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Am J Physiol Renal Physiol 293: F927-F937, 2007. First published June 13, 2007; doi:10.1152/ajprenal.00199.2007
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Pore-forming epsilon toxin causes membrane permeabilization and rapid ATP depletion-mediated cell death in renal collecting duct cells

C. Chassin,1,2,* M. Bens,1,2,* J. de Barry,3,* R. Courjaret,3 J. L. Bossu,3 F. Cluzeaud,1,2 S. Ben Mkaddem,1,2 M. Gibert,4 B. Poulain,3 M. R. Popoff,4 and A. Vandewalle1

1Institut National de la Santé et de la Recherche Médicale, U773, Centre de Recherche Biomédicale Bichat-Beaujon CRB3, Paris; 2Université Paris 7-Denis Diderot, Paris; 3Institut des Neurosciences Cellulaires et Intégratives, UMR7168-LC2 CNRS-Université L. Pasteur, Département de Neurotransmission et Sécrétion Neuroendocrine, Strasbourg; and 4Unité des Bactéries Anaérobies et Toxines, Institut Pasteur, Paris, France

Submitted 26 April 2007 ; accepted in final form 9 June 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Clostridium perfringens epsilon toxin (ET) is a potent pore-forming cytotoxin causing fatal enterotoxemia in livestock. ET accumulates in brain and kidney, particularly in the renal distal-collecting ducts. ET binds and oligomerizes in detergent-resistant membranes (DRMs) microdomains and causes cell death. However, the causal linkage between membrane permeabilization and cell death is not clear. Here, we show that ET binds and forms 220-kDa insoluble complexes in plasma membrane DRMs of renal mpkCCDcl4 collecting duct cells. Phosphatidylinositol-specific phospholipase C did not impair binding or the formation of ET complexes, suggesting that the receptor for ET is not GPI anchored. ET induced a dose-dependent fall in the transepithelial resistance and potential in confluent cells grown on filters, transiently stimulated Na+ absorption, and induced an inward ionic current and a sustained rise in [Ca2+]i. ET also induced rapid depletion of cellular ATP, and stimulated the AMP-activated protein kinase, a metabolic-sensing Ser/Thr kinase. ET also induced mitochondrial membrane permeabilization and mitochondrial-nuclear translocation of apoptosis-inducing factor, a potent caspase-independent cell death effector. Finally, ET induced cell necrosis characterized by a marked reduction in nucleus size without DNA fragmentation. DRM disruption by methyl-beta-cyclodextrin impaired ET oligomerization, and significantly reduced the influx of Na+ and [Ca2+]i, but did not impair ATP depletion and cell death caused by the toxin. These findings indicate that ET causes rapid necrosis of renal collecting duct cells and establish that ATP depletion-mediated cell death is not strictly correlated with the plasma membrane permeabilization and ion diffusion caused by the toxin.

ion diffusion; lipid raft; oncosis

EPSILON TOXIN (ET), which is produced by Clostridium perfringens types B and D, is a potent pore-forming protein toxin responsible for fatal enterotoxemia, mainly in sheep. Overproliferation of clostridial bacteria in the intestine produces large amounts of ET, which enters the bloodstream via the intestinal mucosal barrier and then diffuses into all organs, preferentially accumulating into the brain and kidneys (31). The toxin increases the permeability of the cerebral blood vessels and induces major neurological disorders (13). ET also alters neuronal cells, leading to the release of glutamate (27). In infected domestic animals, the toxin promotes extensive edema in the lungs, which is associated with elevated blood pressure and induces extensive kidney damage, known as "pulpy kidney disease," characterized postmortem by tubule cell necrosis and medullary interstitial hemorrhage (40, 45).

The pathogenesis of the enteroxemia caused by ET is still not understood. Soler-Jover et al. (42) showed that recombinant ET-GFP injected into mice binds to the apical membranes of distal-collecting duct cells and that ET-GFP-infected mice exhibit severe kidney damage, including selective degeneration of the distal nephron. It has been suggested that renal distal-collecting duct cells may possess a specific receptor for ET. ET has been shown to bind to detergent-resistant membrane (DRM) microdomains (or lipid rafts) and to oligomerize to form an heptameric complex in rat synaptosomal and renal Madin-Darby canine kidney (MDCK) cell membranes (28, 29, 34). The toxin can also assemble and form nonselective diffusion channels when inserted into artificial lipid bilayers (35). The resolution of the crystal structure of ET has revealed that the toxin forms a beta-barrel pore exhibiting some structural similarities with aerolysin, another pore-forming toxin (22). Binding to specific membrane receptor(s) remains an absolute prerequisite for toxin toxicity in living cells (22). However, among the many cell lines tested, only MDCK and human renal leiomyoblastoma G-402 cells have been found to retain susceptibility to ET in vitro (34). In MDCK cells, the toxin has been shown to bind specifically to plasma membranes but not to intracellular organelles (34, 36). ET also induces rapid cell death without altering tight junction permeability, the actin cytoskeleton network, or endocytotic processes (36). With regard to the pathophysiological action of ET, two major issues remain to be clarified. First, the relevance to physiology of the insights gained from analyzing the effects of ET on MDCK cells is disputable because these cells have lost the main characteristic features and specific ion transport properties of the distal-collecting duct cells from which they are derived. Second, ET displays a lethal activity (in mice, the LD50 is 100 ng/kg), which is 100-fold lower than that of the structurally related pore-forming toxins such as aerolysin (26). This raised the question of the cause-to-effect link between the membrane permeabilization and induction of death caused by ET.

In this study, we analyzed the effects of ET on ion diffusion processes and the induction of cell death in a highly differentiated murine renal cortical collecting duct principal (CCD) cell line, mpkCCDcl4 (4), which has retained the main characteristics of the parental CCDs from which they were derived (5). We show that the binding and oligomerization of ET in DRMs of mpkCCDcl4 cells stimulated the electrogenic absorption of Na+ and induced a sustained rise in intracellular Ca2+. Furthermore, ET induced a rapid depletion of cellular ATP content, the permeabilization of mitochondrial membranes, and the mitochondrial-nuclear translocation of apoptosis-inducing factor (AIF), a potent, caspase-independent cell death effector (10, 11, 43). However, ET did not induce nuclear fragmentation, although it did trigger rapid cell necrosis. Rather unexpectedly, DRM disruption by methyl-beta-cyclodextrin (MbetaCD), which impairs the membrane effects of ET, had no impact on the rapid fall in cell ATP and subsequent cell death caused by ET. These results indicate that ET, the most potent clostridial toxin after botulinum and tetanus neurotoxins, induces rapid cell necrosis of renal collecting duct cells, which by the way of a mechanism that differs from that reported for many other pore-forming toxin and appears uncoupled to the membrane alterations.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. ET was purified from an overnight culture of C. Perfringens type D strain NCT2062 as described by Petit et al. (34). The purity of ET and epsilon prototoxin (EPT; >90%) was checked by SDS-PAGE electrophoresis. Iodination of the purified toxin and prototoxin was performed as described (34). Experiments were carried out on mpkCCDcl4 cells grown in DMEM:Ham's F-12, 1:1 vol/vol, supplemented with hormones and 2% fetal calf serum at 37°C in 5% CO2-95% air atmosphere, as previously described (4). Cell viability was assayed by the MTT dye assay as described (34). When required, nuclei were stained with Hoechst H33342 [GenBank] (Sigma, St. Louis, MO).

Immunofluorescence studies. Imaging studies were conducted on living cells incubated with 10–7 M Alexa 488-tagged ET, either alone or with 100 nM Mitotracker Deep Red 633 (Molecular Probes, Eugene, OR). Indirect immunofluorescence was also performed on cells fixed with ice-cold methanol, using a purified rabbit anti-AIF polyclonal antibody (1:100 in PBS/0.3% Triton X-100, Cell Signaling Technology, Danvers, MA) and goat anti-rabbit-CY3-conjugated IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) as the secondary antibody. DNA was stained with SYTOX Green (1:500 in PBS, Molecular Probes) for 5 min at room temperature. All specimens were examined using a confocal laser-scanning microscope (CLSM-510-META, Zeiss, Germany).

Transmission electron microscopy. Confluent cells grown on filters were fixed with 2.5% glutaraldehyde in 100 mM cacodylate buffer, pH 7.4, for 20 min at room temperature and processed for transmission electron microscopy as described (4).

Cellular ATP content. The cellular ATP content was determined using the ATP assay kit (Calbiochem-Merck, Darmstadt, Germany) according to the manufacturer's instructions. The medium from confluent cells incubated under various experimental conditions was removed and replaced by 100 µl nucleotide releasing buffer (NRB). Cells were then incubated for 5 min at room temperature with gentle shaking, and 1 µl of ATP monitoring enzyme was added to the resulting cell lysate. To estimate the release of ATP from the cells, the culture supernatant (50 µl) was mixed with 50 µl NRB and 1 µl ATP monitoring enzyme. The sample was read within 1 min using a Beta counter programmed for luminescence mode. An ATP standard curve was generated. Protein concentration was measured using the Bradford protein assay, and the results were expressed as nanogram of ATP per milligram of protein or as percentage of cellular ATP released into the cell supernatant.

DNA gel electrophoresis. Confluent cells, incubated with or without ET, were lysed using proteinase K and RNase A according to standard protocols. Cell lysates were subjected to agarose gel electrophoresis (1.5%), followed by ethidium bromide staining. Molecular weight standards (SmartLadder) were from Eurogentec (Seraing, Belgium).

Preparation of mitochondrial-enriched membranes and mitochondrial transmembrane potential assay. Enriched-mitochondrial fractions from the cytosol fraction of mpkCCDcl4 cells were separated using the two-step centrifugation ApoAlert Cell Fractionation kit (Clontech Laboratories, Palo Alto, CA) according to the manufacturer's instructions. {Delta}{psi}m Was assessed using the fluorescent probe 3,3'-dihexyl-oxacarbocyanine (DiOC6, Molecular Probes). Detached cells (5 x 104 in 100 µl PBS) were incubated with DiOC6 (40 nM in PBS) for 15 min at 37°C. The reaction was blocked by adding 400 µl ice-cold PBS. The fluorescence emitted was monitored by flow cytometry using a Becton Dickinson FACScalibur (Immunocytometry Systems, San Jose, CA).

Immunoblot analysis. Confluent cells or cytosol and mitochondrial-enriched fractions were lysed in 100 µl of 62.5 mM Tris·HCl (pH 6.8), 2% wt/vol SDS, 10% glycerol, and 50 mM DTT and sonicated for 20 s at 4°C. Cell lysates (50 µg) were electrophoresed using 10% SDS-PAGE and transferred to a nitrocellulose membrane. Antibodies against AMPK-{alpha} (1:500, Cell Signaling Technology) and Bax (1:500, Upstate Chemicon, Lake Placid, NY), cytochrome c (1:100) and cytochrome c oxidase subunit IV (COX4, 1:500; Clontech Laboratories), AIF polyclonal antibody (1:1,000, BD Biosciences, San Jose, CA), and beta-actin (1:1,000, Sigma) were used to detect the corresponding antigens. Protein phosphorylation was analyzed using antibodies against phosphorylated AMPK (1:500) and S6 ribosomal protein (1:500; Cell Signaling Technology). Protein bands were revealed using peroxidase-conjugated goat anti-rabbit or sheep-anti-mouse IgG (Jackson ImmunoResearch Laboratories) and detected using the ECL Plus Western blotting detection system (GE Healthcare, Saclay, France).

Preparation of mpkCCDcl4 cell membranes and electrophoresis. Confluent cells were scraped off, rinsed in ice-cold PBS, and centrifuged (300 g, 5 min) at 4°C. Pelleted cells were resuspended and incubated for 10 min at 4°C in 500 µl of sucrose buffer (250 mM sucrose, 3 mM imidazole, 1 mM EDTA, pH 7.2) containing a protease inhibitor cocktail (Complete, Roche, France) and 1 mM PMSF. The cell suspension was then homogenized and centrifuged (900 g, 10 min) at 4°C. The procedure was repeated once on the resulting pellet. The lysed supernatants were pooled and then centrifuged (19,000 g, 50 min) at 4°C. The resulting pellet was resuspended in 100 µl sucrose buffer containing protease inhibitor cocktail and PMSF. Pelleted membranes were resuspended in 25 µl Laemmli buffer and submitted to SDS-PAGE followed by autoradiography.

Preparation of detergent-insoluble membrane fractions and flotation centrifugation on sucrose gradient. Cell membranes were incubated in PBS containing protease inhibitors 1 mM PMSF and 1% Triton X-100 by rotary shaking at 4°C for 30 min. Triton X-100-insoluble pellets, corresponding to DRMs, were resuspended in 200 µl Triton X-100-free PBS. Supernatant proteins were precipitated by adding 700 µl of 6% trichloracetic acid in the presence of 0.4 mg of Na deoxycholate as a carrier. Resuspended pellets and precipitated supernatant proteins were centrifuged (19,000 g, 30 min) at 4°C, and the final pellets were resuspended in 25 µl Laemmli buffer. Separation of DRMs from apical and basolateral plasma membranes was also performed by floating centrifugation on a gradient sucrose (29). Confluent cells grown on filters were incubated for 30 min at 37°C in the presence of apical or basal 125I-ET (10–8 M) and then rinsed twice in ice-cold PBS, scraped off the filters, and centrifuged (300 g, 5 min) at 4°C. Pelleted cells were incubated in 200 µl PBS containing a protease inhibitor cocktail, 1 mM PMSF, and 1% Triton X-100 at 4°C for 30 min by rotary shaking. The lysates were brought to 40% sucrose in a final volume of 500 µl in polyallomer tubes (Beckman Coulter, Fullerton, CA) and overlaid with 3.5 ml of 30% sucrose in PBS, followed by 1 ml of 5% sucrose in PBS. The gradients were centrifuged at 150,000 g for 18 h at 4°C. Fractions (500 µl) were collected from the tops of the tubes and then precipitated as described above and centrifuged (19,000 g, 30 min) at 4°C. Resuspended pellets in 25 µl Laemmli were boiled for 5 min and run on a 10% SDS-PAGE. Membranes were then subjected to Western blotting using an anti-caveolin-1 antibody (1:200, Santa Cruz Biotechnology, Santa Cruz, CA).

Treatment with PI-PLC. Confluent cells transfected or not with a plasmid encoding for the decay accelerating factor (DAF), a GPI-anchored protein, fused to GFP, kindly provided by P. Bocquet and A. Galmiche (15), were preincubated for 2 h with 10 µg/ml cycloheximide to inhibit protein neosynthesis and 6 U/ml of phosphatidylinositol-specific phospholipase C (PI-PLC; Sigma) in DM medium at 37°C (1, 30). mpkCCDcl4 cells were also preincubated with cycloheximide and PI-PLC, as described above, and then with unlabeled ET (10–8 M) for 60 min at 37°C or with purified C. septicum alpha toxin (10–6 M) for 3 h at 37°C. Triton-insoluble membrane fractions were then prepared and subjected to Western blotting. The blots were revealed using immunopurified rabbit antibodies directed against ET or alpha toxin (37).

Electrophysiological studies. The transepithelial electrical resistance (RT), potential (PD), and short-circuit current (Isc) were measured on confluent mpkCCDcl4 cells grown on Transwell filters (0.4-µm pore size, Corning Costar, Cambridge, MA) as described (4). Isc recordings were also performed on sets of confluent cells in which the apical Na+-containing hormonal-free medium (HFM) was replaced by a Na+-free solution (in mM): 156 N-methyl-D-glucamine, 4 KCl, 0.7 MgCl2, 0.4 MgSO4, 1.05 CaCl2, 20 glucose, 8 HEPES, pH 7.4, while the basal side of the filter was bathed with Na+-containing HFM. Whole cell patch-clamp recordings were performed on cells plated and grown on glass coverslips. Patch pipettes with tip resistance of 4 to 6 M{Omega} were filled with an internal solution containing (in mM) 132 K+-gluconate, 1 EGTA/KOH, 2 MgCl2, 2 NaCl, 10 HEPES/KOH, 2 MgATP, and 0.5 GTP. Cells were voltage-clamped to a steady-state holding potential of –60 mV, and the whole cell current was monitored throughout the experiment and sampled at 10 kHz (Axopatch 200A amplifier, Digidata 1322A and pClamp8 suite from Axon Instruments, Foster City, CA). When required, the transmembrane resistance at –60 mV was determined by the square voltage step method under the current clamp mode. Except when ET was added, cell cultures were superfused continuously using an extracellular solution containing (in mM) 130 NaCl, 2.7 KCl, 5 CaCl2, 0.5 MgCl2, 10 HEPES/Tris, 5.6 glucose, and 0.1% BSA (pH 7.4).

Intracellular Ca2+ measurements. [Ca2+]i recordings were performed on cells grown on glass coverslips and loaded with fura-2-AM (2 µM for 20 min). Cells were placed on an inverted epifluorescence microscope (Axiovert, Zeiss, Germany) and continuously superfused (1 ml/min) with Krebs solution containing 5 mM glucose and 10 mM HEPES buffer (pH 7.2). They were illuminated alternately at 350 and 380 nm and image pairs were recorded at 520 nm every 10 s for 20 min. The ratio of the fluorescence intensities (Ex 350 nm/Ex 380 nm) was calculated on a pixel basis for each image pair. The mean value of the fluorescence ratio was normalized for each region of interest, and the changes in [Ca2+]i were estimated without further calibration.

Data presentation and analysis. Results are given as means ± SE from (n) separate experiments. Statistical analyses were done using a two-tailed Student's t-test. P values <0.05 were considered significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
ET binds to plasma membranes of living mpkCCDcl4 cells. Imaging studies revealed that Alexa 488-tagged ET (10–8 M, 30 min) binds to apical and basolateral membranes of mpkCCDcl4 cells grown on filters (Fig. 1A). Alexa 488-tagged ET labeling remained restricted to the apical or basal cell surface membrane domain directly exposed to ET, with no fluorescent label being detected on the opposite basal or apical cell membrane domain (Fig. 1A). Alexa 488-tagged ET binding was prevented by preincubating the mpkCCDcl4 cells with an excess of unlabeled EPT (not shown), suggesting that EPT and ET may share a common membrane receptor. Additional imaging studies with organelle-specific dyes also confirmed that the toxin did not bind to any intracellular organelles (not shown).


Figure 1
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  		Fig. 1. Epsilon toxin (ET) forms large insoluble complexes in mpkCCDcl4 cell membranes. A: confocal micrographs of cells grown on filters and incubated with 10–8 M Alexa 488-tagged ET for 30 min added either on the apical (left) or basal side (right) of the filters. ap, Apical; ba, basal. Scale bar = 10 µm. B and C: SDS-PAGE and autoradiography of cell membrane preparations (10 µg per lane) from confluent cells grown on filters incubated with 125I-ET (10–8 M; B) or with cell membranes at 37°C with 125I-epsilon prototoxin (EPT; lane 1), 125I-ET (lane 2), 125I-ET plus x1-fold excess (lane 3) or x10-fold excess (lane 4) of unlabeled EPT (C). D: SDS-PAGE and autoradiography of whole cell membranes (lane 1), Triton X-100-insoluble (lane 2), and -soluble (lane 3) membrane extracts incubated with 125I-ET. E: sucrose gradient fractionation of Triton X-100-insoluble membrane extracts from cells incubated with apical 125I-ET (10–8 M, 60 min). The 30-kDa ET monomers and 220-kDa ET complexes were recovered in the same fractions (lanes 2–5) containing the detergent-resistant membrane (DRM) marker caveolin-1. F: no 220-kDa labeled band was detected in Triton-insoluble (lanes 1 and 3) and Triton-soluble (lanes 2 and 4) fractions prepared from cells incubated with 10 mM methyl-beta-cyclodextrin (MbetaCD) or 0.4% saponin and then with 125I-ET.

 
ET forms large insoluble complexes in the detergent-resistant plasma membrane microdomains of filter-grown mpkCCDcl4 cells. Previous studies suggested that the formation of membrane pores by ET is correlated to the appearance of toxin heptameric complexes (28, 34). We investigated the ability of ET to form multimeric complexes in both apical and basal membranes of mpkCCDcl4 cells. SDS-PAGE analysis of lysed cell layers and quantification studies (not shown) revealed that 125I-labeled 220-kDa complexes were ~1.5-fold more abundant in the apical than in the basolateral membranes (Fig. 1B). 125I-ET, but not 125I-EPT, formed large insoluble complexes in purified membrane preparations (Fig. 1C, lanes 1 and 2). The formation of 125I-ET complexes was not altered when membranes were incubated with 10–8 M 125I-ET plus 10–8 M unlabeled EPT but was inhibited when membranes were incubated with a 10-fold excess of unlabeled EPT (Fig. 1C, lanes 3–4). When 125I-ET-labeled cells were lysed with Triton X-100, the 125I 220-kDa band was mainly recovered in the Triton X-100-insoluble fraction, and virtually none of it in the Triton X-100-soluble fraction (Fig. 1D, lanes 1–3). SDS-PAGE analysis of Triton X-100-insoluble fraction from mpkCCDcl4 cells incubated with 125I-ET on the apical side of the filters and fractionated by flotation-centrifugation revealed that monomeric 125I-ET and 125I 220-kDa complexes were recovered in fractions enriched with the DRM marker caveolin-1 (Fig. 1E). These results indicate that ET forms complexes in DRMs, mainly on the apical side of renal mpkCCDcl4 cells. Cholesterol is essential for maintaining the integrity of DRMs (6). Pretreating mpkCCDcl4 cell membranes with MbetaCD (10 mM for 60 min at 37°C), an efficient drug that extracts cholesterol from membranes (21), or with saponin (0.4% for 60 min at 4°C), a cholesterol-sequestering drug (41), completely prevented the formation of 125I 220-kDa complexes in both the Triton X-100-soluble and -insoluble fractions (Fig. 1F). However, live mpkCCDcl4 cells treated with MbetaCD did exhibit some Alexa 488-tagged ET labeling; this was much weaker than that of MbetaCD-untreated cells, although some irregular, more intense patches were detectable (not shown), suggesting that ET binding can occur in noncholesterol-enriched areas of membrane.

ET does not bind to GPI-anchored proteins. Previous studies demonstrated that two other pore-forming toxins, areolysin and C. septicum alpha toxin, which share structural similarities with ET, bind to glycosyl phosphatidylinositol (GPI)-anchored protein receptors (1, 16). DRMs microdomains are highly enriched in GPI-anchored proteins (6). This raises the question of whether the putative receptor for ET and EPT is bound to the plasma membrane via a GPI anchor. GPI-anchored proteins can be removed from the cell surface using PI-PLC (23). Rather unexpectedly, after PI-PLC pretreatment with Alexa 488-tagged ET, the labeling of mpkCCDcl4 cells was the same as that of untreated cells (Fig. 2A). The efficiency of the PI-PLC treatment was checked by using mpkCCDcl4 cells transfected with a plasmid encoding for the GPI-anchored tag DAF-GFP. GFP fluorescence was detected in the perinuclear region, in cytoplasmic patches and delineated the plasma membranes of untreated cells, whereas the DAF-GFP tag was removed from the plasma membrane following PI-PLC treatment (Fig. 2B). We then investigated the relationship between GPI-anchored proteins and the formation of 220-kDa complexes. The formation of insoluble 125I-labeled complexes by ET was then compared with that induced by C. septicum alpha toxin, which forms oligomers in DRMs (17) in the same Triton-insoluble membrane preparation of mpkCCDcl4 cells pretreated or not with PI-PLC. As expected, the insoluble complex formed by the alpha toxin was almost undetectable after PI-PLC treatment, whereas the 220-kDa complexes formed by ET were unaffected (Fig. 2C). In addition, when tested with MTT, ET retained the same level of cytotoxic activity in mpkCCDcl4 cells treated with PI-PLC as in untreated cells (data not shown). These results provide evidence that, in contrast to the receptors for structurally related toxins (i.e., aerolysin or C. septicum alpha toxin), the ET receptor is not a GPI-anchored protein.


Figure 2
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  		Fig. 2. PI-PLC does not alter the binding of ET to mpkCCDcl4 cells. A and B: pattern of membranous Alexa 488-tagged ET (10–7 M, 5 min) labeling (A) or DAF-GFP (B) in cells treated without (–PI-PLC) or with 10 µg/ml cycloheximide plus 6 U/ml PI-PLC (+PI-PLC) for 2 h. Scale bars = 10 µm. C: untreated cells and cells preincubated with cycloheximide plus PI-PLC were incubated either with unlabeled ET (10–8 M, 60 min at 37°C) or with C. septicum alpha toxin (10–6 M, 3 h at 37°C). Triton X-100-insoluble membranes fractions were run on a 10% SDS-PAGE and subjected to Western blotting. Multimeric complexes were revealed using specific anti-C. septicum alpha toxin and anti-ET antisera. Membrane blots were also revealed with an anti-beta-actin antibody used as internal standard.

 
ET causes rapid membrane permeabilization in mpkCCDcl4 cells. Confluent cells grown on filters formed tight epithelia with high transepithelial resistance (RT > 3,000 {Omega}·cm2) and negative transepithelial potential (PD ranging from –60 to –80 mV). Adding increasing concentrations of ET to the apical or basal side induced a parallel and dose-dependent decrease in both RT and PD (Fig. 3A). EPT did not affect RT or PD when it was added to the basal side of the cells (Fig. 3B). In contrast, the apical addition of high concentrations of EPT (>10–9 M) induced significant decreases in RT and PD (Fig. 3B). This suggested that inactive EPT was cleaved by protease(s) to form active ET. mpkCCDcl4 cells synthesize a serine protease, mCAP1 (46), which stimulates epithelial Na+ channel (ENaC) activity, the rate-limiting step for Na+ entry into collecting duct principal cells. This finding led us to test whether aprotinin, an inhibitor of serine proteases, could prevent the apical proteolytic cleavage of EPT caused by serine proteases. Preincubating cells for 18 h with 1 mg/ml aprotinin on the apical side significantly decreased PD, an effect that reflects inhibition of the stimulatory action of serine protease on ENaC activity (46), but it also protected mpkCCDcl4 cells against the membrane depolarization caused by subsequent apical addition of EPT (Fig. 3C).


Figure 3
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  		Fig. 3. Dissociated effects of ET and EPT on apical and basolateral cell membrane depolarization. A and B: transepithelial electrical resistance (RT) and potential (PD) were measured on confluent mpkCCDcl4 cells grown on filters and incubated without ({circ}) or with (bullet) ET or EPT (10–10 to 10–7 M), which were added to the apical or basal side of the cell layers. C: RT and PD were measured on confluent cells grown on filters and sequentially incubated without (– Aprotinin) or with (+ Aprotinin) aprotinin for 18 h at 37°C and then without ({circ}) or with (bullet) 10–8 M EPT for 60 min at 37°C. Aprotinin and EPT were both added to the apical medium bathing cells. Values are means ± SE from 4 to 6 separate experiments.

 
ET stimulates Na+ absorption when added to the apical membrane side of filter-grown mpkCCDcl4 cells. The impact of the presence of large ET complexes in plasma membranes on electrogenic ion transport was studied using the short-circuit current (Isc) method. Apical addition of 10–9 M ET caused a twofold increase in Isc within 20 min (Fig. 4A). In contrast, the depolarization of the basolateral membrane produced by adding ET to the basal side of the cells induced a rapid decline in Isc (Fig. 4A). These effects on Isc were completely abolished by apical or basal preincubation of the cells with a 100-fold excess of anti-ET antiserum on the same side as ET (Fig. 4B), whereas applying anti-ET antiserum to the opposite side did not impair the rise in Isc caused by ET (data not shown). Apical addition of the potent ENaC inhibitor amiloride (10–5 M) did not prevent the increase in Isc caused by subsequently adding ET to the apical side of the cells (not shown). This suggests that ET induced the influx of Na+ through cation nonspecific pores. This view is supported by two other observations. 1) The blockade of Na+-K+-ATPase by incubating cells with basal ouabain (5 x 10–5 M) blunted the rise in Isc caused by the subsequent apical addition of ET (Fig. 4C). 2) When the apical Na+-rich HFM was replaced by Na+-free medium (HFM medium), Isc decreased significantly and the apical addition of ET did not significantly increase Isc: 23.3 ± 1.7 µA/cm–2 (n = 7) vs. HFM medium + ET: 38.9 ± 3.8 µA/cm–2 (n = 7), P < 0.01; apical Na+-free medium: 6.2 ± 0.8 µA/cm–2 (n = 6) vs. apical Na+-free medium + ET: 7.9 ± 1.1 µA/cm–2 (n = 6).


Figure 4
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  		Fig. 4. ET induces an increase in short-circuit current (Isc), a decrease in membrane resistance, and a rise in [Ca2+]i in mpkCCDcl4 cells. A: Isc recordings performed on confluent cells before ({circ}) and after adding (arrows) ET ({varepsilon}T, 10–9 M) to the apical side (bullet) or basal side ({blacktriangleup}) of the cell layers. B: Isc recordings performed on cells incubated without ({square}) or with a x100-fold excess of anti-{varepsilon}T antiserum (bullet) for 30 min and then with 10–9 M {varepsilon}T (arrows). Anti-{varepsilon}T antiserum and {varepsilon}T were both added on the same side (apical or basal) of the cell layers. C: Isc recordings performed before ({circ}) and after basal addition of ouabain (5 x 10–5 M; {square}) and subsequent apical addition of ET ({varepsilon}T, 10–9 M, bullet). Values are means ± SE from 4 to 6 experiments. D: representative recordings of membrane potential responses to hyperpolarizing pulses before and during 10–7 M ET application. E: recordings of [Ca2+]i performed on cells loaded with fura-2-AM (2 µM for 20 min) and incubated without (bullet; control) or with 10–7 M {varepsilon}T ({circ}). Values, expressed as normalized ratios of fluorescence intensities, are means ± SE from 4 to 6 experiments. The increase in fluorescence ratio indicates a rise in [Ca2+]i.

 
ET induces an inward current and a rise in [Ca2+]i. Adding ET (10–7 M) to mpkCCDcl4 cells grown on glass, maintained under voltage clamping at a holding potential of –60 mV, induced an inward current which developed in a time-dependent manner parallel to a decrease in the cell membrane resistance (Fig. 4D and inset). The whole cell current change induced by ET, measured as {Delta}I (i.e., current recorded – holding current), appeared after 1 to 3 min of toxin exposure and increased as a function of time: it had an amplitude of –267.6 ± 65.2 pA (n = 8) after 10 min of ET application. These findings are consistent with the fact that ET inserts into membranes and forms nonspecific cationic pores (35). Imaging studies on living fura-2-loaded mpkCCDcl4 cells also showed that ET induced a sustained rise in [Ca2+]i, which was detected after adding ET (Fig. 4E). The time course of this [Ca2+]i rise was similar to that observed for the inward current, suggesting a cause-effect link between the two phenomena.

ET induces rapid ATP depletion and mitochondrial permeabilization. Adding the toxin to the apical side of the cells caused a rapid dose- and time-dependent decrease in cell viability at concentrations of ET above 10–9 M (Fig. 5A), and a concomitant decrease in cellular ATP content, with an almost complete depletion of cellular ATP 30 min after adding concentrations of ET of more than 10–9 M (Fig. 5B). Only a small fraction of total cellular ATP was recovered in the supernatant after incubating for 60 (not shown) or 90 min with either 10–9 M ET (3.6 ± 1.5%, n = 5) or 10–7 M ET (10.4 ± 4.5%, n = 5). Furthermore, adding extracellular ATP (5 or 10 mM) to the culture medium did not protect exposed cells against the loss in cell viability caused by the toxin (not shown).


Figure 5
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  		Fig. 5. ET induces ATP depletion and mitochondrial membrane permeabilization. A and B: cell viability (A) was measured using the MTT test and cellular ATP content (B) was measured on mpkCCDcl4 cells incubated with various concentrations of epsilon toxin ({varepsilon}T). C: Western blot analysis of the expression of AMPK and beta-actin and phosphorylated AMPK (p-AMPK) and ribosomal protein S6 (p-rpS6) from cells incubated without (0) or with {varepsilon}T. D: Western blot analysis of cytochrome c (cyt. C), apoptosis-inducing factor (AIF), and COX4 in cytosol and enriched-mitochondrial fractions from mpkCCDcl4 cells incubated without (–) or with (+) ET. E: electronmicrographs of confluent mpkCCDcl4 cells incubated without (– {varepsilon}T) or with (+ {varepsilon}T) 10–8 M ET for 60 min at 37°C. Bar = 10 µm. Note the integrity of tight junctional complexes and the presence of focal areas of altered mitochondria (arrowheads). F: representative illustrations of {Delta}{psi}m mitochondrial potential analyzed by flow cytometry on cells incubated without (– {varepsilon}T) or with (+ {varepsilon}T) 10–8 M ET for 30 min at 37°C. In this experiment, 38% of the cells exhibited a loss of {Delta}{psi}m. G: confocal micrograph showing the absence of colocalization between the Mitotracker Red (red) and Alexa 488-tagged {varepsilon}T (green).

 
AMP-activated protein kinase (AMPK) is a ubiquitous, heterotrimeric, metabolic-sensing Ser/Thr kinase that is activated by any stress that depletes cellular ATP (18). We therefore tested whether ET could affect AMPK. The toxin did not alter the total amount of AMPK-{alpha} but did induce concentration-dependent phosphorylation of AMPK-{alpha} (Fig. 5C). Calmodulin-dependent protein kinase kinases (CaMKK{alpha} and beta) phosphorylate and activate AMPK, at least in some cell types (19). Preincubating the cells with the CaMKK inhibitor STO-609 (5 µM, 30 min) failed to prevent the loss of cell viability caused by ET (not shown). Activation of AMPK has been found to inhibit the phosphorylation of downstream ribosomal S6 kinases (20). Concomitantly to the increase in p-AMPK-{alpha}, binding of ET to plasma membrane resulted in a complete inhibition of S6 ribosomal protein phosphorylation (Fig. 5C).

We next tested whether ET-induced cellular ATP depletion altered mitochondrial permeability. Incubating mpkCCDcl4 cells with 10–8 M ET for 60 min caused the release of cytochrome c from the mitochondria into the cytosol, while COX4, a constituent of the inner mitochondrial membrane, was only detected in the mitochondrial-enriched membrane fraction, and not in the cytosol fraction of either untreated and ET-treated cells (Fig. 5D). Consistently with the permeabilization of the mitochondrial outer membrane, we observed the disappearance of mitochondrial crests in mpkCCDcl4 cells incubated with 10–8 M ET for 60 min (Fig. 5E) and a significant loss of the {Delta}{psi}m mitochondrial potential (Fig. 5F). These effects cannot have been produced directly by ET, which did not diffuse into the cell, and did not bind to mitochondria (Fig. 5G), but are likely to have been the consequence of the ET-induced ATP depletion.

ET causes mitochondrial-nuclear translocation of AIF and cell necrosis. We next investigated the possible mechanisms responsible for the rapid cell death caused by ET. At all the concentrations tested (10–10 to 10–7 M for 60 min), the toxin failed to induce large-scale DNA fragmentation (Fig. 6A), and no induction of the apoptotic caspase 3 was detected in ET-treated cells (not shown). Moreover, preincubating the cells with 20 µM Z-VAD.fmk, a wide-spectrum caspase inhibitor, did not protect the cells against the rapid loss in cell viability caused by the toxin (not shown). As shown in Fig. 6B, ET stimulated the expression of Bcl-2-associated protein X (Bax), which is known to induce mitochondrial membrane permeabilization and cytoplasmic release of cytochrome c (25, 32). These findings led us to test whether ET also induced nuclear translocation of AIF (43), a mitochondrial caspase-independent cell death effector that has been shown to induce both chromatin condensation and DNA fragmentation (10, 11). Western blot analysis revealed that AIF was present in mitochondrial-enriched fractions from untreated and ET-treated mpkCCDcl4 cells, but not in the cytosol (see Fig. 5D). Immunofluorescence studies demonstrated that AIF had translocated from the mitochondria into the nuclei of cells incubated with increasing concentrations of ET (Fig. 6, C and D). Little or no AIF staining was seen in the untreated cells (Fig. 6C, left), whereas AIF aggregates could be detected in multiple nuclei after incubating for 30–45 min with 10–8 M ET (Fig. 6C, right). In addition, the pan-caspase inhibitor Z-VAD.fmk (20 µM, 30 min) did not prevent the mitochondrial-nuclear translocation of AIF caused by 10–8 M ET (Fig. 6D). Imaging studies also revealed that ET induced in a concentration-dependent manner a marked reduction in the size of nuclei (Fig. 6E). Overall, these findings strongly suggest that ET may promote oncosis (cell necrosis) as a result of a rapid ATP depletion process associated with mitochondrial-nuclear translocation of AIF.


Figure 6
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  		Fig. 6. Effects of ET on DNA fragmentation and AIF distribution. A: nuclear DNA was analyzed by agarose gel electrophoresis from untreated mpkCCDcl4 cells (0) and cells exposed to increasing concentrations of ET for 60 min. B: Western blot analyses of Bax and beta-actin, used as the internal control, in cells without (0) or with 10–8 M ET treatment for various times. C: confocal micrographs of cells dual-stained with an anti-AIF antibody (red) and SYTOX green DNA dye (green). In contrast to untreated cells (left), nuclear AIF staining was detected in multiple nuclei (arrowheads) after incubating for 45 min with 10–8 M ET (right). Bars = 5 µm. D: quantitative analysis of the time course of AIF staining in nuclei from cells incubated with 10–8 M ET for various times or with Z-VAD.fmk (20 µM, 30 min) and then with 10–8 M ET for an additional 60 min. Values are means ± SE from 25 to 35 individual cell recordings from 2 separate experiments. *P < 0.05 vs. time 0 value. E: confluent cells incubated in the absence (none) or presence of various concentrations of ET for 60 min were stained with the nuclear dye Hoechst H33342 (10 µg/ml). ET induced a dose-dependent reduction in the size of nuclei (arrowheads).

 
Extraction of cholesterol from DRMs impairs ion diffusion, but does not prevent ATP depletion and subsequent cell death caused by ET. The cause-to-effect linkage between membrane permeabilization and cell death induction has been established for many pore-forming toxins. Thus we tested whether cholesterol-affecting drugs, which impair ET oligomerization in DRMs, could protect cells against the toxin. Apical addition of MbetaCD (1 and 10 mM, 60 min) produced reduction in the rapid fall in RT and PD caused by the subsequent apical addition of 10–8 M ET (Fig. 7A). Preincubating the cells with MbetaCD also inhibited the rise in Isc (Fig. 7B), and in [Ca+2]i (Fig. 7C), and produced a reduction of 75 ± 12% (n = 6) in the inward ionic current elicited by ET (Fig. 7D), suggesting that the ion diffusion and transport processes stimulated by ET occur mainly in the DRMs. However, MbetaCD treatment, which on its own had no effect on the mitochondrial release of AIF in non-ET-treated cells, did not impair the mitochondrial-nuclear translocation of AIF caused by 10–8 M ET (Fig. 8, A and B). Incubating the cells with 5 mM MbetaCD alone did not alter the ATP content of the untreated cells and had no protective action against the rapid decrease in cellular ATP caused by ET (Fig. 8C). Consistent with these results, MbetaCD (1 and 10 mM) was found to have only a transient protective effect on cell viability for the first 15–30 min after adding 10–8 M ET (Fig. 8D). These findings demonstrate that removing cholesterol from DRMs resulted in a significant inhibition of the cellular entry of Na+ and Ca2+ but is not sufficient to protect cells against the rapid necrosis caused by ET. Moreover, preincubating the cells with 5 mM MbetaCD plus STO-609, Z-VAD.fmk, ouabain, or amiloride also failed to protect the cells against the rapid loss of viability caused by 10–8 M ET (not shown). The potent inhibitor of calpains, ALLN (25 µM, 30 min), used alone or with 5 mM MbetaCD, also failed to prevent cell death caused by the toxin (not shown).


Figure 7
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  		Fig. 7. Lipid raft disruption by MbetaCD reduces ion diffusion caused by ET in mpkCCDcl4 cells. A: RT and PD were measured on mpkCCDcl4 cells incubated without or with MbetaCD (1 or 10 mM, 60 min) and then with ET ({varepsilon}T, 10–8 M, 60 min). B: Isc recordings were performed on cells preincubated without ({circ}) or with 1 mM MbetaCD alone ({square}) and then with 10–9 M {varepsilon}T alone (bullet) or MbetaCD plus {varepsilon}T ({blacksquare}). Both MbetaCD and {varepsilon}T were added to the apical side of the cells. C: time lapse recordings of [Ca2+]i changes were performed on cells loaded with fura-2-AM (2 µM for 20 min). Recordings were performed on untreated cells (none), cells incubated with 1 mM MbetaCD alone, or with 10–7 M {varepsilon}T or MbetaCD plus 10–7 M {varepsilon}T. D: whole cell patch-clamp experiments in voltage clamp mode (HP = –60 mV) showing that the ionic inward current induced by 10–7 M {varepsilon}T (bullet) is strongly reduced when cells were treated for 15 min with 5 mM MbetaCD before adding {varepsilon}T ({circ}). Values are means ± SE from 5–7 separate experiments. *P < 0.05 between groups.

 

Figure 8
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  		Fig. 8. Lipid raft disruption by MbetaCD does not impair the depletion in ATP and subsequent oncosis caused by ET in mpkCCDcl4 cells. A and B: cells were preincubated or not with 5 mM MbetaCD and then with ET ({varepsilon}T) for an additional 45 min. No differences were observed in the expression (A) or in the quantitative analysis of AIF nuclear staining (B). Values are means ± SE of 43–73 individual cell recordings from 3 separate experiments. C: cellular ATP content in cells incubated with 1 mM MbetaCD alone (bullet), {varepsilon}T alone ({blacksquare}), {varepsilon}T plus 1 mM MbetaCD ({triangleup}), or {varepsilon}T plus 10 mM MbetaCD ({circ}) for 90 min at 37°C. D: time course of cell viability assessed by the MTT test on confluent mpkCCDcl4 cells incubated with 10–8 M {varepsilon}T alone ({blacksquare}), {varepsilon}T plus 1 mM MbetaCD ({triangleup}), or {varepsilon}T plus 10 mM MbetaCD ({circ}). Values are means ± SE from 3 separate experiments.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
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 REFERENCES
 
ET is an extraordinarily potent pore-forming toxin, the lethal dose of which in mice is 100-fold lower than that of structurally related, pore-forming toxins such as aerolysin (26). Here, we show that the immortalized mpkCCDcl4 cells (4) remained highly sensitive to ET. ET preferentially binds to the apical membranes of cells grown on filters, and oligomerizes there, and does diffuse intracellularly. We also show that the inactive EPT can be processed to form fully active ET by proteases present in the apical plasma membrane, but not in basal plasma membrane. These findings are of physiological relevance, since the kidney is actively involved in the elimination of ET and in the mouse kidney ET accumulates particularly on the luminal (i.e., apical) side of the distal collecting duct, where it attacks the cells (42). Binding studies strongly suggest that ET and EPT bind to the same mpkCCDcl4 cell membrane receptor. The number of receptors for ET appears to be greater in the apical than basolateral membranes of mpkCCDcl4 cells. A similar asymmetric in vitro distribution of toxin receptors has been already reported for other pore-forming toxins: iota toxin (38), Vibrio cholerae RTX (14), and aerolysin (3). So far, the nature of the ET receptor(s) is still unknown. Many clostridial toxins, such as C. tetani toxin (30), C. septicum alpha toxin, and aerolysin produced by Aeromonas hydrophilia, which display many similarities with ET (1, 12, 16), specifically bind to GPI-anchored proteins. However, PI-PLC, which removes most GPI-anchored proteins (23) and prevents binding and subsequent biological activities of the toxins listed above, does not prevent 125I-ET binding or the subsequent formation of large 125I complexes in mpkCCDcl4 cell membranes. These findings indicate that the receptor for ET is not a GPI-anchored protein receptor. This is consistent with structural studies showing that, despite ET displays remarkable overall structural similarity with aerolysin, its domain II which is involved in the interaction with GPI anchors (24) differs widely from that of aerolysin (9). Preliminary studies using an ET overlay procedure similar to that used by Abrami et al. (1) revealed that ET predominantly bound to a protein with an apparent molecular weight of 42 kDa (not shown). This binding was prevented by incubating membranes with an identified 37-kDa protein previously isolated from MDCK cells using affinity purification with ET linked to Sepharose beads (34). The fact that cholesterol sequestration does not fully abolish the binding of ET-Alexa488 to living mpkCCDcl4 cells suggests that the receptors for ET may be nonexclusively localized in DRMs.

The formation of the toxin heptamer (28, 29, 34) induces membrane permeabilization, which is prevented by the removal of cholesterol by MbetaCD. This suggests that cholesterol-rich lipid raft microdomains certainly act as concentration platforms favoring oligomerization of ET (2). Binding and oligomerization of ET to MDCK cells grown on a solid support induce rapid decrease of intracellular K+ and a concomitant increase in Cl–, Na+, and Ca2+ (34). Here, we show that ET triggers the electrogenic entry of ENaC-independent Na+ driven by the basolateral Na+-K+-ATPase when the toxin is applied to the apical side of mpkCCDcl4 cells grown on filters. The fact that ET had no effect on Isc under apical Na+-free conditions suggests that the toxin induces a cation-permeable pore in the apical membrane of collecting duct cells, which allows the massive entry of Na+ into mpkCCDcl4 cells. Accordingly, the following possible scenario can be proposed for the first phase of the disease: circulating ET and inactive EPT in the bloodstream are filtered by the glomeruli, thus exposing the apical side of the distal collecting duct cells to relatively high amounts of the active toxin. Moreover, the proteases present at the apical surface of the collecting duct cells (i.e., the tubule lumen) may convert the inactive prototoxin into the fully active toxin. ET pores formed in the apical membrane cause a massive intracellular influx of Na+ and are then extruded by basolateral Na+-K+-ATPase, which is responsible for an ensuing sudden increase in Na+ absorption. This hypothesis could explain, at least in part, the sudden rise in blood pressure observed in intoxicated animals (38, 44, 45).

ET also induces a rapid rise in [Ca2+]i which is correlated to the fall in membrane resistance and induction of sustained inward ionic current. Hence, it is conceivable that the Ca2+ ions flow into the cells through the toxin-induced membrane pores. Indeed, ET induces an efflux of K+, sustained influxes of Ca2+ and Na+, and the entry of small molecules into MDCK cells, which would be consistent with the formation of a rather large pore in the cell membrane (35). The possibility that ET may bind and directly activate endogenous Ca2+ conductance is rather unlikely, because the inhibition of heptameric complex formation by cholesterol-affecting drugs protects mpkCCDcl4 cells against a sudden rise in [Ca2+]i and, furthermore, Ca2+ channels are not known to be associated with DRMs. However, we cannot exclude the possibility that ET-induced depolarization of mpkCCDcl4 cells could trigger the opening of endogenous voltage-dependent Ca2+ channels. Further experiments are needed to test this latter possibility, and to find out whether the release of Ca2+ from intracellular Ca2+ stores contributes to the observed increase in [Ca2+]i.

Importantly, our findings reveal that the mechanism by which ET induces rapid cell demise differs from that of other bacterial pore-forming toxins. Under many pathological conditions, apoptosis and necrosis often coexist (33). Interestingly, even though low concentrations of ET activate the apoptotic Bax, they do not induce nuclear cell fragmentation in mpkCCDcl4 cells, but, instead, they induce oncosis. This is reminiscent of a previous report, in which ET-dependent necrosis in MDCK cells was monitored by the entry of propidium iodide and found to correlate with cell mortality (36). This may be due to the fact that these renal collecting duct cells, which exhibit high metabolic and glycolytic activities, are highly susceptible to rapid ATP depletion and subsequent oncosis. The mechanism by which ET induces cell death clearly differs from that caused by chemical agents producing ATP depletion in many other pore-forming toxins. For example, the C. perfringens enterotoxin (CPE), which also causes a rapid increase in [Ca2+]i in intestinal-like CaCo-2 cells, induces DNA fragmentation and caspase- and calpain-dependent apoptosis or oncosis, at low and high doses, respectively (8). In sharp contrast, even at low concentrations ET failed to activate caspase 3 or to induce advanced DNA fragmentation. Consistent with the rapid fall in cellular ATP, ET binding to mpkCCDcl4 plasma membranes leads to the activation of AMPK, which is a sensitive indicator of reduced cellular energy status (19), and subsequent inhibition of many phosphorylation processes, including the phosphorylation of S6 ribosomal protein downstream of AMPK, which reflects inhibition of the TOR pathway, a major stimulus of protein synthesis (7). The ET effects also result in inducing permeabilization of mitochondrial membranes reflected by the release of cytochrome c into the cytoplasm and the stimulated expression of Bax (25, 32), leading to mitochondrial-nuclear translocation of AIF. AIF is a potent apoptogenic protein (43), which seems to be sufficient by itself to cause cell death in a variety of cell systems (39). In response to various stress effectors, AIF exits from the outer mitochondrial membrane and enters the nucleus, where it activates endonucleases that cause nuclear condensation and DNA degradation (10). Although ATP has been shown to be required for AIF-chromatin condensation in cell-free systems (43), nuclear redistribution of AIF can be observed even in a context of ATP depletion (11, 39). The results from imaging studies have identified abundant AIF immunostaining in nuclei from ET-treated cells, very similar to that reported in ATP-depleted OK cells (39). These findings suggest that the mitochondrial permeabilization caused by ET promotes the mitochondrial-nuclear translocation of AIF, homogeneous chromatin condensation and nuclear shrinkage, which are considered to be typical features of oncosis. However, in contrast with many other pore-forming toxins, ET failed to induce DNA fragmentation, suggesting that the energy-dependent processes required for the apoptotic program are rapidly dissipated following the rapid ATP depletion process.

Is the oncosis induced by the toxin directly linked to the increase in [Ca2+]i and/or to the concomitant increase in ion transport through the toxin pores formed in cell membranes? Moderate increase in [Ca2+]i or massive changes in [Ca2+]i secondary to the formation of large plasma membrane pores induced by low or high doses of CPE in intestinal-like Caco-2 cells have been shown to be responsible for calmodulin- and calpain-dependent apoptosis or oncosis, respectively (8). In the present study, calpain and calmodulin inhibitors did not prevent cell death induced by ET, albeit one cannot rule out the possibility that the formation of large-size ET channels (35) may facilitate the leakage of cytoplasmic molecules, thereby impairing the effect of the inhibitors tested. Moreover, even transient activation of calmodulin- and calpain-dependent apoptosis appears unlikely. Indeed, the removal of cholesterol by MbetaCD, which prevented ET oligomerization in lipid rafts and massively reduced Ca2+ and Na+ influx, did not prevent the sudden decrease in cellular ATP or the subsequent oncosis. Therefore, these findings suggest that a massive and sudden rise in [Ca2+]i is not an absolute prerequisite for the induction of cell necrosis by ET. The possibility that some wide cation channels unaffected by MbetaCD treatment could be sufficient to allow a modest rise in [Na+] and [Ca2+]i, leading to the induction of oncosis in renal collecting duct cells, cannot be firmly ruled out. Further studies will be needed to identify the mechanism by which the binding of ET to the plasma membrane of collecting duct cells can trigger cell ATP depletion and subsequent cell necrosis.

To conclude, the cytotoxic effects of ET in mpkCCDcl4 cells are dual and comprised of a pore-forming, cholesterol-dependent phase that occurs in DRMs, and an ATP depletion-induced oncosis that is almost completely resistant to the removal of cholesterol. The renal collecting duct mpkCCDcl4 cells therefore offer an attractive new cell system that could be used to analyze the cytotoxic action of ET, and to identify its membrane receptor.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
These studies were supported by a grant from Sanofi-Aventis France and Bayer Pharma as part of a multiorganism call for proposals (to A. Vandewalle, M. R. Popoff, and B. Poulain). C. Chassin was supported by a PhD student's grant from the Ministère de la Défense (D.G.A./M.R.I.S). R. Courjaret was supported by a grant from Association Française contre les Myopathies (AFM).

Present address of R. Courjaret: Abteilung für Allgemeine Zoologie, Fachbereich Biologie, Universität Kaiserslautern, 67663 Kaiserslautern, Germany.


   ACKNOWLEDGMENTS
 
We thank J. P. Duong van Huyen and M. F. Brelaire for help in electron microscopy. We thank F. Cluzeaud (INSERM U773) and C. Pouzet (IFR02, Faculté de Médecine Xavier Bichat, Paris, France) for expert technical assistance in confocal microscopy.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. Vandewalle, Institut National de la Santé et de la Recherche Médicale U773, Faculté de Médecine Xavier Bichat, BP 416, 16 rue Henri Huchard, F-75870, Paris Cedex 18, France (e-mail: vandewal{at}bichat.inserm.fr)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* C. Chassin, M. Bens, and J. de Barry equally contributed to this work. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Abrami L, Fisaz M, Glauser PE, Parton RG, van der Goot FG. A pore-forming toxin interacts with a GPI-anchored protein and causes vacuolation of the endoplasmic reticulum. J Cell Biol 140: 525–540, 1998.[Abstract/Free Full Text]
  2. Abrami L, van der Goot FG. Plasma membrane microdomains act as concentration platforms to facilitate intoxication by aerolysin. J Cell Biol 147: 175–184, 1999.[Abstract/Free Full Text]
  3. Abrami L, Fivaz M, Glauser PE, Sugimoto N, Zurzolo C, van der Goot FG. Sensitivity of polarized epithelial cells to the pore-forming toxin aerolysin. Infect Immun 71: 739–746, 2003.[Abstract/Free Full Text]
  4. Bens M, Vallet V, Cluzeaud F, Pascual-Letallec L, Kahn A, Rafestin-Oblin ME, Rossier BC, Vandewalle A. Corticosteroid-dependent sodium transport in a novel immortalized mouse collecting duct principal cell line. J Am Soc Nephrol 10: 923–934, 1999.[Abstract/Free Full Text]
  5. Bens M, Chassin C, Vandewalle A. Regulation of NaCl transport in the renal collecting duct: lessons from cultured cells. Pflügers Arch 453: 133–146, 2006.[CrossRef][Web of Science][Medline]
  6. Brown DA, London E. Functions of lipid rafts in biological membranes. Annu Rev Cell Dev Biol 14: 111–136, 1998.[CrossRef][Web of Science][Medline]
  7. Carerra AC. TOR signaling in mammals. J Cell Sci 117: 4615–4616, 2004.[Free Full Text]
  8. Chakrabarti G, McClane BA. The importance of calcium influx, calpain and calmodulin for the activation of CaCo-2 cell death pathways by Clostridium perfringens enterotoxin. Cell Microbiol 7: 129–146, 2005.[CrossRef][Web of Science][Medline]
  9. Cole AR, Gibert M, Popoff M, Moss DS, Titball RW, Basak AK. Clostridium perfringens {varepsilon}-toxin shows structural similarity to the pore-forming toxin aerolysin. Nature Struct Mol Biol 11: 797–798, 2004.[CrossRef]
  10. Daugas E, Nochy D, Ravagnan L, Loeffler M, Susin SA, Zamzami N, Kroemer G. Apoptosis-inducing factor (AIF): a ubiquitous mitochondrial oxidoreductase involved in apoptosis. FEBS Lett 476: 118–123, 2000a.[CrossRef][Web of Science][Medline]
  11. Daugas E, Susin SA, Zamzami N, Ferri KF, Irinopoulou T, Larochette N, Prevost MC, Leber B, Andrews D, Penninger J, Kroemer G. Mitochondrio-nuclear translocation of AIF in apoptosis and necrosis. FASEB J 14: 729–739, 2000.[Abstract/Free Full Text]
  12. Diep DB, Nelson KL, Raja SM, Pleshak EN, Buckley JT. Glycosylphosphatidylinositol anchors of membrane glycoproteins are binding determinants for the channel-forming toxin aerolysin. J Biol Chem 273: 2355–2360, 1998.[Abstract/Free Full Text]
  13. Finnie JW, Blumbergs PC, Manavis J. Neuronal damage produced in rat brains by Clostridium perfringens type D epsilon-toxin. J Comp Pathol 120: 415–420, 1999.[CrossRef][Web of Science][Medline]
  14. Fullner KJ, Lencer WI, Mekalanos JJ. Vibrio cholerae-induced cellular responses of polarized T84 intestinal epithelial cells are dependent on production of cholera toxin and the RTX toxin. Infect Immun 69: 6310–6317, 2001.[Abstract/Free Full Text]
  15. Gauthier NC, Monzo P, Kaddai V, Doye A, Ricci V, Boquet P. Helicobacter pylori VacA cytotoxin: a probe for a clathrin-independent and Cdc42-dependent pinocytic pathway routed to late endosomes. Mol Biol Cell 16: 4852–4866, 2005.[Abstract/Free Full Text]
  16. Gordon VM, Nelson KL, Buckley JT, Stevens VL, Tweten RK, Elwood PC, Leppla SH. Clostridium septicum alpha toxin uses glycosylphosphatidylinositol-anchored protein receptors. J Biol Chem 274: 27274–27280, 1999.[Abstract/Free Full Text]
  17. Hang'ombe MB, Mukamoto M, Kohda T, Sugimoto N, Kozaki S. Cytotoxicity of Clostridium septicum alpha-toxin: its oligomerization in detergent resistant membranes of mammalian cells. Microb Pathog 37: 279–286, 2004.[CrossRef][Web of Science][Medline]
  18. Hardie DG. The AMP-activated protein kinase pathway–new players upstream and downstream. J Cell Sci 117: 5479–5487, 2004.[Abstract/Free Full Text]
  19. Hardie DG, Sakamoto K. AMPK: a key sensor of fuel and energy status in skeletal muscle. Physiology 21: 48–60, 2006.[Abstract/Free Full Text]
  20. Hay N, Sonenberg N. Upstream and downstream of mTOR. Genes Dev 18: 1926–1945, 2004.[Abstract/Free Full Text]
  21. Kilsdonk EP, Yancey PG, Stoudt GW, Bangerter FW, Johnson WG, Phillips MC, Rothblat GH. Cellular cholesterol efflux mediated by cyclodextrins. J Biol Chem 270: 17250–17256, 1995.[Abstract/Free Full Text]
  22. Lesieur C, Vécsey-Semjén B, Abrami L, Fivaz M, van der Goot FG. Membrane insertion: the strategies of toxins. Mol Membr Biol 14: 45–64, 1997.[Web of Science][Medline]
  23. Low MG. Biochemistry of the glycosyl-phosphatidylinositol membrane protein anchors. Biochem J 244: 1–13, 1987.[Web of Science][Medline]
  24. MacKenzie CR, Hirama T, Buckley JT. Analysis of receptor binding by the channel-forming toxin aerolysin using surface plasmon resonance. J Biol Chem 274: 22604–22609, 1999.[Abstract/Free Full Text]
  25. Marzo I, Brenner C, Zamzami N, Jurgensmeier JM, Susin SA, Vieira HL, Prevost MC, Xie Z, Matsuyama S, Reed JC, Kroemer G. Bax and adenine nucleotide translocator cooperate in the mitochondrial control of apoptosis. Science 281: 2027–2031, 1998.[Abstract/Free Full Text]
  26. Minami J, Katayama S, Matsushita O, Matsushita C, Okabe A. Lambda-toxin of Clostridium perfringens activates the precursor of epsilon-toxin by releasing its N- and C-terminal peptides. Microbiol Immunol 41: 527–535, 1997.[Web of Science][Medline]
  27. Miyamoto O, Sumitani K, Nakamura T, Yamagami S, Miyata S, Itano T, Negi T, Okabe A. Clostridium perfringens epsilon toxin causes excessive release of glutamate in the mouse hippocampus. FEMS Microbiol Lett 189: 109–113, 2000.[CrossRef][Web of Science][Medline]
  28. Miyata S, Matsushita O, Minami J, Katayama S, Shimamoto S, Okabe A. Cleavage of a C-terminal peptide is essential for heptamerization of Clostridium perfringens {varepsilon}-toxin in the synaptosomal membrane. J Biol Chem 276: 13778–13783, 2001.[Abstract/Free Full Text]
  29. Miyata S, Minami J, Tamai E, Matsushita O, Shimamoto S, Okabe A. Clostridium perfringens {varepsilon}-toxin forms a heptameric pore within the detergent-insoluble microdomains of Madin-Darby canine kidney cells and rat synaptosomes. J Biol Chem 277: 39463–39468, 2002.[Abstract/Free Full Text]
  30. Munro P, Kojima H, Dupont JL, Bossu JL, Poulain B, Boquet P. High sensitivity of mouse neuronal cells to tetanus toxin requires a GPI-anchored protein. Biochem Biophys Res Commun 289: 623–629, 2001.[CrossRef][Web of Science][Medline]
  31. Nagahama M, Sakurai J. Distribution of labeled Clostridium perfringens epsilon toxin in mice. Toxicon 29: 211–217, 1991.[Medline]
  32. Narita M, Shimizu S, Ito T, Chittenden T, Lutz RJ, Matsuda H, Tsujimoto Y. Bax interacts with the permeability transition pore to induce permeability transition and cytochrome c release in isolated mitochondria. Proc Natl Acad Sci USA 95: 14681–14686, 1998.[Abstract/Free Full Text]
  33. Nicotera P, Melino G. Regulation of the apoptosis-necrosis switch. Oncogene 23: 2757–2765, 2004.[CrossRef][Web of Science][Medline]
  34. Petit L, Gibert M, Gillet D, Laurent-Winter C, Boquet P, Popoff MR. Clostridium perfringens epsilon-toxin acts on MDCK cells by forming a large membrane complex. J Bacteriol 179: 6480–6487, 1997.[Abstract/Free Full Text]
  35. Petit L, Maier E, Gibert M, Popoff MR, Benz R. Clostridium perfringens epsilon toxin induces a rapid change of cell membrane permeability to ions and forms channels in artificial bilayers. J Biol Chem 276: 15736–15740, 2001.[Abstract/Free Full Text]
  36. Petit L, Gibert M, Gourch A, Bens M, Vandewalle A, Popoff MR. Clostridium perfringens epsilon toxin rapidly decreases the membrane barrier permeability of polarized renal MDCK cells. Cell Microbiol 5: 155–164, 2003.[CrossRef][Web of Science][Medline]
  37. Popoff MR. Purification and characterization of Clostridium sordellii lethal toxin and cross-reactivity with Clostridium difficile cytotoxin. Infect Immun 55: 35–43, 1987.[Abstract/Free Full Text]
  38. Richards JF, Mainguy G, Bibert M, Marvaud JC, Stiles B, Popoff MR. Transcytosis of iota toxin across polarized CaCo-2 cell monolayers. Mol Microbiol 43: 901–917, 2002.
  39. Ruchalski K, Mao H, Singh SK, Wang Y, Mosser DD, Li F, Schwartz JH, Borkan SC. HSP72 inhibits apoptosis-inducing factor release in ATP-depleted renal epithelial cells. Am J Physiol Cell Physiol 285: C1483–C1493, 2003.[Abstract/Free Full Text]
  40. Sakurai J. Toxins of Clostridium perfringens. Rev Med Microbiol 6: 175–185, 1995.
  41. Schroeder RJ, Ahmed SN, Zhu Y, London E, Brown DA. Cholesterol and sphyngolipid enhance the Triton X-100 insolubility of glycosylphosphatidylinositol-anchored proteins by promoting the formation of detergent-insoluble ordered membrane domains. J Biol Chem 273: 1150–1157, 1998.[Abstract/Free Full Text]
  42. Soler-Jover A, Blasi J, Gómez de Aranda I, Navarro P, Gibert M, Popoff MR, Martin-Satué M. Effect of epsilon toxin-GFP on MDCK cells and renal tubule in vivo. J Histochem Cytochem 52: 931–942, 2004.[Abstract/Free Full Text]
  43. Susin SA, Lorenzo HK, Zamzami N, Marzo I, Snow BE, Brothers GM, Mangion J, Jacotot E, Costantini P, Loeffler M, Larochette N, Goodlett DR, Aebersold R, Siderovski DP, Penninger JM, Kroemer G. Molecular characterization of mitochondrial apoptosis-inducing factor. Nature 397: 441–446, 1999.[CrossRef][Medline]
  44. Tamai E, Ishida T, Miyata S, Matsushita O, Suda H, Kobayashi S, Sonobe H, Okabe A. Accumulation of Clostridium perfringens epsilon-toxin in the mouse kidney and its possible biological significance. Infect Immun 71: 5371–5375, 2003.[Abstract/Free Full Text]
  45. Uzal FA, Kelly WR, Morris WE, Assis RA. Effects of intraveinous injection of Clostridium perfringens type D epsilon toxin in calves. J Comp Pathol 126: 71–75, 2002.[CrossRef][Web of Science][Medline]
  46. Vuagniaux G, Vallet V, Jaeger NF, Pfister C, Bens M, Farman N, Courtois-Coutry N, Vandewalle A, Rossier BC, Hummler E. Activation of the amiloride-sensitive epithelial sodium channel by the mouse serine protease mCAP1 expressed in a mouse cortical collecting duct cell line. J Am Soc Nephrol 11: 828–834, 2000.[Abstract/Free Full Text]




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