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Glucosylation of Rho GTPases by Clostridium difficile toxin A triggers apoptosis in intestinal epithelial cells

Institute of Toxicology, Hannover Medical School, 30625 Hannover, Germany

Correspondence
Ralf Gerhard
gerhard.ralf{at}mh-hannover.de

Received 16 November 2007
Accepted 2 March 2008

Abbreviations: FACS, fluorescence-activated cell sorting.

	INTRODUCTION

TOP INTRODUCTION METHODS RESULTS AND DISCUSSION REFERENCES

 
Clostridium difficile is a pathogen that causes antibiotic-associated diarrhoea and severe pseudomembranous colitis. Toxin A (TcdA) and toxin B (TcdB) are the main pathogenic factors released by C. difficile (Just & Gerhard, 2004). The toxins are homologous glucosyltransferases that mono-glucosylate Rho GTPases (Rho, Rac and Cdc42) (Just et al., 1995). This glucosylation leads to functional inactivation. As Rho GTPases regulate the dynamics of the actin cytoskeleton (Hall, 1998), their inactivation leads to a reorganization of filamentous actin structures. Thus, the predominant effect of TcdA and TcdB is the breakdown of the actin cytoskeleton, accompanied by a change in cell morphology. Rho GTPase inhibition results in inhibition of migration, morphogenesis, cell division (Jaffe & Hall, 2005) and membrane trafficking (Ridley, 2001). Additionally, Rho GTPases are involved in transcriptional regulation and apoptosis (Aznar & Lacal, 2001; Symons, 1996). They play a critical role in apoptosis associated with morphological changes in cells (Coleman & Olson, 2002; Coniglio et al., 2001), or contribute in a pro- or anti-apoptotic manner as a response to genotoxic stress (Fritz & Kaina, 2006).

The apoptotic properties of TcdA and TcdB are well known. These toxins induce apoptosis in intestinal epithelial cells (Brito et al., 2002, 2005; Carneiro et al., 2006; Fiorentini et al., 1998; Liu et al., 2003; Mahida et al., 1998) and in all other cells tested such as neurons (Le et al., 2005), endothelial cells (Hippenstiel et al., 2002), HeLa cells (Qa'Dan et al., 2002) and monocytes (Warny & Kelly, 1999). It is common sense that the executioner caspase-3 is involved in TcdA-induced apoptosis (Brito et al., 2002; Carneiro et al., 2006; Hippenstiel et al., 2002; Le et al., 2005; Qa'Dan et al., 2002; Solomon et al., 2005). Additionally, activation of caspase-6, another executioner caspase, can also be observed (Brito et al., 2002; Carneiro et al., 2006). The initiator caspase-8 represents the extrinsic apoptotic pathway, which can be activated via transmembrane death receptors such as the Fas receptor. Caspase-9, representing the intrinsic pathway, is activated by an apoptosome containing cytochrome c released from damaged mitochondria (Boatright & Salvesen, 2003). Both initiator caspases (8 and 9) are known to cleave procaspase-3 into the activated caspase-3. The activation of the intrinsic pathway by clostridial toxins has been described by different authors. Cytochrome c is released from mitochondria in response to treatment with TcdA or TcdB (Brito et al., 2002; Gerhard et al., 2005; Kim et al., 2005; Le et al., 2005; Liu et al., 2003). As a consequence, caspase-9 is consecutively activated (Brito et al., 2002; Carneiro et al., 2006; Hippenstiel et al., 2002; Le et al., 2005; Liu et al., 2003). However, caspase-8 also plays a role in TcdA-induced apoptosis (Brito et al., 2002; Carneiro et al., 2006). Intracellular effects of TcdA that trigger pro-apoptotic events are not known. The glucosylation of Rho GTPases as a prerequisite for apoptosis is controversial and is discussed below. Mutant forms of TcdA (Teichert et al., 2006) were used here to elucidate the impact of Rho glucosylation on the apoptotic processes.

	METHODS

TOP INTRODUCTION METHODS RESULTS AND DISCUSSION REFERENCES

 
Chemicals and reagents. The antibodies used were: polyclonal anti-caspase-3 antibody recognizing procaspase-3 and cleaved caspase-3, polyclonal antibody recognising cleaved caspase-8 (Asp374) and polyclonal antibody recognising cleaved caspase-9 (Asp315) (all from Acris Antibodies), monoclonal anti-Rac1 antibody (clone 102; BD PharMingen), monoclonal anti-Rac1 antibody (clone 23A8; Upstate), horseradish-conjugated anti-mouse IgG and anti-rabbit IgG (Rockland Immunochemicals). The pan-caspase inhibitor z-VAD-fmk was from Merck Biosciences. The Bacillus megaterium expression system was from MoBiTec.

Cell culture. The colonic crypt cell line HT-29 was cultured under standard conditions in Dulbeccos' modified Eagle's medium/F12 mix supplemented with 10 % fetal bovine serum, 100 µM penicillin and 100 µg streptomycin ml^–1. For stimulation experiments, cells were seeded in Petri dishes (3.5 cm diameter) coated with fibronectin.

Preparation of toxins. All experiments were performed with expressed recombinant TcdA. The recombinant and genetically engineered mutants were expressed in a B. megaterium expression system as His-tagged fusion proteins. The constructs were sequenced to ensure that the sequence was identical compared with the reference gene of C. difficile TcdA, strain VPI 10463 (GenBank accession no. X51797), except for the desired mutations. Purification was performed by Ni²⁺ affinity chromatography as described previously (Burger et al., 2003). Toxins were stored until use in 10 mM Tris/HCl (pH 7.4), 50 mM NaCl at –80 °C. The recombinant toxins exhibited all features of the native TcdA, i.e. glucosyltransferase activity (except when specifically reduced by directed mutagenesis), cell binding, endocytotic uptake and autocatalytic cleavage of the catalytic domain (data not shown). Different conditions for expression and purification of native TcdA from C. difficile and of the recombinant TcdA did not induce alteration of any of these features (Gerhard et al., 2005). The absence of cytotoxic impurities acting on isolated mitochondria was checked using a cytochrome c release assay.

Flow cytometry. The number of apoptotic or necrotic cells was counted by fluorescence-activated cell sorting (FACS). Detached and adherent cells, which were suspended by trypsinization, were pooled before FACS analysis. The DNA content was measured using the fluorescent nucleotide acid dye propidium iodide. Cells (0.5x10⁶) were fixed in 70 % ice-cold ethanol for 30 min and washed once with 1 % BSA in PBS. The DNA was stained with propidium iodide [150 µg ml^–1 in Tris/HCl (pH 7.4) containing 1 % BSA and 1 % Triton X-100]. RNA was removed by incubating the cells with 0.5 % RNase for 30 min. The cells were then subjected to flow cytometry (FACScan flow cytometer; Becton Dickinson). A fluorescence area (FL2) of 400 was set to correlate with a 2n set of chromosomes within the G₁ phase. Cells found in the sub-G₁ phase were considered to be apoptotic and necrotic due to the decrease in DNA content.

Western blotting. Activation of caspase-9, -8, and -3 was shown by Western blot analysis, using antibodies that recognize the cleaved and thus activated caspases. Whole-cell homogenates were resolved by SDS-PAGE and transferred to nitrocellulose. Immunodetection was performed in Tris-buffered saline containing 0.2 % (v/v) Tween 20. The nitrocellulose was incubated overnight at 4 °C with primary antibodies, followed by the appropriate secondary antibody for 45 min at room temperature. Femto SuperSignal ECL Substrate (Pierce) was used for detection.

Statistical analysis. A two-sided Student's t-test was performed to determine significant effects. A value of P<0.01 was considered significant and is indicated with asterisks in the figures. The t-test was only used for studies with n5 experiments. All values are given as means±SD.

	RESULTS AND DISCUSSION

TOP INTRODUCTION METHODS RESULTS AND DISCUSSION REFERENCES

 
All experiments were performed using expressed recombinant TcdA. The recombinant toxins exhibited all features of the native TcdA, i.e. glucosyltransferase activity (except when specifically reduced by directed mutagenesis), cell binding, endocytotic uptake and autocatalytic cleavage of the catalytic domain (data not shown). Different conditions for expression and purification of native TcdA from C. difficile and of the recombinant TcdA did not induce alteration of any of these features (Gerhard et al., 2005).

Glucosyltransferase activity of TcdA is crucial for induction of apoptosis

TcdA (3 nM) induced morphological changes in HT-29 cells and the cells rounded up within 8 h (Fig. 1a). The kinetics of Rac glucosylation, which was assayed using a specific antibody (Genth et al., 2006), was in strong correlation with cell rounding (data not shown). The activation of caspase-3 followed the glucosylation of Rho GTPases. The results of Western blot analyses with specific antibody recognizing uncleaved procaspase-3 (35 kDa) as well as the cleaved and thus activated caspase-3 (12 kDa) are shown in Fig. 1(b). There was a strong correlation between caspase activation (Fig. 1b, upper panel) and Rac1 glucosylation (Fig. 1b, middle panel). Total Rac1 is shown in the lower panel. The maximum activation of caspase-3 was detectable only when Rac1 was quantitatively glucosylated. The 50 % effective concentration required to activate caspase-3 after 36 h was approximately 50 ng TcdA ml^–1 (0.2 nM). Recombinant TcdA with mutations that specifically affect the glucosyltransferase activity (Fig. 2a) is an ideal tool to distinguish between glucosyltransferase-dependent and -independent effects. Different mutants were used to check for glucosyltransferase-independent effects. The purity of toxin preparation (Fig. 2b, left panel) and the absence of cleaved N-terminal glucosyltransferase (Fig. 2b, right panel) were checked by SDS-PAGE and Western blot analysis with specific anti-TcdA1–542 antibody, respectively. TcdA W101A had 50-fold reduced glucosyltransferase activity compared with wild-type TcdA, and was applied in an equimolar or equipotent ratio to distinguish between Rho glucosylating effects and those that are independent. The double mutant TcdA D285/287N does not glucosylate Rho GTPases when applied to cells, even at the highest achievable concentration of 0.3 µM (Teichert et al., 2006). The cytotoxic effect of TcdA resulting from caspase activation was measured by flow cytometry. As shown in Fig. 2(c), 1 nM TcdA induced apoptosis in 28±6 % of cells after 36 h incubation compared with untreated cells (5±2 %). The double mutant TcdA D285/287N, which had no Rho glucosylating properties in a cell culture assay, did not induce apoptosis (4±3 %). Mutant TcdA W101A led to a slight and non-significant increase (8±4 %) in the number of apoptotic cells at a concentration of 1 nM. However, at a 50-fold concentration, TcdA W101A had similar apoptotic features (21±15 %) to wild-type TcdA. Thus, the apoptotic effect of TcdA correlates with the glucosyltransferase activity that is displayed intracellularly. Destruction of the actin cytoskeleton alone by latrunculin B (5 µg ml^–1) did not induce apoptosis in HT-29 cells. Fig. 2(d) shows the activated, i.e. cleaved, caspase-3 after treatment of cells with either wild-type TcdA, TcdA W101A or TcdA D285/287N (1 nM each). This set of experiments showed that apoptosis was induced only when Rho GTPases were maximally glucosylated (compare with Fig. 1b). Breakdown of the actin cytoskeleton was a sequel of Rho glucosylation, but did not cause apoptosis.

View larger version (86K): [in this window] [in a new window]   Fig. 1. (a) Morphological changes in HT-29 cells treated with TcdA for the indicated times. (b) Western blot analysis showing activation of caspase-3 after 36 h of treatment with TcdA at the indicated concentrations. The 50 % effective concentration of caspase-3 was determined by treatment with 50 ng TcdA ml–1 (upper panel). Glucosylation of Rac1 was detected by Western blot analysis with a specific antibody that specifically recognizes non-glucosylated Rac1 (middle panel) and total Rac1 (lower panel). The blots shown are representative of at least three separate experiments.

View larger version (26K): [in this window] [in a new window]   Fig. 2. (a) Structure of TcdA. The N-terminal glucosyltransferase domain (GTD) consisting of aa 1–542 is cleaved intracellularly by an autoproteolytic cleavage. The transmembrane domain (TMD) and the receptor-binding domain (RBD) facilitate binding to the target cell and translocation of the GTD into the cytosol. Two conserved motifs within the GTD participate in binding of the co-substrate UDP–glucose. Mutation of the tryptophan within the WIGG motif (W101A) results in a 50-fold reduction in glucosyltransferase activity. TcdA D285/287N is a double mutant where the DXD motif was altered to extinguish the glucosyltransferase activity. (b) Left panel: Coomassie gel showing the purity of the toxin preparation. Right panel: Western blot analysis using an antibody specific for the N-terminal fragment of TcdA (aa 1–542) showing that preparations did not contain cleaved glucosyltransferase domain. (c) Apoptosis of cells treated with TcdA and mutant TcdA, estimated by flow cytometry. Wild-type TcdA (1 nM), TcdA W101A at an equipotent (50 nM) or equimolar concentration (1 nM), TcdA D285/287N (50 nM) or latrunculin B (5 µg ml–1) were applied to investigate glucosyltransferase-independent apoptotic effects. The results shown are means±SD (n=5). *, P<0.01. (d) Western blot analysis showing cleaved caspase-3 of HT-29 cells after 36 h treatment with wild-type TcdA, TcdA W101A or TcdA D285/287N (1 nM each).

View larger version (23K): [in this window] [in a new window]   Fig. 3. Time-dependent effect of TcdA on G2/M arrest and apoptosis. The number of apoptotic cells (black bars) and cells that arrested within the G2/M transition (grey bars) are shown, as determined by flow cytometry. Cells were treated with TcdA (1 nM) for the indicated times. Latrunculin B (LB) (5 µg ml–1) was applied for 48 h as a control for cytoskeleton-induced G2/M arrest. The results are shown are means±SD (n=5).

View larger version (19K): [in this window] [in a new window]   Fig. 4. (a) Flow cytometry data from cells treated with TcdA (1 nM) in the presence or absence of 50 µM z-VAD-fmk (pan-caspase inhibitor). Cells with a DNA content less than cells within the G1 phase (sub-G1) were considered apoptotic. (b) Western blot of caspase-3, -8 and -9. The pan-caspase inhibitor abolished cleavage of caspase-8 and -9, but did not prevent cleavage of procaspase-3 into intermediate fragments (indicated by arrows).

TcdA induced activation of caspase-3, -8 and -9 in HT-29 cells and thereby triggered apoptotic cell death. Activation of the executioner caspase-3 strongly correlated with the glucosylation of Rac1. Significant cleavage of procaspase-3 was detected when TcdA was applied at concentrations that were sufficient to achieve quantitative glucosylation of the Rac1 pool. It is not known which GTPase is the crucial one for induction of activation of caspases when glucosylated. The kinetics of TcdA-catalysed Rac and Rho glucosylation are almost identical (Teichert et al., 2006), and thus Western blot analysis of non-glucosylated Rac1 is an ideal surrogate for the intracellular action of TcdA (Genth et al., 2006). The correlation of the extent of glucosylation of Rho GTPases and the increasing activation of caspase-3 strongly suggest that inhibition of Rho GTPases is a cause of apoptotic cell death. A single report has described the Rho-independent effect of TcdA (Kim et al., 2005), and it is likely that TcdA exhibits functions additional to its glucosyltransferase activity when applied to cells. To study the effect of glucosylation events in TcdA-induced cell death, we used a set of TcdA mutants that differed solely in their glucosyltransferase activity. These experiments clearly showed that apoptotic cell death strictly depended on the glucosyltransferase activity of these toxins in our model system. No effect on caspases or cell viability was observed when TcdA W101A (with reduced glucosyltransferase activity) or TcdA D285/287N (no glucosyltransferase activity) was applied at concentrations higher than that of wild-type TcdA sufficient to induce maximum effects. It is not known whether inhibition of Rho, Rac, Cdc42 or a combination of these GTPases is responsible for the TcdA-induced apoptosis. Rho GTPases are positively involved in transduction of receptor-mediated apoptotic signals (Brenner et al., 1997), and can also be negatively involved (Esteve et al., 1998; Rattan et al., 2006). Inhibition of Rho by exoenzyme C3-catalysed ADP ribosylation (Bobak et al., 1997; Ikeda et al., 2003; Li et al., 2002), as well as dominant-negative Rac1 and Cdc42 mutants, induces apoptosis (Lassus et al., 2000). Specific glucosylation of Rac/Cdc42 by lethal toxin from Clostridium sordellii also induces apoptosis (Linseman et al., 2001; Petit et al., 2003). The predominance of either these Rho GTPases may be the reason for the different sensitivities, and indicates the difficulties in estimating the relative contribution of these Rho GTPases to the apoptotic properties of Rho-glucosylating toxins.

Caspase-3 is the central player in TcdA-induced cell death. Inhibition of the executioner caspase-3 prevented an apoptotic outcome for cells treated with TcdA for at least 48 h. The most important finding was that the first cleavage of procaspase-3 into intermediate fragments was carried out by a protease that was not susceptible to a pan-caspase inhibitor. This indicates that a non-caspase protease triggers activation of the cascade of caspases, where caspase-3 is simultaneously the initiator and executioner caspase. Caspase-8 and -9 are also involved, but seem to participate only in an amplifying manner.

The results of this study are summarized in a schematic diagram (Fig. 5). Treatment of cells with TcdA causes inhibition of Rho GTPases. This causes a reorganization of the actin cytoskeleton, which in turn results in arrest at the G₂/M transition. This effect is referred to as the cytopathic effect. Glucosylation of Rho GTPases also causes activation of caspases, which is independent of the effect on the actin cytoskeleton. Active caspase-3 induces apoptosis, i.e. the cytotoxic effect of TcdA. By application of different mutants of TcdA, we have shown that the glucosyltransferase activity of TcdA is responsible for both the cytopathic and the cytotoxic effect.

View larger version (24K): [in this window] [in a new window]   Fig. 5. Schematic diagram of the cytopathic and cytotoxic effects of TcdA. Glucosylation of Rho GTPases induces reorganization of the actin cytoskeleton with subsequent G2/M arrest (cytopathic effect). Independently of the cytoskeleton, Rho glucosylation results in activation of caspase-3 to execute apoptosis (cytotoxic effect). Caspase-8 and -9 participate in an amplifying manner.

	REFERENCES

TOP INTRODUCTION METHODS RESULTS AND DISCUSSION REFERENCES

 
Aznar, S. & Lacal, J. C. (2001). Rho signals to cell growth and apoptosis. Cancer Lett 165, 1–10.[CrossRef][Medline]

Boatright, K. M. & Salvesen, G. S. (2003). Mechanisms of caspase activation. Curr Opin Cell Biol 15, 725–731.[CrossRef][Medline]

Bobak, D., Moorman, J., Guanzon, A., Gilmer, L. & Hahn, C. (1997). Inactivation of the small GTPase Rho disrupts cellular attachment and induces adhesion-dependent and adhesion-independent apoptosis. Oncogene 15, 2179–2189.[CrossRef][Medline]

Brenner, B., Koppenhoefer, U., Weinstock, C., Linderkamp, O., Lang, F. & Gulbins, E. (1997). Fas- or ceramide-induced apoptosis is mediated by a Rac1-regulated activation of Jun N-terminal kinase/p38 kinases and GADD153. J Biol Chem 272, 22173–22181.[Abstract/Free Full Text]

Brito, G. A. C., Fujji, J., Carneiro, B. A., Lima, A. A. M., Obrig, T. & Guerrant, R. L. (2002). Mechanism of Clostridum difficile toxin A-induced apoptosis in T84 cells. J Infect Dis 186, 1438–1447.[CrossRef][Medline]

Brito, G. A., Carneiro-Filho, B., Oria, R. B., Destura, R. V., Lima, A. A. & Guerrant, R. L. (2005). Clostridium difficile toxin A induces intestinal epithelial cell apoptosis and damage: role of Gln and Ala-Gln in toxin A effects. Dig Dis Sci 50, 1271–1278.[CrossRef][Medline]

Burger, S., Tatge, H., Hofmann, F., Just, I. & Gerhard, R. (2003). Expression of recombinant Clostridium difficile toxin A using the Bacillus megaterium system. Biochem Biophys Res Commun 307, 584–588.[CrossRef][Medline]

Carneiro, B. A., Fujii, J., Brito, G. A., Alcantara, C., Oria, R. B., Lima, A. A., Obrig, T. & Guerrant, R. L. (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. Infect Immun 74, 81–87.[Abstract/Free Full Text]

Coleman, M. L. & Olson, M. F. (2002). Rho GTPase signalling pathways in the morphological changes associated with apoptosis. Cell Death Differ 9, 493–504.[CrossRef][Medline]

Coniglio, S. J., Jou, T. S. & Symons, M. (2001). Rac1 protects epithelial cells against anoikis. J Biol Chem 276, 28113–28120.[Abstract/Free Full Text]

Esteve, P., Embade, N., Perona, R., Jimenez, B., del Peso, L., Leon, J., Arends, M., Miki, T. & Lacal, J. C. (1998). Rho-regulated signals induce apoptosis in vitro and in vivo by a p53-independent, but Bcl2 dependent pathway. Oncogene 17, 1855–1869.[CrossRef][Medline]

Fiorentini, C., Fabbri, A., Falzano, L., Fattorossi, A., Matarrese, P., Rivabene, R. & Donelli, G. (1998). Clostridium difficile toxin B induces apoptosis in intestinal cultured cells. Infect Immun 66, 2660–2665.[Abstract/Free Full Text]

Fritz, G. & Kaina, B. (2006). Rho GTPases: promising cellular targets for novel anticancer drugs. Curr Cancer Drug Targets 6, 1–14.[CrossRef][Medline]

Genth, H., Huelsenbeck, J., Hartmann, B., Hofmann, F., Just, I. & Gerhard, R. (2006). Cellular stability of Rho-GTPases glucosylated by Clostridium difficile toxin B. FEBS Lett 580, 3565–3569.[CrossRef][Medline]

Gerhard, R., Burger, S., Tatge, H., Genth, H., Just, I. & Hofmann, F. (2005). Comparison of wild type with recombinant Clostridium difficile toxin A. Microb Pathog 38, 77–83.[CrossRef][Medline]

Hall, A. (1998). Rho GTPases and the actin cytoskeleton. Science 279, 509–514.[Abstract/Free Full Text]

Hippenstiel, S., Schmeck, B., N'Guessan, P. D., Seybold, J., Krüll, M., Preissner, K., Von Eichel-Streiber, C. & Suttorp, N. (2002). Rho protein inactivation induced apoptosis of cultured human endothelial cells. Am J Physiol Lung Cell Mol Physiol 283, L830–L838.[Abstract/Free Full Text]

Ikeda, H., Nagashima, K., Yanase, M., Tomiya, T., Arai, M., Inoue, Y., Tejima, K., Nishikawa, T., Omata, M. & other authors (2003). Involvement of Rho/Rho kinase pathway in regulation of apoptosis in rat hepatic stellate cells. Am J Physiol Gastrointest Liver Physiol 285, G880–G886.[Abstract/Free Full Text]

Jaffe, A. B. & Hall, A. (2005). Rho GTPases: biochemistry and biology. Annu Rev Cell Dev Biol 21, 247–269.[CrossRef][Medline]

Just, I. & Gerhard, R. (2004). Large clostridial cytotoxins. Rev Physiol Biochem Pharmacol 152, 23–47.[CrossRef][Medline]

Just, I., Selzer, J., Wilm, M., Von Eichel-Streiber, C., Mann, M. & Aktories, K. (1995). Glucosylation of Rho proteins by Clostridium difficile toxin B. Nature 375, 500–503.[CrossRef][Medline]

Kim, H., Kokkotou, E., Na, X., Rhee, S. H., Moyer, M. P., Pothoulakis, C. & LaMont, J. T. (2005). Clostridium difficile toxin A-induced colonocyte apoptosis involves p53-dependent p21(WAF1/CIP1) induction via p38 mitogen-activated protein kinase. Gastroenterology 129, 1875–1888.[CrossRef]

Lassus, P., Roux, P., Zugasti, O., Philips, A., Fort, P. & Hibner, U. (2000). Extinction of Rac1 and Cdc42Hs signalling defines a novel p53-dependent apoptotic pathway. Oncogene 19, 2377–2385.[CrossRef][Medline]

Le, S. S., Loucks, F. A., Udo, H., Richardson-Burns, S., Phelps, R. A., Bouchard, R. J., Barth, H., Aktories, K., Tyler, K. L. & other authors (2005). Inhibition of Rac GTPase triggers a c-Jun and Bim-dependent mitochondrial apoptotic cascade in cerebellar granule neurons. J Neurochem 94, 1025–1039.[CrossRef][Medline]

Li, X., Liu, L., Tupper, J. C., Bannerman, D. D., Winn, R. K., Sebti, S. M., Hamilton, A. D. & Harlan, J. M. (2002). Inhibition of protein geranylgeranylation and RhoA/RhoA kinase pathway induces apoptosis in human endothelial cells. J Biol Chem 277, 15309–15316.[Abstract/Free Full Text]

Linseman, D. A., Laessig, T., Meintzer, M. K., McClure, M., Barth, H., Aktories, K. & Heidenreich, K. A. (2001). An essential role for Rac/Cdc42 GTPases in cerebellar granule neuron survival. J Biol Chem 276, 39123–39131.[Abstract/Free Full Text]

Liu, T. S., Musch, M. W., Sugi, K., Walsh-Reitz, M. M., Ropeleski, M. J., Hendrickson, B. A., Pothoulakis, C., LaMont, J. T. & Chang, E. B. (2003). Protective role of HSP72 against Clostridium difficile toxin A-induced intestinal epithelial cell dysfunction. Am J Physiol Cell Physiol 284, C1073–C1082.[Abstract/Free Full Text]

Mahida, Y. R., Galvin, A., Makh, S., Hyde, S., Sanfilippo, L., Borriello, S. P. & Sewell, J. L. (1998). Effect of Clostridium difficile toxin A on human colonic lamina propria cells: early loss of macrophages followed by T-cell apoptosis. Infect Immun 66, 5462–5469.[Abstract/Free Full Text]

Petit, P., Bréard, J., Montalescot, V., El Hadj, N. B., Levade, T., Popoff, M. & Geny, B. (2003). Lethal toxin from Clostridium sordellii induces apoptotic cell death by disruption of mitochondrial homeostasis in HL-60 cells. Cell Microbiol 5, 761–771.[CrossRef][Medline]

Qa'Dan, M., Ramsey, M., Daniel, J., Spyres, L. M., Safiejko-Mroczka, B., Ortiz-Leduc, W. & Ballard, J. D. (2002). Clostridium difficile toxin B activates dual caspase-dependent and caspase-independent apoptosis in intoxicated cells. Cell Microbiol 4, 425–434.[CrossRef][Medline]

Rattan, R., Giri, S., Singh, A. K. & Singh, I. (2006). Rho/ROCK pathway as a target of tumor therapy. J Neurosci Res 83, 243–255.[CrossRef][Medline]

Ridley, A. J. (2001). Rho proteins: linking signaling with membrane trafficking. Traffic 2, 303–310.[CrossRef][Medline]

Solomon, K., Webb, J., Ali, N., Robins, R. A. & Mahida, Y. R. (2005). Monocytes are highly sensitive to Clostridium difficile toxin A-induced apoptotic and nonapoptotic cell death. Infect Immun 73, 1625–1634.[Abstract/Free Full Text]

Symons, M. (1996). Rho family GTPases: the cytoskeleton and beyond. Trends Biochem Sci 21, 178–181.[CrossRef][Medline]

Teichert, M., Tatge, H., Schoentaube, J., Just, I. & Gerhard, R. (2006). Application of mutated Clostridium difficile Toxin A for determination of glucosyltransferase-dependent effects. Infect Immun 74, 6006–6010.[Abstract/Free Full Text]

Warny, M. & Kelly, C. P. (1999). Monocytic cell necrosis is mediated by potassium depletion and caspase-like proteases. Am J Physiol 276, C717–C724.[Medline]

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