Clostridium difficile Toxin B Causes Apoptosis in Epithelial Cells by Thrilling Mitochondria
2007; Elsevier BV; Volume: 282; Issue: 12 Linguagem: Inglês
10.1074/jbc.m607614200
ISSN1083-351X
AutoresPaola Matarrese, Loredana Falzano, Alessia Fabbri, Lucrezia Gambardella, Claudio Frank, Blandine Geny, Michel R. Popoff, Walter Malorni, Carla Fiorentini,
Tópico(s)Toxin Mechanisms and Immunotoxins
ResumoTargeting to mitochondria is emerging as a common strategy that bacteria utilize to interact with these central executioners of apoptosis. Several lines of evidence have in fact indicated mitochondria as specific targets for bacterial protein toxins, regarded as the principal virulence factors of pathogenic bacteria. This work shows, for the first time, the ability of the Clostridium difficile toxin B (TcdB), a glucosyltransferase that inhibits the Rho GTPases, to impact mitochondria. In living cells, TcdB provokes an early hyperpolarization of mitochondria that follows a calcium-associated signaling pathway and precedes the final execution step of apoptosis (i.e. mitochondria depolarization). Importantly, in isolated mitochondria, the toxin can induce a calcium-dependent mitochondrial swelling, accompanied by the release of the proapoptogenic factor cytochrome c. This is consistent with a mitochondrial targeting that does not require the Rho-inhibiting activity of the toxin. Of interest, the mitochondrial ATP-sensitive potassium channels are also involved in the apoptotic response to TcdB and appear to be crucial for the cell death execution phase, as demonstrated by using specific modulators of these channels. To our knowledge, the involvement of these mitochondrial channels in the ability of a bacterial toxin to control cell fate is a hitherto unreported finding. Targeting to mitochondria is emerging as a common strategy that bacteria utilize to interact with these central executioners of apoptosis. Several lines of evidence have in fact indicated mitochondria as specific targets for bacterial protein toxins, regarded as the principal virulence factors of pathogenic bacteria. This work shows, for the first time, the ability of the Clostridium difficile toxin B (TcdB), a glucosyltransferase that inhibits the Rho GTPases, to impact mitochondria. In living cells, TcdB provokes an early hyperpolarization of mitochondria that follows a calcium-associated signaling pathway and precedes the final execution step of apoptosis (i.e. mitochondria depolarization). Importantly, in isolated mitochondria, the toxin can induce a calcium-dependent mitochondrial swelling, accompanied by the release of the proapoptogenic factor cytochrome c. This is consistent with a mitochondrial targeting that does not require the Rho-inhibiting activity of the toxin. Of interest, the mitochondrial ATP-sensitive potassium channels are also involved in the apoptotic response to TcdB and appear to be crucial for the cell death execution phase, as demonstrated by using specific modulators of these channels. To our knowledge, the involvement of these mitochondrial channels in the ability of a bacterial toxin to control cell fate is a hitherto unreported finding. Today, it is widely accepted that bacterial pathogens can manipulate the eukaryotic machinery to suit their own needs, frequently interfering with pathways controlling apoptotic cell death (reviewed in Ref. 1Fiorentini C. Falzano L. Travaglione S. Fabbri A. Cell Death Differ. 2003; 10: 147-152Crossref PubMed Scopus (51) Google Scholar). Apoptosis occurs, depending on the stimuli, via two major pathways that converge on caspase 3 activation and are initiated by death receptors or alternatively by mitochondria (reviewed in Ref. 2Hengartner M.O. Nature. 2000; 407: 770-776Crossref PubMed Scopus (6275) Google Scholar). Bacteria can exploit both apoptotic pathways, often by producing protein toxins that mediate a long range cross-talk with host cells. In recent years, the targeting of mitochondrial membranes is emerging as a widespread strategy employed by bacterial pathogens in controlling the host cell destiny (reviewed in Ref. 3Blanke S.R. Trends Microbiol. 2005; 13: 64-71Abstract Full Text Full Text PDF PubMed Scopus (43) Google Scholar). The direct targeting of mitochondria, in fact, can allow bacteria to bypass upstream checkpoints of cell death, thus straightforwardly handling one of the central executioners of apoptosis. In this context, mitochondria have very recently been indicated as a key target for toxin activity, especially for certain pore-forming toxins that besides acting at the level of the eukaryotic cell membranes also organize pores in mitochondrial membranes (4Galmiche A. Rassow J. Doye A. Cagnol S. Chambard J.C. Contamin S. de Thillot V. Just I. Ricci V. Solcia E. Van Obberghen E. Boquet P. EMBO J. 2000; 19: 6361-6370Crossref PubMed Scopus (304) Google Scholar, 5Müller A. Gunther D. Brinkmann V. Hurwitz R. Meyer T.F. Rudel T. EMBO J. 2000; 19: 5332-5343Crossref PubMed Scopus (108) Google Scholar, 6Müller A. Rassow J. Grimm J. Machuy N. Meyer T.F. Rudel T. EMBO J. 2002; 21: 1916-1929Crossref PubMed Scopus (72) Google Scholar, 7Haslinger B. Stranfeld K. Peters G. Schulze-Osthoff K. Sinha B. Cell Microbiol. 2003; 5: 729-741Crossref PubMed Scopus (90) Google Scholar). This allows in fine a leakage of proapoptotic factors, such as cytochrome c, in the cytosol and the consequent stimulation of apoptosis. The direct targeting of mitochondria, however, has been reported also for toxins devoid of pore-forming activity, such as those belonging to the large clostridial toxin family (reviewed in Ref. 8Just I. Gerhard R. Rev. Physiol. Biochem. Pharmacol. 2004; 152: 23-47Crossref PubMed Scopus (183) Google Scholar) and in particular for the lethal toxin from Clostridium sordellii (LT) 4The abbreviations used are: LT, C. sordelli lethal toxin; mKATP channel, ATP-sensitive mitochondrial potassium channel; 5HD, 5-hydroxydecanoic acid; TcdA, C. difficile toxin A; TcdB, C. difficile toxin B; WT, wild-type; rec, recombinant; cyt, cytochorome; FCCP, carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone; mAb, monoclonal antibody; MMP, mitochondrial membrane potential; MMHP, mitochondrial membrane hyperpolarization; JC-1, 5-5′,6-6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazol-carbocyanine iodide; TMRM, tetramethylrhodamine ester; OLM, oligomycin; PI, propidium iodide; MOPS, 3-(N-morpholino)propanesulfonic acid; NterTcdB, recombinant N-terminal domain of TcdB; NterLT82, recombinant N-terminal domain of C. sordellii LT82; fmk, fluoromethyl ketone; FITC, fluorescein isothiocyanate. (9Petit P. Breard J. Montalescot V. El Hadj N.B. Levade T. Popoff M.R. Geny B. Cell Microbiol. 2003; 5: 761-771Crossref PubMed Scopus (33) Google Scholar) and toxin A from Clostridium difficile (TcdA) (10He D. Hagen S.J. Pothoulakis C. Chen M. Medina N.D. Warny M. Lamont J.T. Gastroenterology. 2000; 119: 139-150Abstract Full Text Full Text PDF PubMed Scopus (96) Google Scholar). The exact mechanism by which the large clostridial toxins perturb the mitochondrial functionality, however, remains still to be defined. C. difficile toxin B (TcdB) is another large clostridial toxin, largely accepted as a proapoptotic factor (11Fiorentini C. Fabbri A. Falzano L. Fattorossi A. Matarrese P. Rivabene R. Donelli G. Infect. Immun. 1998; 66: 2660-2665Crossref PubMed Google Scholar, 12Qa'Dan M. Ramsey M. Daniel J. Spyres L.M. Safiejko-Mroczka B. Ortiz-Leduc W. Ballard J.D. Cell Microbiol. 2002; 4: 425-434Crossref PubMed Scopus (71) Google Scholar). It is produced, together with TcdA, by pathogenic strains of C. difficile, recognized as the principal cause of antibiotic-associated pseudomembranous colitis (reviewed in Ref. 13von Eichel-Streibe r.C. Boquet P. Sauerborn M. Thelestam M. Trends Microbiol. 1996; 4: 375-382Abstract Full Text PDF PubMed Scopus (215) Google Scholar). TcdA and TcdB are large protein toxins (TcdB, 269 kDa; TcdA, 308 kDa) that encompass three functional domains: the receptor binding domain, the intermediate part responsible for membrane translocation, and the N-terminal part harboring the glucosyltransferase activity. Both toxins are internalized by receptor-mediated endocytosis and require passage through an acidic compartment for activation (13von Eichel-Streibe r.C. Boquet P. Sauerborn M. Thelestam M. Trends Microbiol. 1996; 4: 375-382Abstract Full Text PDF PubMed Scopus (215) Google Scholar). Once in the cytoplasm, they transfer from UDP-glucose a glucose moiety on the threonine 35/37 residue of Rho, Rac, and Cdc42, thus inactivating these GTPases (14Just I. Selzer J. Wilm M. von Eichel-Streiber C. Mann M. Aktories K. Nature. 1995; 375: 500-503Crossref PubMed Scopus (883) Google Scholar). Rho, Rac, and Cdc42 are important regulatory proteins of mammalian cells that control many cellular processes (reviewed in Ref. 15Etienne-Manneville S. Hall A. Nature. 2002; 420: 629-635Crossref PubMed Scopus (3861) Google Scholar), including the assembly/disassembly of the cell cytoskeleton. Cells exposed to these toxins lose their ability to maintain a proper cytoskeleton architecture, undergoing cell retraction and rounding (8Just I. Gerhard R. Rev. Physiol. Biochem. Pharmacol. 2004; 152: 23-47Crossref PubMed Scopus (183) Google Scholar). These two potent cytotoxins differ in certain aspects, TcdB being more powerful, at least on cultured cells, than TcdA (13von Eichel-Streibe r.C. Boquet P. Sauerborn M. Thelestam M. Trends Microbiol. 1996; 4: 375-382Abstract Full Text PDF PubMed Scopus (215) Google Scholar). This work, however, has entirely been focused on TcdB and on its possible interaction with mitochondria. Our results disclose the ability of TcdB to provoke an early hyperpolarization of mitochondria in intact cells that follows a contained rise of cytosolic calcium and precedes the final execution step of apoptosis. Notably, the toxin can induce the swelling of isolated mitochondria in a calcium-dependent fashion, and this effect does not require the enzymatic activity of the toxin. Moreover, we have also shown that the apoptotic response to TcdB involves the mitochondrial ATP-sensitive potassium channels (mKATP channels), which appear to be crucial for the cell death execution phase. These mitochondrial channels, detected in various types of cells, have been reported to play a key role in the protection against different insults, including those leading to apoptosis (16Pozniakovsky A.I. Knorre D.A. Markova O.V. Hyman A.A. Skulachev V.P. Severin F.F. J. Cell Biol. 2005; 168: 257-269Crossref PubMed Scopus (224) Google Scholar). To our knowledge, this finding is hitherto uncharted for a bacterial protein toxin. Cell Cultures—The human epithelial HEp-2 cells were grown in minimal essential medium supplemented with 10% (v/v) fetal calf serum, 50 units/ml penicillin, and 50 μg/ml streptomycin in a humidified atmosphere with 5% (v/v) CO2 at 37 °C. For all experiments, 5 × 105 cells were seeded into 3-cm diameter Petri dishes. Twenty-four hours after seeding, cells were treated as specified (see below). Toxins—TcdB was prepared and purified according to previously published methods (17von Eichel-Streiber C. Harperath U. Bosse D. Hadding U. Microb. Pathog. 1987; 2: 307-318Crossref PubMed Scopus (96) Google Scholar). Recombinant TcdB (rec TcdB, cloned in the PQE 30 plasmid vector with a His6 tag at the N terminus and expressed into the XL1 blue Escherichia coli) and Helicobacter pylori VacA were generous gifts from P. Boquet (Nice). VacA was activated following the procedure previously reported (4Galmiche A. Rassow J. Doye A. Cagnol S. Chambard J.C. Contamin S. de Thillot V. Just I. Ricci V. Solcia E. Van Obberghen E. Boquet P. EMBO J. 2000; 19: 6361-6370Crossref PubMed Scopus (304) Google Scholar). The N-terminal domain (amino acids 1–546) of TcdB was prepared as follows. DNA encoding for the 546 N-terminal amino acids was amplified from C. difficile strain VPI10463 and cloned into pET28a vector. The recombinant N-terminal domain was produced in E. coli strain BL21 DE3 and purified on a cobalt column (Talon; Clontech), according to the manufacturer's recommendations. To test the enzymatic activity of the TcdB N-terminal domain, in vitro glucosylation of recombinant glutathione S-transferase-Rac (1 μg) was carried out in 50 mm triethanolamine, pH 7.5, containing 2 μlof UDP-[14C]glucose (286.2 mCi/mmol; PerkinElmer Life Sciences), 2 mm MgCl2, 1 mm dithiothreitol, 0.3 mm GDP, and variable concentrations of recombinant N-terminal domain of TcdB (NterTcdB) or C. sordellii LT82 (NterLT82). The reaction was performed for 1 h at 37 °C and stopped by adding sample buffer followed by boiling for 3 min. Samples were then electrophoresed on 12% SDS-PAGE and autoradiographed. Treatments—Before starting the experiments, we performed TcdB titration in living cells and tested three different concentrations among those causing an evident cell retraction (the morphological response that precedes the rounding) within 1 h of incubation. For all of the experiments, we have then chosen the concentration of 3 ng/ml toxin B (corresponding to 10-11 m) that induced cell retraction in the whole population within 30 min. HEp-2 cells were treated with 3 ng/ml wild-type (WT) TcdB or rec TcdB for different time lengths (1, 3, 6, 18, and 32 h). Isolated mitochondria (see below for the preparation) were challenged with (i) WT TcdB (1.5, 3, or 6 ng/ml), (ii) rec TcdB (3 ng/ml), (iii) NterTcdB (1.5, 3, 4.5, or 6 ng/ml), and (iv) VacA (5 μg/ml). It is important to underline that, except when compared with rec TcdB, the WT TcdB will be always referred to simply as TcdB. For experiments with inhibitors, cells were pretreated for 30 min with the following compounds before TcdB administration: (i) carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP; 40 nm) (Sigma), an uncoupler of respiratory chain; (ii) oligomycin (OLM; 1 μm) (Sigma), an inhibitor of ATP synthase known to increase mitochondrial membrane potential (MMP); (iii) diazoxide or pinacidil (both 10 μm; Sigma), activators of mKATP channels; (iv) 5-hydroxydecanoic acid (5HD; 10 μm) (Sigma), which blocks the mKATP channel; and (v) monensin (10 μm) (Sigma), a lysosomotropic agent that alters the endosomal pH. Cells treated with each drug (FCCP, OLM, diazoxide, pinacidil, 5HD, or monensin) alone were considered as controls. For caspase inhibitor experiments, cells were pretreated with a 30 μm concentration of the specific inhibitor for 2 h before TcdB exposure. In particular, the following inhibitors were used: IETD-fmk for caspase 8, LEHD-fmk for caspase 9, and DEVD-fmk for caspase 3. As a negative control, we used HEp-2 cells treated with a heat-inactivated (98 °C for 10 min) TcdB. In this case, all of the toxin-induced effects were abolished. Cell Death Assays—Quantitative evaluation of apoptosis was performed by using the following flow cytometry methods: (i) double staining using an FITC-conjugated annexin V/propidium iodide (PI) apoptosis detection kit (Eppendorf, Milan, Italy), which allows discrimination between early apoptotic, late apoptotic, and necrotic cells, and (ii) evaluation of DNA fragmentation in ethanol-fixed cells using PI (Sigma). Activation of Caspases in Living Cells—The activation state of caspase 8, 9, and 3 was evaluated by using the CaspGLOW fluorescein active caspase staining kit (MBL, Woburn, MA). This kit provides a sensitive means for detecting activated caspases in living cells. The assay utilizes specific caspase inhibitors (IETD-fmk for caspase 8, LEHD-fmk for caspase 9, and DEVD-fmk for caspase 3) conjugated to FITC as the fluorescent marker. These inhibitors are cell-permeant and nontoxic and irreversibly bind to the caspase active form. The FITC label allows detection of activated caspases in apoptotic cells directly by flow cytometry. Control and treated HEp-2 cells were incubated with FITC-IETD, FITC-LEHD-fmk, and FITC-DEVD-fmk for 1 h at 37 °C following the manufacturer's instructions. Samples were thereafter washed three times and immediately analyzed on a cytometer by using the FL-1 channel. Two additional experimental controls were also considered: (i) samples prepared by pretreating cells with specific caspase 8, caspase 9, or caspase 3 inhibitor before TcdB administration and (ii) unlabeled HEp-2 cells (negative control). MMP in Living Cells—The MMP of controls and treated-HEp-2 cells was studied by using a 5–5′,6–6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazol-carbocyanine iodide (JC-1; Molecular Probes, Inc., Eugene, OR) probe. In line with this method, living cells were stained with 10 μm JC-1, as previously described (18Cossarizza A. Franceschi C. Monti D. Salvioli S. Bellesia E. Rivabene R. Biondo L. Rainaldi G. Tinari A. Malorni W. Exp. Cell Res. 1995; 220: 232-240Crossref PubMed Scopus (272) Google Scholar). Tetramethylrhodamine ester (TMRM; 1 μm) (Molecular Probes) (red fluorescence) was also used to confirm data obtained by JC-1. Single Cell Assay for [Ca2+]i Recording—Optical fluorimetric recordings with fura-2/AM were used to evaluate the intracellular calcium concentration ([Ca2+]i). Fura-2/AM stock solutions were obtained by adding 50 μg of Fura-2/AM to 50 μlof 75% Me2SO plus 25% pluronic acid. Cells were bathed for 60 min at room temperature with 5 μl of stock solution diluted in 1 ml of extracellular solution (125 mm NaCl, 1 mm KCl, 5 mm CaCl2, 1 mm MgCl2, 8 mm glucose, and 20 mm HEPES, pH 7.35) for a final Fura concentration of 5 μm. This solution was then removed and replaced with extracellular solution, and the dishes were quickly placed on the microscope stage. To measure fluorescence changes, a Hamamatsu (Shizouka, Japan) Argus 50 computerized analysis system was used, recording every 12 s the ratio between the values of light intensity at 340- and 380-nm stimulation. The basal level of [Ca2+]i was estimated as ∼70 nm using the calibration standard kit (Molecular Probes), equivalent to a ratio value of about 0.7. Thapsigargin (100 nm; Alomone Laboratories) was used to evaluate the role played by internal stores in the fast increase of [Ca2+]i. Immunofluorescence-intensified Video Microscopy—After 1 and 18 h of TcdB incubation, control and treated cells were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 30 min at room temperature and then permeabilized with 0.5% Triton X-100 in PBS for 5 min at room temperature. After washing in the same buffer, samples were incubated at 37 °C for 1 h with monoclonal antibodies (mAbs) against mitochondria (Chemicon International, Inc.) and with polyclonal antibodies, prepared as previously described (19Popoff M.R. Infect. Immun. 1987; 55: 35-43Crossref PubMed Google Scholar), specific for LT and cross-reacting with TcdB. After washing, samples were double-labeled with anti-mouse Alexa Fluor 594 (Molecular Probes) and anti-rabbit Alexa Fluor 488 (Molecular Probes). Following extensive washings, samples were mounted with glycerol/PBS (2:1) and observed with a Olympus BX51 fluorescence microscope. Images were captured by a color chilled 3CCD camera (Hamamatsu, Japan), and normalization and background subtraction were performed for each image. Figures were obtained by the OPTILAB (Graftek) software for image analysis. Preparation of Isolated Mitochondria—HEp-2 cells grown in monolayer were harvested by a solution containing 0.25% (w/v) trypsin and 0.02% (w/v) EDTA, in calcium- and magnesium-free PBS and collected by centrifugation. After three washings in PBS, cells were resuspended in Homo-buffer (10 mm Hepes, pH 7.4, 1 mm EGTA, 0.1 m sucrose, 5% bovine serum albumin, 1 mm phenylmethylsulfonyl fluoride, and complete protease inhibitor mixture (Roche Applied Science) and maintained for 10 min on ice. After this time, cells were homogenized with about 100 strokes of a Teflon homogenizer with a B-type pestle as previously reported (20Zamzami N. Maisse C. Metivier D. Kroemer G. Methods Cell Biol. 2001; 65: 147-158Crossref PubMed Google Scholar) for 10 min at 4 °C to remove intact cells and nuclei, and the supernatants were further centrifuged at 10,000 × g at 4 °C for 10 min to precipitate the heavy membrane fractions (enriched in mitochondria). These fractions were then purified by standard differential centrifugation. The mitochondrial pellet obtained was resuspended in swelling buffer containing 0.1 m sucrose, 0.5 m sodium succinate, 50 mm EGTA at pH 7.4, 1 m phosphoric acid (H3PO4), 0.5 m MOPS, and 2 mm rotenone, kept on ice, and used within 2 h from the preparation. Protein content in the mitochondrial preparation was determined by a spectrophotometric method using bovine serum albumin as a standard. The purity of mitochondria preparation was assessed by Western blot, checking subunit I of cytochrome c oxidase (mAb by Chemicon International). The main problem in obtaining a very purified mitochondria preparation is represented by the possible contamination with other intracellular organelles, such as vesicles from the endolysosomal compartment and Golgi apparatus. Thus, before performing the swelling experiments, we tested the purity of our mitochondrial preparation by flow cytometry after staining with mAbs specific to endolysosomal compartment or Golgi vesicle antigens, Rab-5 and GM130, respectively (both from Santa Cruz Biotechnology). Samples were incubated at 4 °C for 1 h, and, after washing, they were labeled with anti-mouse Alexa Fluor 488 (Molecular Probes). After a 45-min incubation at 4 °C, samples were washed and immediately analyzed by a cytometer. As negative and positive controls, we used purified mitochondria incubated with mouse IgG1 immunoglobulin or with mAb to VDAC-1 (Santa Cruz Biotechnology), respectively, followed by anti-mouse Alexa Fluor 488. Swelling Induction in Isolated Mitochondria—Mitochondria (0.5 mg of protein/ml) were resuspended in swelling buffer at the final volume of 1.5 ml. As a general rule, the reagents (toxins and drugs) under investigation were added 5 min after the recording had initiated. Total recording time was 25 min. A useful positive control for these experiments consisted of the addition of 300 μm Ca2+, a Ca2+ concentration that opens the protein transition pore. This causes amplitude swelling that is accompanied by a decrease of ΔΨ and an increase of the outer membrane permeability, leading to the release of proteins (i.e. cytochrome c (cyt c)) that are normally stored in the intramembrane space. The ΔΨ of isolated mitochondria can be quantified by multiple methods. Here, we used a cytofluorimetric analysis after mitochondria staining with 1 μm tetramethylrhodamine/methyl ester/perchlorate (TMRM; Molecular Probes). By this method, the incorporation of dye TMRM was measured in the FL3 channel: low levels of TMRM incorporation (revealed by a decrease of red fluorescence) indicated a low ΔΨ (21Rodolfo C. Mormone E. Matarrese P. Ciccosanti F. Farrace M.G. Garofano E. Piredda L. Fimia G.M. Malorni W. Piacentini M. J. Biol. Chem. 2004; 279: 54783-54792Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar). We herein tested the effects produced on mitochondria ΔΨ by (i) WT TcdB, (ii) rec TcdB, (iii) NterTcdB, and (iv) VacA, either in the presence or absence of 10 μm Ca2+. These measurements were also performed on mitochondria preincubated with diazoxide or FCCP. In parallel, the supernatants of swelling reactions, carried out in the absence of TMRM, were examined by a commercial enzyme-linked immunosorbent assay kit (R&D Systems, Minneapolis, MN) for the cyt c detection. Flow Cytometry Analysis of Isolated Mitochondria—All samples were analyzed with a FACScan cytometer (BD Biosciences) equipped with a 488-nm argon laser. To exclude debris, during analyses, samples were gated based on light scattering properties in the side scattering and forward scattering modes. The red fluorescence emission (due to TMRM dye) of untreated mitochondria was put up in correspondence with the 102 channels and considered as basal emission. Basal emission was recorded for 5 min, and after this time reagents to be tested were added, and the effect was monitored for the following 25 min. Dot plots of red fluorescence emission as a function of the time, obtained in each condition we used, were statistically analyzed by using CellQuest software for a Macintosh computer (BD) in order to determine the percentage of mitochondria with depolarized membrane. Assay for Cytochrome c Release— Release of cyt c was evaluated by a sensitive and specific immunoassay, using a commercial enzyme-linked immunosorbent assay kit (R&D Systems), according to the manufacturer's instructions. The light emitted was quantified by using a microtiter plate reader at 405 nm. Cyt c concentration was expressed as ng/ml. Apoptosis Induced by C. difficile Toxin B Is Mediated by Caspases 9 and 3—Although several lines of evidence have pointed to TcdB as a proapoptotic factor (11Fiorentini C. Fabbri A. Falzano L. Fattorossi A. Matarrese P. Rivabene R. Donelli G. Infect. Immun. 1998; 66: 2660-2665Crossref PubMed Google Scholar, 12Qa'Dan M. Ramsey M. Daniel J. Spyres L.M. Safiejko-Mroczka B. Ortiz-Leduc W. Ballard J.D. Cell Microbiol. 2002; 4: 425-434Crossref PubMed Scopus (71) Google Scholar), the actual mechanism that leads to cell death is not yet precisely defined. Hence, to address this question, we first assayed which of the two main apoptotic pathways was involved. First, we conducted dose-dependent experiments by using different concentrations of TcdB (1.5, 3, and 6 ng/ml). The results obtained (data not shown) indicated that the minimum dose of the toxin able to induce some apoptotic effects within 24 h was represented by 3 ng/ml. In fact, 1.5 ng/ml toxin was completely ineffective in living HEp-2 cells. In light of this, we performed specific time course experiments by using 3 ng/ml TcdB. The time-dependent effects of the toxin in terms of cell death and caspase activation are reported in Fig. 1. After a 1-h incubation of cells with TcdB, the toxin induced the activation of caspase 9 in a low percentage of cells (Fig. 1, middle column,11 ± 1% versus 6 ± 9% in the control). Importantly, neither a significant activation of caspases 8 (not shown) and3(right column;6 ± 2% versus 4 ± 3% in the control) nor cell death, in terms of AV-positive cells (left column), were detectable (right column). After 6 h of TcdB intoxication, a significant (p < 0.01) increase in the percentage of either cells positive to annexin V (left column, third row) or with active caspase 9 (middle column, third row) was evident. By contrast, a small, but not significant (p > 0.05 versus control cells) increase in caspase 3 activation was observed at this time point (right column, third row). After 18 h, the percentage of cells with active caspase 9 (29 ± 4%) and active caspase 3 (11 ± 5%) significantly increased (Fig. 1A), whereas caspase 8 failed to reveal any activation state (not shown). It is noteworthy that at this time, almost 20% of cells showed apoptotic features (Fig. 1, 18 h, left panel). Prolonging TcdB exposure time up to 32 h, a further increase of caspase 9 (54 ± 7%; middle panel) and caspase 3 activation (51 ± 3%; right panel), together with a significant augmentation in the percentage of apoptotic cells, was observed (more than 40%; left panel). Importantly, caspase 8, which defines the receptor-mediated pathway, was also activated in cells after a 32-h exposure to TcdB. The fact that the observed TcdB-induced caspase 9 activation preceded that of caspase 8, allowed us to hypothesize that the toxin may trigger apoptosis in HEp-2 cells via an effect, either direct or indirect, on mitochondria. To verify this assumption, we performed experiments by using specific caspase inhibitors. Results reported in Fig. 1B clearly show that pretreatment with LEHD-fmk (a caspase 9 inhibitor) or DEVD-fmk (a caspase 3 inhibitor) significantly (p < 0.01) prevented the TcdB-induced apoptosis. By contrast, IETD-fmk (a caspase 8 inhibitor) was unable to protect HEp-2 cells from the effects induced by the toxin. These data suggest that TcdB can induce apoptosis via the intrinsic pathway, thus ascribing an ancillary role to caspase 8 in TcdB-induced cell death in our experimental system. Role of Mitochondrial Membrane Potential in TcdB-induced Apoptosis in Living Cells—Therefore, going backward into the pathway controlled by caspase 9, we measured the MMP during time course experiments (0, 1, 18, and 32 h of TcdB treatment) using the JC-1 probe. JC-1, a very sensitive reagent largely used to evaluate MMP changes occurring in apoptosis, allows the measurement of both the earlier events (i.e. hyperpolarization) (22Matarrese P. Di Biase L. Santodonato L. Straface E. Mecchia M. Ascione B. Parmiani G. Belardelli F. Ferrantini M. Malorni W. Am. J. Pathol. 2002; 160: 507-520Abstract Full Text Full Text PDF Scopus (23) Google Scholar) and later events (i.e. depolarization) (18Cossarizza A. Franceschi C. Monti D. Salvioli S. Bellesia E. Rivabene R. Biondo L. Rainaldi G. Tinari A. Malorni W. Exp. Cell Res. 1995; 220: 232-240Crossref PubMed Scopus (272) Google Scholar) state of mitochondria. Strikingly, after 1 h of TcdB incubation (Fig. 2A, second panel), a peculiar change occurred in toxin-treated cells, represented by the mitochondrial membrane hyperpolarization (MMHP) (43 ± 9% versus 15 ± 2% in control cells). This increase of MMP was followed, as a late event, by the characteristic drop of MMP (typical of cell apoptosis) after 18 and 32 h (Fig. 2A, third and fourth panels, respectively). To explore the possible causal link between MMHP and apoptosis in our experimental system, we used two different "mitochondriotropic" agents able to modulate MMP. Pretreatment with a noncytotoxic dose (40 nm)of FCCP, which causes the mitochondrial proton gradient dissipation (23Giovannini C. Matarrese P. Scazzocchio B. Sanchez M. Masella R. Malorni W. FEBS Lett. 2002; 523: 200-206Crossref PubMed Scopus (93) Google Scholar) and abolishes ΔΨ increase (24Skulachev V.P. IUBMB Life. 2005; 57: 305-310Crossref PubMed Scopus (71) Google Scholar), led to a significant reduction of TcdB-induced MMP changes (Fig. 2, B and D). In fact, FCCP inhibited the MMHP found in cells exposed to TcdB for 1 h (Fig. 2B, 7.1%), and, consequently, also the drop of MMP indu
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