Escherichia coli Shiga toxins induce apoptosis in epithelial cells that is regulated by the Bcl-2 family

Nicola L. Jones1,2,3, Avinash Islur1, Rizwan Haq4, Mariola Mascarenhas1, Mohamed A. Karmali2, Mary H. Perdue5, Brent W. Zanke4, and Philip M. Sherman1,2,3

1 Research Institute, Hospital for Sick Children, and Departments of 2 Molecular Microbiology and Medical Genetics and 3 Pediatrics, University of Toronto, Toronto M5G 1X8; 4 Department of Medicine, University of Toronto, Princess Margaret Hospital, and Ontario Cancer Institute, Toronto M5G 2M9; and 5 Intestinal Diseases Research Program, McMaster University, Hamilton, Ontario, Canada L8N 3Z5


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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Human intestinal cells lack globotriaosylceramide (Gb3), the receptor for Shiga toxin-1 (Stx1) and Shiga toxin-2 (Stx2). Therefore, the role of these toxins in mediating intestinal disease during infection with Shiga toxin-producing Escherichia coli is unclear. The aims of this study were to determine whether Stx1 and Stx2 induce apoptosis in epithelial cells expressing (HEp-2, Caco-2) or lacking (T84) Gb3 and to characterize the role of the Bcl-2 family. Stx1 (12.5 ng/ml) induced apoptosis in both HEp-2 (21.9 ± 7.9% vs. 0.8 ± 0.3%, P = 0.01) and Caco-2 (10.1 ± 1.2% vs. 3.1 ± 0.4%, P = 0.006) cells but not in Gb3-deficient T84 cells. Toxin-mediated apoptosis of HEp-2 cells was associated with enhanced expression of the proapoptotic protein Bax. Inhibition of caspase activation prevented toxin-stimulated apoptosis. In addition, overexpression of Bcl-2 by transient transfection blocked Stx1-stimulated cell death. These findings indicate that Shiga toxins produced by E. coli signal Gb3-expressing epithelial cells to undergo apoptosis in association with enhanced Bax expression, thereby resulting in activation of the caspase cascade.

caspases; Bax; programmed cell death


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

INFECTION WITH Shiga toxin-producing Escherichia coli (STEC) causes diarrhea and can result in the more severe sequelae of hemorrhagic colitis and the hemolytic uremic syndrome (34). The exact factors involved in disease pathogenesis are not known. However, bacteriophage-encoded Shiga-like toxins, called Stx1 and Stx2, are considered to be major virulence factors of STEC infection (27). Stx1 and Stx2 are A-B subunit toxins in which the B subunit binds as a pentamer to a specific glycolipid receptor, globotriaosylceramide (Gb3) (28). After binding to Gb3, the holotoxin is endocytosed and transported to the Golgi apparatus and then to the endoplasmic reticulum. The A subunit inhibits protein synthesis by acting on the 28S rRNA of the 60S ribosomal subunit, resulting in cell cytotoxicity (27).

The role of Stx1 and Stx2 in mediating intestinal disease is controversial (34). Studies in animal models of infection suggest that Stx can modulate disease severity, including the production of diarrhea and hemorrhagic colitis. Purified toxin induces fluid accumulation in ligated rabbit intestinal loops in association with the presence of apoptotic intestinal villus epithelial cells (20). Furthermore, infection of rabbits with a natural E. coli enteropathogen, strain RDEC-1 (serotype O15: H-) when expressing Stx1 results in more severe disease than that caused by infection with the isogenic toxin-negative strain (42). In contrast, in gnotobiotic piglets (45) and infant rabbits (26), infection with Stx toxin-positive or -negative strains results in no detectable difference in the diarrheal response.

The variation in disease in response to Stx in these animal models could be caused by the presence or the absence of the receptor for Stx (Gb3) in the intestinal epithelium (27). Because the human intestine lacks detectable Gb3 (14), it has been suggested that Stx mediates only the vascular complications of disease including hemorrhagic colitis and the hemolytic uremic syndrome (9). However, previous studies showed that Stx1 also undergoes directed translocation to the Golgi and endoplasmic reticulum in human intestinal cells deficient in Gb3 (37).

Apoptosis, a distinct form of cell death, plays a crucial role in the development and maintenance of homeostasis of tissues (17). Therefore, deregulation of the apoptotic pathway can result in pathological processes (17, 18, 49). A major molecular modulator of apoptosis is the Bcl-2 family (1). This family of proteins contains both proapoptotic members such as Bax and Bak and antiapoptotic members such as Bcl-2 (1). Although the exact mechanisms by which these proteins regulate apoptosis are unclear, the ability of the different pro- and antiapoptotic family members to form a dynamic equilibrium between hetero- and homodimers appears to regulate the apoptotic pathway (25). Recent evidence indicates that the ratio of suppressors to inducers determines the sensitivity of the target cell to an apoptotic stimulus by modulating mitochondrial cytochrome c release. For example, Bax induces the release of mitochondrial cytochrome c and triggers caspase activation. Bcl-2 and Bcl-xL are capable of preventing Bax-mediated cytochrome c release and cell death (8).

In human intestinal epithelium, the distribution of the death antagonist Bcl-2 is limited to the colonic crypts (32) whereas Bax, an apoptotic agonist, is expressed near the gut lumen (24). This topographic expression of pro- and antiapoptotic Bcl-2 family members correlates with the location of intestinal cells destined for apoptosis and proposed stem cells, respectively (17). In addition, aberrant Bcl-2 expression has been implicated in the pathogenesis of the adenoma-to-carcinoma sequence in colon carcinogenesis (3, 32). Furthermore, enhanced Bak expression in association with apoptosis of gastric epithelial cells is observed during infection with the pathogen Helicobacter pylori (4). These findings suggest that the Bcl-2 family plays a crucial role in regulating apoptosis in the intestine.

A growing list of bacterial pathogens are able to modulate apoptosis of eukaryotic cells (49). For example, Shigella flexneri induces apoptosis of infected macrophages in vitro (5) and in vivo (50), mediated by the secreted bacterial product IpaB (5). In contrast, epithelial cells are resistant to S. flexneri-stimulated programmed cell death in vitro, suggesting cell type specificity for the apoptotic signaling pathway (31). Purified Stx1 is capable of causing apoptosis in Burkitt's lymphoma cells (30), Vero cells (15) and human renal tubular epithelial cells (19). In Burkitt's lymphoma cells, the susceptibility to apoptosis is related to expression of the receptor Gb3 (30, 43). Therefore, the aims of the present study were to determine whether Stx1 and Stx2 induce apoptosis of mucosal epithelial cells, to delineate the requirement for the Gb3 receptor, and to define the regulation of apoptosis by members of the Bcl-2 family after exposure to Stx.


    MATERIALS AND METHODS
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Bacterial Toxins

Stx1 and Stx2 were purified by methods described previously (2, 36). Briefly, cell lysates were obtained from E. coli strain R82pJES 120 DH5a (for Stx2 purification) and strain JB28 (for Stx1 purification) by polymyxin B (0.1 mg/ml) treatment. After precipitation by ultracentrifugation, the preparation underwent column chromatography purification by applications to hydroxylapatite column (Calbiochem, La Jolla, CA), chromatofocusing column of Polybuffer Exchanger 94 (Pharmacia, Baie d'Urfe, PQ, Canada), and Cibachron blue F3Ga column (Pierce, Rockford, IL). The purified B subunit of Stx1 was kindly provided by Dr. Clifford Lingwood (Hospital for Sick Children, Toronto, ON, Canada).

Cell Culture

HEp-2 cells, a human laryngeal epithelial cell line expressing Gb3 (37), were grown as monolayers in tissue culture flasks at 37°C in 5% CO2. The culture medium was MEM (Life Technologies GIBCO BRL, Grand Island, NY) supplemented with 15% (vol/vol) heat-inactivated FCS (Cansera International, Rexdale, ON, Canada), 0.1% sodium bicarbonate, and 2% penicillin-streptomycin. After overnight serum starvation, cells were incubated for up to 48 h with varying concentrations of Stx1, Stx2, or the purified B subunit of Stx1, which were filtered through a 0.2-µm syringe filter (Gelman Sciences, Ann Arbor, MI). In some studies, HEp-2 cells were incubated with cycloheximide (10 µg/ml) to inhibit protein synthesis (38).

The human intestinal cell line Caco-2, which expresses Gb3 (16), was grown as a monolayer in tissue culture flasks. Cells were maintained in MEM with Earle's salts, nonessential amino acids, 25 mM HEPES, 1 mM sodium pyruvate (all from Life Technologies GIBCO BRL), and 2% penicillin-streptomycin supplemented with 10% (vol/vol) heat-inactivated FCS (Cansera International). Cells that had been incubated for 14 days were incubated with Stx1 (concentration range 12.5-113 ng/ml), which was filtered through a 0.2-µm syringe filter (Gelman Sciences).

T84 cells, a human intestinal cell line that lacks detectable Gb3 (37), were grown as monolayers in tissue culture flasks. The cells were cultured in a 1:1 mixture of DMEM and F-12 nutrient mixture (GIBCO) supplemented with 10% (vol/vol) heat-inactivated FCS and 2% (vol/vol) penicillin-streptomycin (GIBCO) and grown at 37°C in 5% CO2. Cells were treated for up to 48 h with Stx1 (concentration range 12.5-113 ng/ml).

Assessment of Apoptosis

Transmission electron microscopy. For transmission electron microscopy, cells were grown to confluence in tissue culture flasks and incubated with Stx1 or Stx2 as described in Cell Culture. Cells incubated in the absence of toxin served as controls. Both cells in suspension and trypsinized cells were pelleted, fixed with 2% glutaraldehyde in 0.1 M phosphate buffer (vol/vol), postfixed in 2% osmium tetroxide, and dehydrated through a series of graded acetone washes (7). Samples were embedded in epoxy resin, and ultrathin sections were placed onto 300-mesh copper grids. The grids were then stained with uranyl acetate and lead salts, as described previously (7). Grids were examined for the presence of apoptotic cells using a transmission electron microscope at an accelerating voltage of 60 kV (21).

Fluorescent dye staining. Cells in suspension and trypsinized cells were pelleted and resuspended in 1 ml of phosphate-buffered saline. Acridine orange-ethidium bromide in phosphate-buffered saline (100 µg/ml) was added to the suspension. A drop of the suspension was applied to a microscope slide, and apoptotic cells were assessed by fluorescent microscopy as previously described (10). The percentage of apoptotic cells was determined by counting 500 cells in multiple randomly selected fields.

Cell death detection by immunoassay. To detect oligonucleosomes formed as a result of DNA fragmentation, the Cell Death Detection ELISAplus kit (Boehringer Mannheim, Indianapolis, IN) was used as described by the manufacturer. Briefly, adherent control and Stx1-treated (12.5 ng/ml) HEp-2 cells were trypsinized and 5 × 104 cells were lysed by incubation in the supplied lysis buffer. Supernatants of cell lysates were transferred to streptavidin-coated microtiter wells, and cytoplasmic nucleosomes were quantitated by using biotinylated anti-histone antibody and peroxidase-conjugated anti-DNA antibody after incubation with the peroxidase substrate 2,2'-azino-di(3-ethylbenzthiazoline-sulfonate). An enrichment factor, to quantitate the relative increase in nucleosomes, was calculated as the ratio of specific absorbances in lysates from Stx1-treated cells compared with untreated cells. The histone-DNA complex provided by the manufacturer served as a positive control for the assay. HEp-2 cells incubated with 1 M sorbitol for 2 h at 37°C, which has previously been demonstrated to induce apoptosis (23), served as an additional positive control. Cells that underwent freeze-thawing at -20°C for 1 h served as a control for necrosis.

Western blotting. Cells were lysed in 0.15 ml of ice-cold lysis buffer containing 150 mM NaCl, 1% Triton X-100, 10 mM Tris (pH 7.4), 5 mM EDTA, 1 mM phenylmethylsulfonylfluoride, 2 µg/ml aprotinin, 50 µg/ml leupeptin, 1 mM benzamidine, 1 µg/ml pepstatin, 1 mM sodium vanadate, 50 mM sodium fluoride, and 2 mM sodium pyrophosphate for 30 min on ice (11). The lysate was centrifuged at 14,000 rpm for 15 min at 4°C. Protein concentrations of the supernatants were measured by the Bradford method, and samples were diluted with Laemmli buffer. Samples with equivalent concentrations of protein were boiled for 2 min, subjected to 14% SDS-PAGE and electroblotted onto an Immobilon-P transfer membrane (Millipore, Bedford, MA). Blots were blocked with 5% skim milk in phosphate-buffered saline plus 0.5% Tween 20 overnight at 4°C. Separate blots were then incubated with anti-Bcl-2 (2.0 µg/ml; Santa Cruz Biotechnology, Santa Cruz, CA), anti-Bak (2.5 µg/ml; Oncogene Research Products, Cambridge, MA) and anti-Bax (2.5 µg/ml; Santa Cruz Biotechnology) antibodies for 1 h at 37°C. Blots were washed in phosphate-buffered saline plus 0.5% Tween 20 and then incubated with peroxidase-conjugated anti-mouse immunoglobulin G antibodies (for Bcl-2 and Bak) or anti-rabbit antibodies (for Bax). Immunoblots were developed with the ECL Western blotting detection system (Amersham Life Science; Little Chalfont, UK). Densitometry was performed to compare protein expression between groups with the National Institutes of Health Gel Plotting Macros program. Differences in sample loading were normalized by comparing the densitometry of immunoblots to an irrelevant protein band in Coomassie brilliant blue-stained polyacrylamide gels.

Transfection of Cultured Cells

DNA used for transfection was purified by anion-exchange column (Qiagen, Valencia, CA). Subconfluent HEp-2 cells were transiently transfected in six-well Costar cell culture dishes (Corning, Corning, NY) using FuGENE transfection reagent (Boehringer-Mannheim, Laval, PQ, Canada) with pcDNA3.Bcl-2 (4 µg) (46) and cotransfected with a green fluorescent protein (GFP) marker plasmid (pEGFP C2) (Clontech Laboratories, Palo Alto, CA) at a molar ratio of 4:1. Control cells were transfected with the vector pCDNA3 (4 µg; Invitrogen, San Diego, CA) and pEGFP C2. After culturing for 48 h, the cells were incubated in the presence or absence of Stx1 (12.5 ng/ml) for an additional 24 h at 37°C. Floating and trypsinized adherent cells were then stained with Hoechst dye (5 µg/ml), and the percentage of apoptotic nuclei among the GFP-positive cells was enumerated by fluorescence microscopy (43).

Caspase Inhibition

HEp-2 cells were preincubated with the broad-spectrum caspase inhibitor benzyloxycarbonyl-Asp-Glu-Val-Asp-fluoromethyl ketone (Z-VAD-fmk) or the CPP-32 family-specific inhibitor benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl ketone (Z-DEVD-fmk) (Calbiochem) at varying concentrations (1-200 µM) for 1 h at 37°C before Stx1 treatment (10). Inhibition of apoptosis was determined by fluorescence microscopy of cells stained with acridine orange and ethidium bromide, as outlined in Fluorescent dye staining.

Statistical Analysis

Results are expressed as means ± SE. To test statistical significance between multiple groups, a one-way analysis of variance was used followed by post hoc comparisons with the Newman-Keuls test. The two-tailed unpaired Student's t-test was used where indicated.


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Effect of Stx1 on Gb3-Expressing Cells

HEp-2 cells incubated with Stx1 underwent apoptosis as assessed both by fluorescent microscopy and by transmission electron microscopy, as shown in Fig. 1. Apoptotic cells displayed the characteristic ultrastructural features of cell shrinkage, condensed and marginated chromatin, and fragmented nuclei.


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Fig. 1.   Shiga toxin-1 (Stx1) induces apoptosis in globotriaosylceramide (Gb3)-expressing HEp-2 cells. Identification of apoptotic cells in untreated (A, C) and Stx1-treated (12.5 ng/ml; B, D) HEp-2 cells after 24 h by acridine orange-ethidium bromide staining and fluorescence microscopy (A, B) and transmission electron microscopy (C, D). Untreated HEp-2 cells demonstrate normal morphology as assessed by fluorescence microscopy (A) and transmission electron microscopy (C). Stx1-treated HEp-2 cells show morphological features of apoptosis including condensed and marginated chromatin with enhanced fluorescence (arrows and arrowhead; B). Nonviable apoptotic cells fluoresce orange (arrowhead). As assessed by transmission electron microscopy (D), Stx1-treated HEp-2 cells (arrows) showed characteristic features of programmed cell death, including cytoplasmic vacuolation and condensed and marginated nuclear chromatin. Original approximate magnifications: A and B, ×400; C, ×7,800; D, ×6,000. Quantitation of dose (E)- and time (F)-dependent induction of apoptosis by Stx1 is also shown. Incubation with varying concentrations of Stx1 (range 1.25-25.0 ng/ml) resulted in increase in apoptosis compared with untreated cells at each concentration (ANOVA, P < 0.05; E). HEp-2 cells incubated with 12.5 ng/ml Stx1 (gray bars) compared with untreated cells (stippled bars) demonstrated an increase in apoptosis at each time point (ANOVA, P < 0.05; F). Results are expressed as mean percentage of apoptotic cells per 500 cells enumerated. Error bars represent SE.

DNA fragmentation and release of oligonucleosomes was detected in toxin-treated cells by use of a cell death immunoassay with an apoptotic enrichment factor (4.9 ± 1.9) comparable to both the histone-DNA complex-positive control (7.7 ± 3.5) and cells treated with sorbitol to induce apoptosis (5.2, n = 1). In contrast, the apoptotic enrichment factor did not increase in cells undergoing necrosis after freeze-thawing (0.27, n = 1).

HEp-2 cells incubated with Stx1 at varying concentrations for 24 h showed a dose-dependent increase in apoptosis (Fig. 1E), with a maximal effect detected at a dose of 12.5 ng/ml (21.9 ± 7.9%, ANOVA, P < 0.05). The Stx1-mediated apoptosis of HEp-2 cells was time dependent, with the largest increase in cell death observed after 24 h of incubation with the toxin (20.7 ± 4.7%; ANOVA, P < 0.05; Fig. 1F). After Stx1 treatment of HEp-2 cells for 48 h, a large percentage of apoptotic cells undergoing secondary necrosis was observed (67.9 ± 5.5%). Treatment of HEp-2 cells with boiled Stx1 (100°C × 30 min) did not result in enhanced cell death compared with controls, indicating that the observed effects were not caused by the presence of contaminating small amounts of lipopolysaccharide.

To determine whether Stx1 would induce programmed cell death in other Gb3-expressing epithelial cells, the human intestinal Caco-2 cell line was used. Similar to the findings with HEp-2 cells, incubation with 12.5 ng/ml of Stx1 for 24 h stimulated Caco-2 cells to undergo programmed cell death (10.1 ± 1.2% vs. 3.1 ± 0.4%; P = 0.006, Student's t-test).

Evaluation of Apoptosis in Stx1-Treated Gb3-Deficient T84 Cells

To determine whether Stx1 functions as a proapoptotic stimulus in an epithelial cell line lacking Gb3, T84 cells were incubated with the toxin. In contrast to HEp-2 and Caco-2 cells, T84 cells were resistant to Stx1-mediated apoptosis, as measured by fluorescent microscopy after staining with acridine orange and ethidium bromide. At Stx1 concentrations (113 ng/ml) 10 times as high as the dose required to achieve a maximal effect in HEp-2 cells, the percentage of apoptotic T84 cells was not increased compared with untreated control cells (0.4% ± 0.6%).

Comparison of Stx1- and Stx2-Mediated Apoptosis in HEp-2 Cells

The induction of apoptosis in Gb3-expressing HEp-2 cells and the resistance to toxin-mediated apoptosis in T84 cells suggested that Gb3 was necessary for the activation of the cell death program by the toxin. Therefore, we investigated the effects of the related Shiga toxin Stx2, which is also synthesized by STEC but which binds less avidly to the Gb3 receptor (12). Stx2-induced programmed cell death of HEp-2 cells was demonstrated by typical morphological features of cell blebbing and condensed and marginated nuclear chromatin (Fig. 2A). HEp-2 cells treated with equivalent concentrations of Stx1 and Stx2 underwent apoptosis (Fig. 2B). However, incubation with Stx1 resulted in a greater increase in cell death compared with treatment with an equivalent concentration of Stx2 (21.2 ± 4.4% and 13.4 ± 3.6%, respectively; ANOVA, P < 0.05).


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Fig. 2.   Apoptotic response of HEp-2 cells to Shiga toxin-2 (Stx2). A: transmission electron photomicrograph of HEp-2 cells incubated with Stx2 (12.5 ng/ml). Typical morphological features of apoptosis including cell membrane blebbing and marginated and condensed chromatin are demonstrated. Approximate magnification ×46,000. B: comparison of apoptotic response in HEp-2 cells exposed to equivalent concentrations of Stx1 and Stx2 (12.5 ng/ml for 24 h at 37°C). Apoptosis was induced in HEp-2 cells treated with Stx1 or Stx2, with a greater induction of cell death mediated by Stx1. Results are expressed as mean percentage of apoptotic cells per 500 cells enumerated. Error bars represent SE. ANOVA, P < 0.05.

Induction of Apoptosis With B Subunit of Stx1

To determine whether binding to Gb3 was sufficient to trigger apoptosis, HEp-2 cells were incubated with the purified B subunit of Stx1. Compared with incubation with the holotoxin, a 100-fold higher concentration of the B subunit was required to induce apoptosis (6.4 ± 0.6%; P < 0.05, ANOVA). These findings indicated that the holotoxin was more effective in transducing the cell death signal.

To determine whether protein synthesis inhibition could enhance the B subunit-stimulated cell death, we compared the degree of apoptosis in B subunit-treated cells in the presence or absence of cycloheximide (10 µg/ml). Compared with cells incubated with the B subunit alone, the addition of cycloheximide did not enhance apoptosis (6.4 ± 0.6% vs. 6.6 ± 1.1%).

Expression of Bcl-2, Bax, and Bak After Treatment With Stx1 and Stx2

To delineate the role of Bcl-2 and related family members in the toxin-mediated apoptotic cascade, expression of Bcl-2 and the proapoptotic homologs Bax and Bak were measured by Western blotting. As shown in Fig. 3, in three separate experiments after incubation with Stx1 for 24 h, there were 2.9-, 9.0-, and 3.2-fold increases in expression of the proapoptotic family member Bax compared with untreated HEp-2 cells. Similarly, cells incubated with Stx2 showed 2.9-, 6.0- and 1.9-fold increases in Bax expression. In contrast, expression of the proapoptotic family member Bak and the apoptotic antagonist Bcl-2 did not differ from that in untreated cells after toxin treatment (Fig. 3).


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Fig. 3.   Immunoblot analysis of Bcl-2 family members in response to Shiga-like toxins. A: Bax, Bak, and Bcl-2 expression in untreated HEp-2 cells and tissue culture cells incubated with 12.5 ng/ml of either Stx1 or Stx2 for 24 h. Both Stx1 and Stx2 enhanced expression of Bax (3.2- and 1.9-fold increase respectively, as measured by densitometry when normalized for protein loading) compared with untreated control cells. B: Coomassie brilliant blue-stained polyacrylamide gel corresponding to Western blots shown in A. Lane M, markers; lane C, control cells.

Prevention of Toxin-Induced Cell Death by Inhibition of Caspases

To determine whether inhibition of caspase activity affects toxin-triggered apoptosis the caspase inhibitors Z-VAD-fmk and Z-DEVD-fmk were used. As shown in Fig. 4, preincubation of HEp-2 cells with the general caspase inhibitor, Z-VAD-fmk, blocked cell death in a dose-dependent manner. The maximum inhibitory effect was observed at a dose of 50 µM (0.9 ± 0.4% vs. 25.4 ± 3.2%, P < 0.001). Higher doses of the specific caspase-3 inhibitor Z-DEVD-fmk (200 µM) were required to prevent toxin-mediated apoptosis (4.6 ± 0.6%, P < 0.05).


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Fig. 4.   Caspase inhibition prevents Stx1-stimulated cell death of HEp-2 cells. Cells preincubated with general caspase inhibitor Z-VAD-fmk showed dose-dependent reduction in apoptosis in response to Stx1 (12.5 ng/ml) (P < 0.001). Preincubation with specific caspase-3 inhibitor Z-DEVD-fmk also blocked cell death, but a higher concentration (200 µM) was required to achieve same effect (P < 0.05). Results are expressed as mean percentage of apoptotic cells per 500 cells enumerated. Error bars represent SE.

Overexpression of Bcl-2 Inhibits Stx1-Mediated Apoptosis

To confirm the biological importance of the observed alterations in expression of the Bcl-2 family during toxin-triggered cell death, HEp-2 cells were transiently transfected with constructs for overexpression of Bcl-2 together with GFP. Although transfection with the empty vector alone resulted in an increase in the baseline level of apoptosis, toxin-treated GFP-positive cells containing the empty vector demonstrated enhanced apoptosis and displayed the characteristic features of condensed and brightly stained chromatin with nuclear fragmentation. In contrast, the morphological features of apoptosis were not detected in GFP-positive cells expressing the Bcl-2 construct. As shown in Fig. 5, compared with cells transfected with the empty vector, transient transfection of HEp-2 cells with Bcl-2 prevented Stx1-induced apoptosis (48.0 ± 2.9% vs. 6.3 ± 1.5%, P < 0.05).


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Fig. 5.   Inhibition of Stx1-mediated apoptosis by overexpression of Bcl-2 in HEp-2 cells. Cells were transiently cotransfected at a molar ratio of 1:4 with green fluorescent protein (GFP) and either Bcl-2 (4 µm) or control vector (4 µm) and incubated with Stx1 (12.5 ng/ml) for 24 h at 37°C. Apoptosis of GFP-positive cells was assessed by fluorescent microscopy after Hoechst staining. Cells transfected with control vector underwent apoptosis in response to Stx1 (48.0 ± 2.9%). In contrast, apoptosis was inhibited in cells transfected with Bcl-2 (6.3 ± 1.5%; P < 0.05). Results are expressed as mean percentage of apoptotic cells per 100 GFP-positive cells enumerated. Error bars represent SE.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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This study demonstrates that Stx1 and Stx2 produced by E. coli induce apoptosis of human mucosal epithelial cells that express the Gb3 receptor for the toxins. The B subunit of Stx1 also triggers cell death of epithelial cells but to a lesser degree than the holotoxin. In contrast, intestinal epithelial cells lacking the glycolipid receptor did not undergo apoptosis. In addition, evaluating the role of the Bcl-2 family of pro- and antiapoptotic proteins in mediating programmed cell death showed that toxin treatment of Gb3-expressing cells was associated with an enhanced expression of the proapoptotic homolog Bax.

In previous studies using immunogold labeling, we showed (37) that Stx1 undergoes transcellular translocation in Gb3-negative T84 intestinal cells and is localized to organelles that are involved in retrograde transport of the toxin to both the endoplasmic reticulum and the Golgi network. Similar subcellular organellar targeting of Shiga toxin from S. dysenteriae type 1 is dependent on the Gb3 receptor (40). These findings suggest that in T84 cells there exists an alternative receptor for Stx1 binding. Indeed, a recent study demonstrated that Stx1 and Stx2 bind to an uncharacterized protein receptor on Vero cells in addition to the previously identified glycolipid receptor, Gb3 (6).

T84 cells were resistant to Shiga toxin-stimulated programmed cell death at doses up to 10 times that required to mediate a maximal apoptotic effect in HEp-2 cells, despite the ability of the toxin to undergo directed retrograde transport. Taken together, these findings indicate that Shiga toxins produced by E. coli are capable of mediating apoptosis and that the Gb3 receptor is required to activate the death signal transduction cascade. In support of this contention, the B subunit binding portion of the toxin lacking the enzymatic activity necessary to inhibit protein synthesis also was capable of inducing apoptosis in Gb3-expressing epithelial cells, albeit to a lesser degree than the holotoxin. These findings in epithelial cells are in agreement with previous studies that reported a reduced degree of apoptosis in Gb3-expressing lymphocytes treated with the B subunit compared with those cells exposed to the holotoxin (30). Fibroblast cells transfected with the Stx1 B subunit undergo apoptosis (33). These findings support the results of the present study showing that the B subunit alone was capable of triggering cell death. Together, these data suggest that factors in addition to binding of the B subunit to the Gb3 receptor mediate apoptosis.

The mechanism by which the holotoxin enhances the cell death signal triggered by Gb3 binding alone remains unclear. Inhibition of protein synthesis can potentiate an apoptotic stimulus, likely through the reduction of short-lived antiapoptotic proteins (41). Therefore, inhibition of protein synthesis by the A subunit could enhance the apoptotic cascade signaled by binding to Gb3. However, treatment of cells with the protein synthesis inhibitor cycloheximide did not enhance the apoptotic effect of the B subunit alone, a finding that suggests that an additional mechanism(s) may well be involved. Indeed, a recent study demonstrated that transfection of fibroblasts with the Stx1 A subunit does not result in apoptosis (33).

Our findings suggest that intestinal epithelial cells are resistant to the cell death cascade activated by Stx1. Whether cytokines, such as tumor necrosis factor-alpha (TNF-alpha ), that are produced in response to infection with STEC (29) alter the expression of Gb3 in intestinal cells and thereby enhance the sensitivity to apoptosis has yet to be determined. Previous in vitro studies of endothelial cells derived from human umbilical vein (35) and saphenous vein (22) demonstrated that Stx1-mediated cytotoxicity is potentiated by pretreatment with lipopolysaccharide, butyrate, and cytokines, including TNF-alpha and interleukin-1 (IL-1). TNF-alpha and IL-1 increase galactosyl transferase activity, resulting in enhanced Gb3 expression by endothelial cells (46). Similarly, butyrate pretreatment of Caco-2A or HT-29 intestinal epithelial cells promotes Stx1 binding and cytotoxicity (16). In contrast, butyrate treatment of T84 cells does not enhance sensitivity to Stx1 (16). In these studies, however, apoptosis was not measured; instead, cytotoxicity was assessed as inhibition of protein synthesis. A more recent study specifically measured the apoptotic response of isolated renal tubular cells after exposure to Stx1 (19). Toxin treatment induced apoptosis of tubular cells, and this effect was enhanced by prestimulation with TNF-alpha . Thus in vivo disease pathophysiology could be altered during infection with STEC by both bacterial factors, including lipopolysaccharide, and by host factors such as the release of proinflammatory mediators.

A study of renal tissue obtained from three children with postenteropathic hemolytic uremic syndrome detected apoptotic cells within renal tubuli and cortices (19). Isolated human renal tubular cells underwent programmed cell death on exposure to Stx1. Shiga toxins produced by E. coli could contribute to the tissue injury observed in the hemolytic uremic syndrome. However, the mechanism(s) by which Shiga toxins mediate programmed cell death have not been elucidated. This is the first study to document the molecular determinants involved in the death signal transduced by Stx1 and Stx2 by demonstrating that induction of apoptosis by these toxins is associated with an enhanced expression of Bax, a proapoptotic homolog. Furthermore, both overexpression of Bcl-2 and caspase inhibition prevented toxin-mediated cell death.

The mechanism by which Bax stimulates apoptosis or Bcl-2 blocks cell death in sensitive cells remains unclear. Through competitive dimerization the ratio of pro- to antiapoptotic Bcl-2 family members determines the fate of a cell after an apoptotic stimulus (25). However, recent evidence suggests that the cell death signal transmitted by these proteins is more complex (1). A feature common to cells undergoing some forms of apoptosis is the disruption of the mitochondrial transmembrane potential through the opening of mitochondrial permeability transition pores (25). The breakdown of the transmembrane potential results in the release of cytochrome c into the cytoplasm. Cytoplasmic cytochrome c complexes with the molecule Apaf-1 and the proform of caspase-9. In the presence of ATP, caspase-9 is activated, thereby triggering the caspase cascade resulting in membrane blebbing and DNA fragmentation---the preeminent features characteristic of programmed cell death (48). Bax, which becomes localized to mitochondria during apoptosis, mediates the release of cytochrome c (39). In cells overexpressing Bax, Bcl-2 overexpression prevents both the release of cytochrome c and apoptosis (8, 39). In contrast, caspase inhibition prevents some features of apoptosis, such as nuclear fragmentation, but does not block Bax-mediated release of cytochrome c (39). Thus the apoptotic cross-talk mediated by Bax and Bcl-2 likely involves factors in addition to the release of cytochrome c.

On the basis of the results of this study, we propose a model to outline the pathways activated during toxin-mediated apoptosis (Fig. 6). Binding to the Gb3 receptor in human mucosal epithelial cells by Shiga toxins produced by STEC mediates an apoptotic signal transduction cascade associated with enhanced expression of Bax. The development of therapies designed to inhibit this cell death cascade could lead to improved treatment options to prevent the complications of infection caused by enteric bacterial pathogens that elaborate Shiga toxins.


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Fig. 6.   Proposed model of pathways mediating apoptosis in epithelial cells exposed to Shiga-like toxins. After binding of holotoxin to Gb3 receptor in sensitive cells, there is an increase in Bax expression. This proapoptotic Bcl-2 family member could result in activation of caspase cascade, thereby signaling cell to undergo apoptosis. Alternatively, caspase activation may occur upstream of changes in Bax. Both overexpression of Bcl-2 and inhibition of caspase activation block toxin-mediated programmed cell death.


    ACKNOWLEDGEMENTS

N. L. Jones is the recipient of a Research Initiative Award from the Canadian Association of Gastroenterology/Astra Zeneca/Medical Research Council of Canada. A. Islur is the recipient of Summer Studentship Awards from the Canadian Association of Gastroenterology and the Crohn's and Colitis Foundation of Canada.


    FOOTNOTES

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. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: P. M. Sherman, Division of Gastroenterology/Nutrition, Rm. 8411, The Hospital for Sick Children, 555 University Ave., Toronto, Ontario, Canada M5G 1X8 (E-mail: philip.sherman{at}sickkids.on.ca).

Received 22 December 1998; accepted in final form 7 December 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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