-cold PBS, and treated with 2% (wt/vol) trichloroacetic acid (TCA) on ice for 20 min to extract protein. TCA-precipitated proteins were solubilized with 1 N NaOH and used for protein determination. The ATP in the supernatant was measured in a luminometer (EGG Berthold, Bad Wildbad, Germany) by using a standard curve generated for each set of samples with freshly prepared Tris-ATP. All reactions were performed in a total volume of 200 µl. The ATP per sample was calculated as nanomoles of ATP per milligram of protein.

Isolation of mitochondria and cytosol for cytochrome c analysis. Mitochondria were isolated from control and from cells treated with toxin A (30 ng/ml) for varying times, as previously described, using a Percoll-metrizimide gradient (39). Cell lysates were centrifuged at 100,000 g for 15 min to yield a cytosolic fraction from the supernatant. Both mitochondrial and cytosolic fractions were immediately resolved on 12.5% SDS-PAGE and analyzed for cytochrome c by Western blotting, as described above for HSP72 but using a monoclonal anti-cytochrome c antibody (Transduction Labs, Lexington, KY).

Caspase-9 activity. Cytosolic extracts from toxin A-stimulated cells were isolated as described for analysis of cytochrome c release. Extracts were immediately assayed for caspase-9 activation using a fluorometric assay and the substrate Ac-Leu-Glu-His-Asp-AMC. A final concentration of substrate of 50 µM was used per manufacturer's directions (Upstate Biotechnology). Ten micrograms of protein were used per assay condition. The fluorescent product AMC was measured in a Hitachi F-2000 spectrofluorometer at 460 nm. Caspase-9 activities were expressed as picomoles of AMC generated per hour per milligram of protein.

Statistical methods. Differences between data points were determined using the Instat software (GraphPad, San Diego, CA). Differences between single points were determined using the Student's t-test. For multiple comparisons, an analysis of variance using a Bonferroni correction was used.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Decreased HSP72 expression results in increased toxin A-induced changes in barrier function. Like many other neoplastic cells, C2 cells express high basal levels of HSP72, a condition that was exploited in these studies to determine whether HSP72 expression mitigates the toxic effects of toxin A. By establishing stable HSP72 antisense-transfected clones (C2AS), HSP72 expression could be significantly reduced without affecting the constitutive heat shock HSC73 homologue (31). These clones have been previously shown to be more sensitive to the injurious effects of the physiologically relevant oxidant monochloramine (31). Vector-only (without antisense)-transfected C2 clones (C2VC) were also established at the same time, serving as controls for all subsequent described experiments because of the high endogenous HSP72 expression.

To determine the effects of toxin A on barrier function of C2 HSP72 sense and antisense transfectants, we measured TER and passive [3H]mannitol flux, as previously described (31). We determined that toxin A, at a high concentration (600 ng/ml), caused a rapid and near-complete decline in TER, which occurred within 2 h, in both C2AS and C2VC cells (data not shown). At lower concentrations (1-60 ng/ml), toxin A caused a steady decline in TER after a 4- to 6-h delay that was particularly pronounced in C2AS cells when a concentration of 30 ng/ml toxin A was used (Fig. 1). Compared with untreated controls, TER of the C2AS decreased significantly at 6 h as opposed to 8 h for the C2VC cells. At 6, 8, and 10 h after toxin A treatment, the TER decrease following toxin A treatment was significantly greater in C2AS cells compared with C2VC cells. Thus inhibition of HSP72 resulted in a nearly 50% greater rate of decline in TER after toxin A treatment.


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Fig. 1.   Transepithelial electrical resistance (TER) response of vector-only (C2VC) and HSP72 antisense (C2AS)-transfected C2 cells following toxin A (TxA; 30 ng/ml) treatment. Cells were grown on collagen-coated permeable Transwell supports for 7 days. Data are means ± SE for 4 separate experiments. In each experiment, triplicate wells were used for each determination and averaged. Statistical difference (*P < 0.05) between toxin A-treated C2AS and C2VC cells is denoted. C2AS decreased significantly (P < 0.05) from untreated paired control at 6 h; C2VC decreased significantly (P < 0.05) from its untreated control at 8 h.

To confirm that these changes in TER reflected perturbations in paracellular permeability, we performed radiolabeled [3H]mannitol flux measurements, as previously described (31). Monolayers were treated with toxin A (30 ng/ml) for 6 h, and [3H]mannitol fluxes measured over the next hour. As shown in Fig. 2, after 6 h, toxin A increased the paracellular flux of [3H]mannitol in C2AS but not in C2VC monolayers, indicating an increase in paracellular permeability in the former. The increase in monolayer permeability caused by toxin A was specific for the paracellular pathway, confirmed by lack of differences in the flux of the larger dextran marker (3 kDa) between the groups. In the C2VC cells, the flux of this larger marker was 13.1 ± 1.9 and 14.7 ± 2.1 pmol · cm-2 · h-1, with and without toxin A, respectively. In the C2AS cells, the flux was 14.1 ± 1.7 and 14.9 ± 2.3 pmol · cm-2 · h-1, with and without toxin A, respectively (n = 4 separate experiments). These data strongly suggest that the effect of toxin A is mediated by selective alteration of tight junction function and not by the compromise of the monolayer integrity.


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Fig. 2.   Fluxes of [3H]mannitol in C2AS and C2VC cells. Cells were treated with TxA (30 ng/ml) or saline vehicle and allowed to incubate for 6 h. [3H]mannitol was added to the apical medium after 5.5 h, and mannitol fluxes were initiated between hours 6 and 7. Samples were taken every 15 min during this period. Values are means ± SE for 4 separate experiments. Statistical difference (*P < 0.05) between toxin A-treated C2AS and C2VC cells is denoted.

Because other studies have reported that toxins such as Escherichia coli endotoxin and toxic shock syndrome toxin (TSST-1) can induce the expression of heat shock proteins in cell cultures (8, 19), we determined whether HSP72 expression was induced by toxin A. By Western blot analysis, at a concentration of 30 ng/ml, toxin A did not have an effect on basal HSP72 expression (data not shown).

Cells without HSP72 show early loss and dissociation of perijunctional actin and tight junction integrity after toxin A stimulation. Alterations in intestinal epithelial barrier function, including those of toxin A, may be associated with changes in the perijunctional actin ring and zonula occludens (16, 33). To determine whether HSP72 can prevent the disruptive effects of toxin A on the actin cytoskeleton, we used Alexa 488-phalloidin to label actin. At 6 h, when a significant drop in TER was observed in C2AS but not in C2VC cells, perijunctional actin was noted to be disrupted with widening of intercellular spaces in the C2AS but not in the C2VC cells (Fig. 3). No changes were observed in untreated C2AS or C2VC cells over this time. C2AS cells displayed dissociation of the perijunctional actin ring and increased intercellular separation, compared with the corresponding C2VC cells (Fig. 3). Of note, no alterations in the basal actin stress filaments were observed in treated or control C2AS or C2VC cells at these time points (data not shown). The changes in perijunctional actin are therefore consistent with the known effects of toxin A on intestinal epithelial barrier function and would explain the differences seen in TER and paracellular [3H]mannitol flux. Furthermore, our data show that HSP72-expressing C2VC cells are able to counteract the actin disrupting effects of toxin A. 


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Fig. 3.   Effect of toxin A on perijunctional actin in C2AS and C2VC cells. Cells were treated with toxin A (6 h, 30 ng/ml) or diluted methanol control and then fixed and stained for actin with Alexa 488-phalloidin as described in MATERIALS AND METHODS. Images shown are representative of those of 3 separate experiments.

HSP72 immunoprecipitates with RhoA. Toxin A glucosylates and inactivates Rho proteins, including RhoA (17). To determine whether HSP72 inhibits these actions by binding RhoA, we immunoprecipitated RhoA from cellular lysates and performed tandem Western blots for HSP72 (and RhoA) to confirm coimmunoprecipitation. As shown in Fig. 4, HSP72 coimmunoprecipitated with RhoA in lysates from control and toxin A (30 ng/ml)-treated cells. Representative immunoblots in Fig. 4, left, confirm that the anti-HSP72 (A) and anti-RhoA antibodies (D) are capable of immunoprecipitating their respective antigen proteins.


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Fig. 4.   HSP72 associates with RhoA. C2VC cells were untreated (C, control), heat shocked (HS; 42°C, 1 h), or treated with TxA (30 ng/ml for varying times) and then solubilized, and immunoprecipitations (IP) using designated antibodies were performed. Western blots (WB) of the immunoprecipitates were analyzed using designated antibodies (left). Bands for RhoA (~20 kDa) are indicated by arrows. Densitometry analyses of the blots using NIH Image 1.54 are represented by the corresponding histograms (right). In all cases, the Western blot band intensity of control cells (C) was set to 100%. Images shown are representative of those of 3 separate experiments. Histogram values represent arbitrary densitometry units relative to control and are means ± SE of 3 experiments. Statistical difference (+P < 0.01) from untreated control cells (C) is denoted.

Figure 4B shows that RhoA (arrow) coimmunoprecipitated with HSP72 when anti-HSP72 antibody was used. Similarly, when anti-RhoA was used to immunoprecipitate RhoA, HSP72 was coimmunoprecipitated (Fig. 4C). No significant differences in HSP72-RhoA interaction were noted during the 10-h incubation period with toxin A compared with untreated control cells (Fig. 4, first lane of each blot and first bar of histograms). The same procedure was performed on cell lysates following heat shock at 42°C for 1 h to determine whether potential HSP72-RhoA binding was specific for toxin A-induced stress. The effect was not specific, because HSP72 and RhoA coimmunoprecipitation could be readily demonstrated following heat shock (Fig. 4, second lane of each blot).

As shown by the corresponding densitometry histograms in Fig. 4, right, no significant changes in immunoreactive HSP72 and RhoA were observed during the 10-h incubation period with toxin A. Of note is the significant difference in the Western blot signal of HSP72 following anti-HSP72 and anti-RhoA immunoprecipitations during heat shock conditions compared with control and toxin A-treated conditions (Fig. 4, A and C, second lane). The finding is not surprising given that thermal stress is a potent stimulator of HSP72 expression (9, 24, 29). The presence of increased quantities of HSP72 during heat shock most likely contributes to a greater amount of binding that results in a stronger Western blot signal after immunoprecipitation with HSP72 and RhoA antibodies. A similar effect is not seen in Western blots of RhoA following anti-HSP72 and anti-RhoA immunoprecipitations (Fig. 4, B and D), since thermal stress is not known to increase RhoA expression.

As a negative control, ras, another small GTP-binding protein, was immunoprecipitated and analyzed for HSP72 coimmunoprecipitation (binding). No interaction between HSP72 and ras could be detected by coimmunoprecipitation (data not shown). Thus HSP72 binding is specific for RhoA and does not include all GTP-binding proteins.

Despite HSP72-RhoA binding, HSP72 does not protect HSP72 against RhoA glucosylation. We next examined whether HSP72 binding with RhoA mitigates toxin A inactivation of RhoA by blocking its glucosylation effects. After toxin A treatment (30 ng/ml), protein lysates from C2AS and C2VC cells were incubated with the C3 exoenzyme of C. botulinum, which is known to ADP-ribosylate the same Rho protein substrates as toxin A (17). It is important to note that the amount of Rho labeled by the C3 exoenzyme ADP-ribosylation assay is inversely related to the amount Rho that is glucosylated by toxin A during the initial exposure. As RhoA is glucosylated by toxin A, less is available for C3-dependent ADP-ribosylation labeling by the in vitro assay, resulting in a weaker radioactive ADP signal. Conversely, if HSP72 blocks toxin A-induced glucosylation of RhoA, more RhoA will be available for ADP-ribosylation by C3, resulting in a stronger radioactive ADP signal.

As shown in Fig. 5, a similar time-dependent rate of toxin A-stimulated glucosylation of RhoA from C2AS and C2VC cells was observed. After 4 h, there was a decrease in ADP-ribosylation signal intensity noted in both cell types, indicative of increased toxin A glucosylation. Increased glucosylation in cells stimulated with RhoA over time in intact cells is reflected by a decrease in C3-stimulated ADP-ribosylation of RhoA with the in vitro assay as progressive glucosylation blocks sites for ADP-ribosylation. Thus HSP72 binding of RhoA does not appear to block the ability of toxin A to glucosylate RhoA and cannot explain the observed protective effects of HSP72 against toxin action in colonic epithelial cells.


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Fig. 5.   ADP-ribosylation of RhoA after toxin A treatment. C2VC (A) and C2AS (B) cells were treated with toxin A (30 ng/ml). At the indicated time points, cytosolic fractions of the cells were isolated. RhoA glucosylation was assayed indirectly using the C3 exoenzyme ADP-ribosylation assay as described in MATERIALS AND METHODS. Toxin A-induced RhoA glucosylation was evident at 4 h with increased RhoA glucosylation (decreased ADP-ribosylation signal) at 8 h in both C2AS and C2VC cells. Images shown are representative of those of 3 separate experiments.

HSP72 protects against mitochondrial damage and ATP depletion. Recent studies have demonstrated that toxin A induces mitochondrial damage and ATP depletion in target cells (14, 15). Cytochrome c oxidase is found exclusively in the mitochondria and participates in ATP generation by facilitating electron transfer (4). Its release from mitochondria during injury is one of the earliest stages of programmed cell death (apoptosis), resulting in the activation of several key caspases. To determine whether HSP72 mitigates these effects, we determined the release of cytochrome c from mitochondria following toxin A treatment of C2AS and C2VC cells. Cytochrome c release from mitochondria to cytoplasm was observed after 2 h of toxin A exposure only in C2AS cells (Fig. 6B). In contrast, cytoplasmic cytochrome c leakage from mitochondria in C2VC cells did not occur until after 4-6 h (Fig. 6A).


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Fig. 6.   Effects of toxin A on mitochondrial and cytosolic cytochrome c levels. C2VC (A) and C2AS (B) cells were treated with toxin A (30 ng/ml). At the indicated time points, mitochondrial (Mito) and cytosolic (Cyto) fractions were prepared as described in MATERIALS AND METHODS. An untreated control was also analyzed after 12 h (12C). Cytosolic fractions (10 µg) and mitochondrial fractions (2 µg) were run on 12.5% SDS-PAGE. Western blots were subsequently generated and analyzed as described in MATERIALS AND METHODS. Images shown are representative of those of 3 separate experiments.

We next measured cellular ATP levels in the two groups after toxin A treatment. Cellular ATP levels are dependent on functional mitochondria, and substantial decreases in ATP concentration have been associated with compromises in barrier function (12, 41, 43). Cells were exposed to toxin A (30 ng/ml), and cellular ATP levels were measured by a luminescence assay. At baseline and without toxin treatment, there were no significant differences in the cellular ATP content of C2AS and C2VC cells (Fig. 7). In the C2AS cells, which lack HSP72 expression, a greater rate of decline in ATP content was observed (Fig. 7). After toxin A treatment, cellular levels of ATP began to decline. In C2AS cells, a significant decrease compared with untreated cells was observed within 1 h, whereas for C2VC cells, a significant decrease from untreated cells was observed at 2 h (Fig. 7). After 2 and 4 h of toxin A exposure, ATP concentrations in C2VC cells were significantly greater than those in C2AS cells (Fig. 7). With continuous toxin exposure, the damaging effects of the toxin are eventually too substantial to reverse, as evidenced by the similar ATP levels at 8 h. These observations demonstrate that HSP72 delays the mitochondrial injury caused by toxin A, but with continued exposure to toxin A, the protective effects of HSP72 were lost.


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Fig. 7.   Cellular ATP levels after toxin A treatment. C2VC and C2AS cells were exposed to TxA (30 ng/ml) or medium. At the indicated time points, medium was aspirated and cells were prepared for measurement of cytosolic ATP levels as described in MATERIALS AND METHODS. Values are means ± SE for 3 separate experiments. Statistical difference (*P < 0.05) between toxin A-treated C2AS and C2VC cells is denoted. The ATP level in C2AS and C2VC cells decreased significantly (P < 0.05) from control after 1 and 2 h, respectively.

HSP72 protects against apoptotic cascade activation of caspase-9. Release of cytochrome c into the cytosol following mitochondrial damage initiates activation of caspase-9, an important early signal in the apoptosis cascade. Subsequently, caspase-9 activates caspase-3, which results in the further activation of downstream effectors, ultimately leading to apoptosis (22). Toxin A has been noted to initiate apoptosis in a number of cell lines, although C. difficile toxin B has been more studied in this respect (2, 20, 25, 27, 37). Thus we explored whether HSP72 might offer additional protection against toxin A through the prevention of caspase-9- induced apoptosis, which would reveal another level of protection in intestinal epithelial cells.

As shown in Fig. 8, activation of caspase-9 was observed in C2AS cells as early as 6 h after toxin A (30 ng/ml) treatment, increasing further with prolonged incubation. In contrast, caspase-9 activation in C2VC cells treated with toxin A (30 ng/ml) was detectable just above baseline after 8 h (Fig. 8). Furthermore, caspase-9 activity was significantly higher in C2AS compared with C2VC cells at every measured time point (Fig. 8). These data are consistent with published reports showing that HSP70 proteins inhibit caspase-3 and cytochrome c release and procaspase-9 processing (21, 28).


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Fig. 8.   Time course of caspase-9 activation in toxin A-stimulated C2VC and C2AS cells. Cells were stimulated with toxin A (30 ng/ml), and cytosolic fractions were isolated at varying times. Caspase-9 activity was measured using a fluorometric substrate. Values shown are means ± SE for 3 separate experiments. Statistical difference (*P < 0.05) between toxin A-treated C2VC and C2AS cells is denoted.

Significance of ATP depletion in barrier alteration. The potential role of ATP depletion in causing the TER changes after toxin A was investigated using the oxidative phosphorylation inhibitor antimycin A. Antimycin A binds and inactivates cytochrome b and decreases cellular ATP. To confirm this, we used nontransfected C2 cells. After preliminary experiments to determine the range of antimycin concentrations to be used, cells were treated with 1 or 10 µM antimycin A for up to 10 h. Cells were grown on collagen-coated Transwells so that TER could be measured. At specified time points, filters were cut from the support and ATP levels measured. Antimycin A caused a concentration- and time-dependent decrease in TER (Fig. 9A). Changes in TER were paralleled by decreases in cell ATP (Fig. 9B). Both concentrations of antimycin significantly decreased TER and ATP by 30 min (Fig. 9). Though some statistical differences were noted between the two concentrations (in particular the ATP decrease), these differences were not the focus of these studies; rather, these experiments support a role for ATP in maintenance of barrier function in C2 cells. Because HSP72 slows down the rate of toxin A-induced ATP decline, this may relate to its protection of barrier function.


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Fig. 9.   Effects of antimycin A on TER (A) and ATP (B) in nontransfected C2 cells. C2 cells were grown on collagen-coated Transwells until a stable TER was achieved (8-10 days; average TER: 328 ± 14 Omega  · cm2, n = 36). Monolayers were treated with the designated concentrations of antimycin A and TER and followed over 4 h. At these times, the cell monolayers were cut from the support and cell ATP was measured as described in MATERIALS AND METHODS. Data are means ± SE for 3 separate experiments. Statistical difference (*P < 0.05; ++P < 0.01) compared with untreated control is denoted.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

This study provides substantial new insights into the mechanisms of HSP72 action in protecting intestinal epithelial cells against enterotoxigenically induced cellular dysfunction and apoptosis. Three critical but not mutually exclusive mechanisms of action were identified. The first involves the protection of toxin A-induced disruption of actin cytoskeleton and tight junction-mediated barrier function. The precise mechanisms underlying this effect are not known but could potentially involve HSP72 binding and stabilization of key cytoskeleton-associated proteins, such as alpha -actinin (preliminary unpublished data) or tight junction-associated proteins, like the zonula occludens family. Our study is consistent with recent publications demonstrating that PKC regulation of tight junction zonula occludens proteins is targeted by toxin A (6). However, this study provides little support for the HSP72 protection of RhoA, a major target of C. difficile toxin A action. HSP72-RhoA binding can be detected under basal as well as stress conditions. Moreover, this binding does not affect toxin A glucosylation of RhoA. Therefore, we speculate that the HSP72-induced stabilization of actin cytoskeleton may be occurring at points downstream of RhoA, possibly through direct actions on actin filaments or other actin-binding proteins.

The second site of action of HSP72 against toxin A appears to involve the protection of mitochondrial function. The presence of HSP72 inhibits mitochondria damage as measured by cytochrome c leakage and depletion of cellular ATP by toxin A. Though we do not know the precise nature of this inhibition, we speculate that HSP72 may be either stabilizing mitochondrial membranes or mitochondrial proteins required for structural integrity and function. The resulting inhibition of cytochrome c leakage by HSP72 may be directly relevant to the third potential site of HSP72 action, the delay and attenuation of caspase-9 activation.

Mitochondrial release of cytochrome c is one of the earliest stages of programmed cell death, activating several caspases that evoke downstream mechanisms of apoptosis. We cannot determine from the present studies whether the HSP72 effect is solely due to its inhibition of cytochrome c release or a direct action on caspase-9. Relevant to the latter possibility, HSP70 has been reported to inhibit the activation of caspase-3 and pro-caspase-9 processing (21). Moreover, we previously demonstrated that HSP72 protects against TNF as well as oxidant-mediated cell death and also TNF-stimulated activation of caspase-9 (30). Because both agents promote cytochrome c leakage and caspase-9 activation, these may be potential sites for the antiapoptotic effects of HSP72.

It is important to note that the effects of toxin A, as well as those of HSP72, may involve a number of pathways that may be interdependent or independent. In the present studies of intestinal epithelial cells, we have examined a number of functions of toxin A on TER, cellular ATP, and apoptosis cascade activation and how each is affected by modulating HSP72 expression. With respect to the TER decrease, we have presented data showing that perijunctional actin rearrangement after toxin A is mitigated by HSP72 expression. The question arises as to what is the cause of the actin rearrangement.

Decreases in cell ATP could potentially lead to changes in TER but may not be the only mechanism. In the case of the specific mitochondrial oxidative phosphorylation inhibitor antimycin A, changes in cell ATP correlate with changes in TER (Fig. 9). However, with toxin A, the time correlation between decreased TER and cell ATP is less clear. As shown in Fig. 7, toxin A-induced decreases in cell ATP can be observed within the first 4 h, well before decreases in TER (Fig. 1). It is possible that a certain ATP threshold may have to be reached before TER integrity is affected, which is perhaps achieved more rapidly with antimycin. Alternatively, this temporal discrepancy could indicate that decreases in ATP and TER caused by toxin A are independent events. However, it is important to point out that these time course comparisons are based on approximations and are subject to technical limitations. They do not provide direct evidence of causal relationships.

Another effect of toxin A is the activation of the apoptosis cascade. This may relate to effects of the toxin on mitochondrial function but could involve additional pathways. Activation of caspase-9, a key early step leading to apoptosis, occurs earlier in cells expressing low levels of HSP72 (antisense). Mitochondrial cytochrome c release is an important regulator of caspase-9 activation. Our data indicate that cytochrome c release precedes caspase-9 activation and also occurs earlier in the HSP72 antisense cells. The mechanism(s) by which HSP72 might prevent mitochondrial damage and cytochrome c release remains to be clarified.

Apart from these possible mechanisms and sites of action, HSP72 has been implicated as having antioxidant effects (29, 31). Though these effects may be small, they may contribute to the overall protective effects of induced HSP72 in C2 cells subjected to oxidants or toxin A treatment. Relevant to this possibility, an important role for effects of toxin A on oxidant generation was recently demonstrated. HT29 cells treated with the antioxidants butylated hydroxyanisole or butylated hydroxytoluene before toxin A treatment suppressed stimulated IL-8 release (15). On the basis of our findings, we believe preinduction of inducible heat shock proteins like HSP72 and HSP25/27 benefits intestinal epithelial cells in situ, particularly in preparing cells against acute effects of injurious toxins and pathogens.

Recent studies from our laboratory demonstrate that the surface colonic epithelial cells do indeed express HSP72 (18, 36). In contrast, there is little or no expression of these proteins in colonic crypt cells, goblet cells, or cells of the lamina propria. Colonic epithelial cells are in direct and continuous contact with bacterial flora and luminal antigens. It is therefore interesting and teleologically important that surface colonocytes express HSP72 and HSP25/27 physiologically. The region-specific expression of these proteins, in colonic and gastric mucosa (18, 36, 40), would further implicate them as important cytoprotective factors, because these are the two most environmentally hostile areas of the gastrointestinal tract. In normal colon, HSP72 and HSP25 expression by surface colonocytes appears dependent on luminal bacterial signals, such as the generation of short-chain fatty acids and lipopolysaccharides (18, 36). The physiological expression of colonic heat shock proteins by surface colonocytes confers increased resistance to potentially injurious agents and pathogens. In the case of C. difficile toxin A, a major causal factor of antibiotic associated colitis, antibiotic suppression of short-chain fatty acid generation or other luminal factors may reduce colonic HSP expression sufficiently, rendering the colonic mucosa more susceptible to the damaging effects of toxin A.

In conclusion, we demonstrate that C2 intestinal epithelial cell lines transfected to express low HSP72 levels are more susceptible to C. difficile toxin A-induced cell injury than cells transfected with the empty transfection vector, which still express high levels of HSP72. The expression of heat shock proteins confers to intestinal epithelial cells a physiological advantage against the stressors of their external environment. The presence of toxin A in cells without HSP72 expression compromises barrier function, through a decline in TER, increase paracellular permeability, and disruption of the actin cytoskeleton. Epithelial cells are also susceptible to mitochondrial injury with the earlier release of cytochrome c, greater and more rapid decrease in cellular ATP levels, and earlier activation of apoptosis cascade elements, like caspase-9. Induction of HSP72 during stress conditions has been shown to be protective against a wide range of injurious agents and conditions. We now report that HSP72 offers protection against a specific bacterial toxin through mechanisms not previously described, including involvement of the actin cytoskeleton, mitochondria, and apoptosis activation, but not by protecting RhoA glucosylation.


    ACKNOWLEDGEMENTS

This work was performed in the Martin Boyer Research Laboratories and supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-38510 and DK-47722, Digestive Disease Core Grant DK-42086, The Gastrointestinal Research Foundation of Chicago, and a Crohn's and Colitis Foundation of America Research Grant.


    FOOTNOTES

Address for reprint requests and other correspondence: E. B. Chang, The IBD Research Center, Dept. of Medicine (MC6084), The Univ. of Chicago, 5841 South Maryland Ave., Chicago, IL 60637 (E-mail: echang{at}medicine.bsd.uchicago.edu).

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.

First published December 21, 2002;10.1152/ajpcell.00134.2002

Received 22 March 2002; accepted in final form 11 December 2002.


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