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. 2006 Jan;74(1):81–87. doi: 10.1128/IAI.74.1.81-87.2006

Caspase and Bid Involvement in Clostridium difficile Toxin A-Induced Apoptosis and Modulation of Toxin A Effects by Glutamine and Alanyl-Glutamine In Vivo and In Vitro

Benedito A Carneiro 1,†, Jun Fujii 2, Gerly A C Brito 3, Cirle Alcantara 1, Reinaldo B Oriá 1,3, Aldo A M Lima 4, Tom Obrig 2, Richard L Guerrant 1,*
PMCID: PMC1346681  PMID: 16368960

Abstract

Clostridium difficile is the leading cause of nosocomial bacterial diarrhea. Glutamine and its stable and highly soluble derivative alanyl-glutamine, have been beneficial in models of intestinal injury. In this study, we extend our work on the mechanisms of Clostridium difficile toxin A (TxA)-induced apoptosis in human intestinal epithelial T84 cells and evaluate the effects of glutamine and alanyl-glutamine on TxA-induced apoptosis in vitro and disruption of ileal mucosa in vivo. T84 cells were incubated with TxA (100 ng/ml) in medium with or without glutamine or alanyl-glutamine (3 to 100 mM). Apoptosis was evaluated by DNA fragmentation in vitro and the terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end-labeling method in vivo. Caspase and Bid involvement were investigated by Western blotting. Ligated rabbit ileal loops were used for the evaluation of intestinal secretion, mucosal disruption, and apoptosis. TxA induced caspases 6, 8, and 9 prior to caspase 3 activation in T84 cells and induced Bid cleavage by a caspase-independent mechanism. Glutamine or alanyl-glutamine significantly reduced TxA-induced apoptosis of T84 cells by 47% and inhibited activation of caspase 8. Both glutamine and alanyl-glutamine reduced TxA-induced ileal mucosal disruption and secretion. Altogether, we further delineated the apoptosis-signaling cascade induced by TxA in T84 cells and demonstrated the protective effects of glutamine and alanyl-glutamine. Glutamine and alanyl-glutamine inhibited the apoptosis of T84 cells by preventing caspase 8 activation and reduced TxA-induced intestinal secretion and disruption.

Clostridium difficile is the leading cause of nosocomial bacterial diarrhea (17, 30) and accounts for 10 to 20% of cases of antibiotic-associated diarrhea (3). C. difficile infection manifestations range from asymptomatic carriage to severe pseudomembranous colitis (3). The estimated 250,000 to 300,000 annual cases of C. difficile in the United States represent a major economic burden, each case costing up to $10,000 of extra care (49). The bacterium causes intestinal damage through the actions of toxin A (TxA) and toxin B. Although the enterotoxic TxA plays a critical role in the pathogenesis of C. difficile-associated diarrhea, strains producing only toxin B have been associated with human disease (26, 35). In addition to its proinflammatory and prosecretory activities, TxA induces apoptosis (programmed cell death) in intestinal epithelial cells, mast cells, and endothelial cells, which might contribute to the characteristic intestinal mucosal disruption (1, 6, 8, 18, 39).

Apoptosis is executed by a family of intracellular cysteine proteases (caspases) present as latent precursors that are activated through two major apoptotic pathways, extrinsic and intrinsic. The extrinsic pathway is triggered upon stimulation of death receptors by agonistic antibodies or ligands, such as tumor necrosis factor alpha (TNF-α) or FasL (48). Activation of CD95 (Fas/Apo-1), the best-characterized death receptor, leads to the formation of a death-inducing signaling complex and recruitment of procaspase 8 via the adapter molecule FADD (Fas-associated death domain protein). Procaspase 8 is then activated at the death-inducing signaling complex by proximity-induced autocatalysis (19). The intrinsic pathway is activated by several stimuli that damage the mitochondria, causing cytochrome c release into the cytosol (15). Cytochrome c in association with dATP and APAF-1 (apoptotic protease-activating factor) leads to activation of procaspase 9 (23). Both pathways converge at the activation of downstream effector caspase 3 (16, 44), which propagates the cascade by activation of other caspases, such as caspase 6 and caspase 7 (43). The effector caspases, including caspases 3 and 6, are ultimately responsible for degradation of several key structural proteins and DNA fragmentation (47). We have shown that both pathways participate in apoptosis induced by TxA in T84 cells (6), as well as the involvement of Bid (BH3 interacting death agonist), a proapoptotic Bcl-2 family member that, after being cleaved in the cytoplasm, translocates to the mitochondria and induces cytochrome c release (6, 54).

Glutamine (Gln) is a major respiratory fuel for the intestinal epithelium (50), and its supplementation is efficacious in repairing intestinal mucosal injury caused chemotherapeutic agents (55) and radiation (53), as well as in driving NaCl absorption in experimental models of infectious diarrhea, such as cholera (42), cryptosporidiosis (2), and rotavirus enteritis (36). These actions support the use of glutamine as a therapeutic adjuvant in infectious diarrhea (9) and catabolic stressful conditions (i.e., cancer chemotherapy, prolonged parenteral nutrition, sepsis, and human immunodeficiency virus-related wasting syndrome) (7, 12, 41, 55). Furthermore, glutamine not only acts as a building block for protein synthesis, it also modulates specific cellular signaling pathways, such as induction of heat shock protein (HSP) expression (52), activation of protein kinases (37), and regulation of redox status (29). In addition, glutamine deprivation induces apoptosis in intestinal epithelial cells (32), and its supplementation delays neutrophil apoptosis (34).

In the current study, we further delineate the apoptosis signaling cascade induced by TxA in T84 cells, and we show the ability of glutamine and its stable and highly soluble derivative, alanyl-glutamine (AlaGln), to inhibit TxA-induced apoptosis and reduce TxA-induced intestinal mucosal disruption and secretion in vivo.

MATERIALS AND METHODS

Reagents.

Anti-caspase primary antibodies were obtained from Cell Signaling Technology, Beverly, MA, and Santa Cruz Biotechnology, Santa Cruz, CA (anti-Bid). Glutamine and alanyl-glutamine were purchased from Sigma (St. Louis, MO) and DeGussa Corp. (Parsippany, NJ), respectively. General caspase inhibitor (Z-VAD-fmk), caspase 1 inhibitor (Z-YVAD-fmk), and caspase 8 inhibitor (Z-IETD-fmk) were purchased from Enzyme System Products (Livermore, CA).

Cell culture.

T84 cells from the American Type Culture Collection (Manassas, VA) were cultured at 5% CO2 in 50% Dulbecco's minimal essential medium, 50% Ham's F-12 medium (DMEM-F-12; Cellgro, Herndon, VA) supplemented with 5% fetal bovine serum (Gibco BRS, Gaithersburg, MD), and 100 μg/ml penicillin G and 0.085 mg/ml streptomycin (Gibco BRL, Grand Island, NY). DMEM-F-12 medium without glutamine (Cellgro, Herndon, VA) was used to evaluate the effect of glutamine or alanyl-glutamine supplementation.

Toxin A.

Purified TxA from C. difficile (strain 10463; 308 kDa) was kindly provided through our collaboration with David Lyerly, Tech Lab, Blacksburg, VA.

Western blot analysis of caspases and Bid.

Bid and caspase cleavages were evaluated as previously described (4). T84 cells were seeded in 75-cm2 tissue culture flasks, grown to ∼80% confluence, and treated with fresh medium with or without TxA (100 ng/ml) for 6, 18, 24, or 48 h. After each incubation period, the supernatants were collected, and the cells were scraped off the flasks and harvested (450 × g; 5 min; 4°C). The cell pellet was washed twice with ice-cold phosphate-buffered saline (PBS), homogenized with 0.25 ml of lysis buffer (50 mM Tris-Cl [pH 8.0]; 50 mM NaCl; 0.1 mM EDTA; 1% Tween 20; 1 mM [each] dithiothreitol, leupeptin, aprotinin, and phenylmethylsulfonyl fluoride), and incubated for 15 min at room temperature. The cell lysate was centrifuged (2,000 × g; 5 min; 4°C), and the supernatant was further clarified (15,000 × g; 15 min; 4°C) and used as a cytosolic extract in Western blotting. After the protein concentration of each sample was determined (BCA protein assay kit; Pierce Chemical Co., Rockford, IL), equal quantities of cytosolic proteins were separated by electrophoresis using 10% and 16% Tris-Glycine gels (21). The gels were transferred onto nitrocellulose membranes. The membranes were blocked with TBS-T buffer (20 mM Tris base, 500 mM NaCl, 0.1% Tween 20, pH 7.5) containing 4 to 5% (wt/vol) nonfat dry milk for 1 h at room temperature and were incubated with primary antibodies overnight at 4°C. The membranes were washed three times for 10 min each time with TBS-T buffer and incubated for 1 h at room temperature with horseradish peroxidase-conjugated secondary antibodies, anti-rabbit immunoglobulin (Ig) (1:2,500), anti-mouse Ig (1:5,000), and anti-goat Ig (1:20,000). The Western blots were developed using the ECL plus system (Amersham Pharmacia Biotech, Piscataway, NJ).

DNA fragmentation assay.

T84 cells were seeded in six-well plates (106 cells/well), and 24 h after the seeding, the medium was changed to fresh medium. Media without glutamine (controls) or media supplemented with glutamine (3 to 100 mM) or with alanyl-glutamine (3 to 100 mM) were added to different wells 1 h before TxA (100 ng/ml). The concentrations of glutamine and alanyl-glutamine used in this study were based on previous reports about the positive effects of Gln (30 to 100 mM) in sodium-dependent cation cotransport in isolated rabbit ileal mucosal preparations mounted in Ussing chambers (24) and on the in vitro protective effect of Gln on the TxA-induced drop in transepithelial resistance (5). After 24 h, the supernatants were collected, and the cells were scraped from the wells and harvested by centrifugation at 200 × g for 5 min at 4°C. The pellets were lysed with 0.3 ml hypotonic lysing buffer (100 mM Tris-HCl, 10 mM EDTA) containing 0.5% Triton X-100 for 30 min on ice. The lysates were centrifuged at 13,000 × g for 10 min to separate intact from fragmented chromatin. The supernatant, containing DNA fragments, was placed in a separate 1.5-ml microcentrifuge tube, and both pellet (containing intact chromatin) and supernatant were treated at 4°C for 30 min with 1 N perchloric acid. The precipitates were sedimented at 13,000 × g for 20 min. The DNA precipitates were hydrolyzed at 70°C for 10 min in 0.15 ml 1 N perchloric acid and quantitated using a modification of the diphenylamine method of Burton (40).

Ligated rabbit ileal loops.

As described previously (1), 2-kilogram New Zealand White rabbits were used in these experiments. After anesthesia with ketamine and xylazine (60 to 80 and 5 to 10 mg/kg intramuscularly, respectively), each rabbit was shaved, and a midline abdominal incision was made. The distal 40 to 60 cm of the ileum was exposed and flushed with PBS. Six animals were used, and 8 to 11 ileal segments 4 cm in length were doubly ligated in each animal. Negative-control loops were injected intraluminaly with PBS only (n = 6), and positive-control loops received PBS plus 10 μg TxA (n = 16), for a total volume of 1 ml/loop. Treatment loops received glutamine or alanyl-glutamine (30 and 100 mM) immediately prior to PBS or TxA (4 and 6 loops received 30 and 100 mM Gln, respectively; 8 and 11 loops received 30 and 100 mM AlaGln, respectively). The ileal loops were returned to the abdominal cavity, and the incision was sutured. After 5 h, the rabbits were euthanized, and the intestinal loops were collected. The length of each segment was measured, and the intraluminal fluid was extracted. We recorded the volume of the fluid and calculated the volume-to-length ratio in milliliters per centimeter for each loop. The intestinal sections were fixed in 10% formalin, stained with hematoxylin-eosin for histopathological evaluation, and also processed for terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end-labeling using an ApopTag Plus Peroxidase in Situ Detection Kit (Serologicals Corporation, Norcross, GA) for analysis of apoptosis.

Briefly, paraffin-embedded sections were hydrated and incubated with 20 μg/ml of proteinase K (Sigma, MO) for 15 min at room temperature. Endogenous peroxidase was blocked with 3% (vol/vol) hydrogen peroxide in PBS for 5 min at room temperature. After being washed, the sections were incubated at 37°C for 1 h with TdT buffer (125 mM Tris-HCl, 1 M sodium cacodylate, 1.25 mg/ml bovine serum albumin, pH 6.6), for 10 min with a stop/wash buffer, and then for 30 min with antidigoxigenin peroxidase conjugate. After being washed with PBS, the slides were covered with peroxidase substrate to develop color, washed three times, and counterstained in 0.5% (vol/vol) methyl green.

RESULTS

Time course of caspase activation.

T84 cells were treated with TxA, and the activation of caspases was evaluated at different intervals by Western blotting. TxA induced activation of caspases 8, 9, and 6 at 18 h, followed by caspase 3 cleavage that occurred at 24 h and peaked at 48 h (Fig. 1). These findings confirm the involvement of both intrinsic and extrinsic apoptotic pathways in TxA-induced apoptosis and their convergence at capase 3, as observed previously using a different method (6).

FIG. 1.

FIG. 1.

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Time course of caspase (Casp) 8, 9, 6, and 3 activation in T84 cells treated with C. difficile toxin A (100 ng/ml). Cleaved caspases 8, 9, and 6 were detected 18 h after incubation with toxin A. Caspase 3 activation occurred at 24 h and peaked at 48 h. Caspase cleavage was analyzed by immunoblotting in cellular lysates of T84 cells incubated with toxin A for the indicated time periods; β-actin was used as a loading control for an equal amount of protein.

TxA induces Bid activation by a caspase-independent mechanism.

We have shown that TxA activates Bid in T84 cells at 6 h, preceding caspase 8 activation (6). This suggested that Bid activation was mediated by a caspase 8-independent mechanism. In the current work, we confirmed the TxA-induced early Bid activation at 6 h prior to immunoblotting detection of caspase 8 cleavage, which occurred initially at 18 h. Moreover, the inability of inhibitors of caspases 8 and 1 or general caspase inhibitor to block Bid activation induced by TxA at 6 h suggested that Bid activation occurred by a caspase-independent mechanism (Fig. 2).

FIG. 2.

FIG. 2.

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Toxin A (100 ng/ml) induced Bid activation at 6 h by a caspase-independent mechanism. Pretreatment with inhibitors of caspase (Casp) 8 (40 μM) and caspase 1 (40 μM) and general caspase inhibitor (z-VAD; 20 μM) did not prevent the cleavage of Bid induced by toxin A. T84 cells were preincubated for 1 h with the respective caspase inhibitor, followed by incubation with toxin A. Bid cleavage was analyzed by immunoblotting in cellular lysate of T84 cells incubated with toxin A for 6 h.

These data raised the question of whether the classical extrinsic pathway of caspase 8 activation and secondary Bid cleavage was functional in T84 cells. To address this question, T84 cells were treated with recombinant TNF-α (5 ng/ml), an activator of the extrinsic pathway and caspase 8, and the time courses of both caspase 8 and Bid cleavage were evaluated. Caspase 8 activation was detected 6 h after TNF-α was added simultaneously with Bid cleavage at T84 cells, demonstrating the functionality of the extrinsic pathway in these cells (data not shown). Therefore, the early caspase 8-independent Bid activation induced by TxA likely represents a specific response to TxA, as opposed to an intrinsic signaling defect of T84 cells.

Glutamine or alanyl-glutamine reduced apoptosis induced by TxA in T84 cells.

Based upon previous data showing an antiapoptotic activity of glutamine in intestinal epithelial cells (5, 13, 32) and neutrophils (34), we investigated the effects of both glutamine and alanyl-glutamine on TxA-induced apoptosis in T84 cells. Treatment with glutamine (100 mM) or alanyl-glutamine (100 mM) 1 h before TxA (100 ng/ml) reduced the DNA fragmentation in T84 cells by 47% 24 h later (Fig. 3).

FIG. 3.

FIG. 3.

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Glutamine (100 mM)- or alanyl-glutamine (100 mM)-enriched medium added 1 h before toxin A (100 ng/ml) challenge reduced the DNA fragmentation in T84 cells by 47% 24 h later. The percentage of DNA fragmentation per total amount of DNA was determined by the Burton method. The bars represent means plus standard errors of the mean of the percentage of fragmented DNA. The control group (C), not treated with either toxin A, glutamine, or alanyl-glutamine, did not show a significant amount of apoptosis. The asterisks indicate statistically significant differences (P < 0.05) in the percentage of DNA fragmentation between the group treated with toxin A plus glutamine or alanyl-glutamine and the group treated with toxin A plus PBS. Statistical significance was determined by analysis of variance and Bonferroni's test. The numbers above each column represent n for each group.

Effect of glutamine or alanyl-glutamine on caspase 8, 6, and 9 activation induced by TxA in T84 cells.

The effects of glutamine and alanyl-glutamine on caspase activation in T84 cells were evaluated to investigate their antiapoptotic mechanisms. Both glutamine and alanyl-glutamine significantly prevented caspase 8 activation induced by TxA at 18 and 24 h (Fig. 4A) without affecting caspase 6 (Fig. 4A) or caspase 9 (Fig. 4B) activation.

FIG. 4.

FIG. 4.

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Glutamine and alanyl-glutamine prevented caspase 8 activation induced by toxin A at 18 and 24 h but had no effect on caspase 6 or caspase 9 activation. (A) Immunoblotting detection of cleaved caspases (Casp) 8 and 6. (B) Cleaved caspase 9 detection. Caspases were detected by immunoblotting in cellular lysate of T84 cells incubated with toxin A (100 ng/ml) or PBS (negative controls) for the indicated time periods. Samples were preincubated with either Gln (100 mM) or AlaGln (100 mM) 1 h prior to toxin A challenge.

Glutamine or alanyl-glutamine reduced mucosal disruption, apoptosis, and intestinal secretion induced by TxA in vivo.

TxA (10 μg/ml) injected into the lumen of ligated ileal loops induced significant intestinal secretion 5 h later, as described previously (1). TxA also caused an intense mucosal disruption (Fig. 5C) and induced a large amount of apoptosis in mucosal cells of rabbit ileal loops (Fig. 5D) in comparison to ileal loops treated with PBS (Fig. 5A and B). Simultaneous administration of glutamine (Fig. 5E and F) or alanyl-glutamine (Fig. 5G and H) (100 mM) with TxA attenuated mucosal disruption and apoptosis, as well as significantly reduced intestinal secretion, in the experimental model of Clostridium difficile TxA enteritis (Fig. 6).

FIG. 5.

FIG. 5.

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Prevention of toxin A-induced mucosal disruption and apoptosis by glutamine (100 mM) or alanyl-glutamine (100 mM) compared to the PBS-treated controls. The left column depicts ileal sections stained with hematoxylin-eosin, and the right column shows the same ileal sections stained by the terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end-labeling method; the brown dots represent apoptotic cells (scale, 1 cm = 70 μm). Severe mucosal disruption (C) and significant amounts of apoptosis in the mucosal cells (D) are seen in the ligated rabbit ileal loops treated with toxin A (10 μg/ml) for 5 h in comparison with controls treated with PBS (A and B). Glutamine (E and F) or alanyl-glutamine (G and H) significantly reduced mucosal disruption and apoptosis induced by toxin A.

FIG. 6.

FIG. 6.

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Effects of glutamine and alanyl-glutamine on Clostridium difficile toxin A-induced secretion in ligated rabbit ileal loops. Rabbit ileal loops were treated simultaneously with toxin A (10 μg/loop) and solutions containing Gln or AlaGln (30 or 100 mM) for 5 h, and then the loop length and the volume of secretion were measured. The bars represent means plus standard errors of the mean of the volume of secretion per centimeter of loop length. The asterisks indicate statistically significant differences (P < 0.05) in the volume of secretion/intestinal-loop length between the groups treated with toxin A plus glutamine or alanyl-glutamine and the group treated with toxin A plus PBS. Statistical significance was determined by analysis of variance and Bonferroni's test. The numbers above each column represent n for each group.

DISCUSSION

The results of this work extend and complement previous studies on the mechanisms of apoptosis induced by TxA in human intestinal epithelial cells and demonstrate the inhibitory effect of glutamine and alanyl-glutamine. In the present study, we demonstrated cleavage of caspases 6, 8, and 9 18 h after incubation with TxA, followed by activation of caspase 3 at 24 h with a peak at 48 h. This time course differs from a previous report, which showed later activation of caspases 6, 8, and 9 using detection of enzymatic activity with a fluorogenic substrate (6). Thus, the data obtained by Western blotting confirms the involvement of both intrinsic and extrinsic pathways in TxA-induced apoptosis and shows more detailed information about the time course of caspase activation. We also showed the caspase-independent activation of Bid.

The proapoptotic Bcl-2 family member Bid amplifies the caspase cascade by connecting the death receptor pathway with the mitochondrial apoptotic pathway (28). After being cleaved by caspase 8 from its inactive 25-kDa form localized in the cytoplasm, the truncated form of Bid translocates to the mitochondria and induces cytochrome c release (22). Besides caspase 8, granzyme B, caspase 1, and lysosomal proteases also cleave Bid, enabling the cross-linking between the two major pathways of caspase activation (22, 33, 45). Two results support the notion that Bid activation in T84 cells exposed to TxA occurs by a caspase-independent mechanism: (i) Bid cleavage occurred before caspase 8 activation; (ii) inhibitors of caspase 8 or caspase 1 or general caspase inhibitors did not block Bid activation. Activation of alternative proteolytic events by TxA, such as the release of lysosomal proteases, might also explain the activation of Bid, but it remains to be demonstrated. Lysosomal extracts have been shown to cleave Bid and cause cytochrome c release, and lysosomal cysteine proteases—cathepsins—have been implicated in caspase activation (38, 45).

Previous studies showing the antiapoptotic properties of glutamine prompted us to study the effects of glutamine and its highly soluble and stable derivative alanyl-glutamine on the T84 cell apoptosis induced by TxA. Glutamine supplementation delayed human neutrophil apoptosis (34) and reduced T-cell apoptosis by increasing glutathione and Bcl-2 levels (11). Glutamine deprivation also induced apoptosis in rat intestinal epithelial cells (32) and rendered premonocytic and HL-60 cells significantly more susceptible to Fas-mediated apoptosis (14, 20). In agreement with these previous studies documenting the antiapoptotic activity of glutamine, both glutamine and alanyl-glutamine reduced the T84 cell apoptosis induced by TxA by almost 50% through blockage of caspase 8 activation.

Glutamine and alanyl-glutamine specifically inhibited caspase 8 activation induced by TxA, without interfering with caspase 6 or 9 activation. This specific effect on caspase 8 is consistent with preliminary data from our laboratory showing that both glutamine and alanyl-glutamine significantly increased the levels of cFLIP (c-Fas-associated death domain-like interleukin-1-converting enzyme-like inhibitory protein) in T84 cells (data not shown). cFLIP binds to the adaptor molecule FADD, preventing the recruitment and activation of caspase 8 (31, 46). Another study also demonstrated the antiapoptotic effect of glutamine through inhibition of the extrinsic pathway. Glutamine prevented HT-29 cell (human intestinal epithelial cells) apoptosis induced by TNF-α-related apoptosis-inducing ligand, which is characterized by caspase 8 activation (13). The predominant modulation of the extrinsic pathway may explain the approximately 50% of residual apoptosis measured by DNA fragmentation when the cells were pretreated with glutamine or alanyl-glutamine.

In addition, in vivo experiments were performed to corroborate the inhibition of TxA-induced apoptosis by both glutamine and alanyl-glutamine. Significant effects of glutamine on preventing the intestinal damage induced by radiation (53) or chemotherapy (10, 55), as well as the effects of glutamine in driving sodium absorption in cholera (25) and cryptosporidium-induced diarrhea (2), provided a strong rationale. Both glutamine and alanyl-glutamine significantly reduced the well-documented intestinal damage caused by TxA in rabbit ileal loops and reduced the amount of epithelial-cell apoptosis. The secretion induced by TxA was also inhibited by glutamine and alanyl-glutamine, in accordance with previous studies from our group using cholera toxin (25). Even though we cannot extrapolate the antiapoptosis mechanisms of glutamine and alanyl-glutamine seen in T84 cells, it is reasonable to suggest that in vivo inhibition of caspase 8 might play a role in the reduction of apoptosis and intestinal damage detected on the rabbit ileal loops.

An additional mechanism to explain the protection of TxA enteritis by glutamine and alanyl-glutamine includes the induction of HSPs, which are critical to cellular survival under stressful conditions, such as heat shock and sepsis. Glutamine induced expression of HSP 72 in the colon of rats with lipopolysaccharide-induced sepsis, resulting in decreased intestinal damage and better survival (51). Accordingly, human intestinal epithelial Caco2 cells overexpressing HSP 72 were protected against TxA-induced toxic effects, such as caspase 9 activation (27).

In summary, the present study shows that TxA induces caspases 6, 8, and 9 prior to caspase 3 cleavage in T84 cells and induces Bid activation by a caspase-independent mechanism. It also demonstrates the roles of glutamine and alanyl-glutamine in inhibiting T84 cell apoptosis by blocking caspase 8 activation and reducing TxA-induced intestinal secretion and mucosal disruption. These results reinforce the potential of glutamine and alanyl-glutamine as adjuvant therapeutic measures in C. difficile enteritis and reveal novel signaling properties of these substances capable of modulating apoptosis cascades in which caspase 8 has a pivotal role.

Acknowledgments

This work was supported in part by National Institutes of Health SBIR Grant 1R43-DK58419-01A1, ICDR Grant 5-UOI AI 26512-14, and Fogarty International Center Action for Building Capacity (ABC) Grant 5D43 TW01136-04. Benedito A. Carneiro, Reinaldo B. Oria, Gerly A. C. Brito, and Cirle Alcantara were recipients of Fogarty International Center ABC research fellowships.

R. L. Guerrant is named as a coinventor on a patent held by the University of Virginia for the use of glutamine derivatives in oral rehydration therapy, but without receiving any patent royalties.

Editor: D. L. Burns

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