Cd2+-induced cytochrome c release in apoptotic proximal tubule cells: role of mitochondrial permeability transition pore and Ca2+ uniporter
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《美国生理学杂志》
Department of Physiology and Pathophysiology, Faculty of Medicine, University of Witten/Herdecke, Witten, Germany
School of Biological Sciences, University of Manchester, Manchester, United Kingdom
ABSTRACT
Cd2+ induces apoptosis of kidney proximal tubule (PT) cells. Mitochondria play a pivotal role in toxic compound-induced apoptosis by releasing cytochrome c. Our objective was to investigate the mechanisms underlying Cd2+-induced cytochrome c release from mitochondria in rat PT cells. Using Hoechst 33342 or MTT assay, 10 μM Cd2+ induced 5–10% apoptosis in PT cells at 6 and 24 h, which was associated with cytochrome c and apoptosis-inducing factor release at 24 h only. This correlated with previously described maximal intracellular Cd2+ concentrations at 24 h, suggesting that elevated Cd2+ may directly induce mitochondrial liberation of proapoptotic factors. Indeed, Cd2+ caused swelling of energized isolated kidney cortex mitochondria (EC50 9 μM) and cytochrome c release, which were independent of permeability transition pore (PTP) opening since PTP inhibitors cyclosporin A or bongkrekic acid had no effect. On the contrary, Cd2+ inhibited swelling and cytochrome c release induced by PTP openers (PO43– or H2O2+Ca2+). The mitochondrial Ca2+ uniporter (MCU) played a key role in mitochondrial damage: 1) MCU inhibitors (La3+, ruthenium red, Ru360) prevented swelling and cytochrome c release; and 2) ruthenium red attenuated Cd2+ inhibition of PO43–-induced swelling. Using the Cd2+-sensitive fluorescent indicator FluoZin-1, Cd2+ was also taken up by mitoplasts. The aquaporin inhibitor AgNO3 abolished Cd2+-induced swelling of mitoplasts. This could be partially mediated by activation of the mitoplast-enriched water channel aquaporin-8. Thus cytosolic Cd2+ concentrations exceeding a certain threshold may directly cause mitochondrial damage and apoptotic development by interacting with MCU and water channels in the inner mitochondrial membrane.
cadmium; aquaporin; apoptosis-inducing factor; cyclosporin A
CD2+ REPRESENTS AN INCREASINGLY prevalent occupational and environmental hazard. Every year, >15,000 tons of Cd2+ are used to manufacture nickel-cadmium batteries, paints, chemical stabilizers, plating, or alloys (25). The wide environmental distribution of Cd2+ has led to an increased interest in its biological effects and toxicity (58). Cd2+ has no known physiological function but shares some toxic properties with other metals (18, 58).
A focal point for chronic Cd2+ nephrotoxicity is the proximal tubule (PT), in particular its S1 segment (18), which is associated with a general transport defect of the PT that mimics the de Toni-Debré-Fanconi syndrome. Renal dysfunction in the form of proteinuria, aminoaciduria, glucosuria, and phosphaturia has been demonstrated in persons occupationally exposed to Cd2+ (for a review, see Refs. 30 and 56). In fact, human studies indicate that up to 7% of the general population may develop renal dysfunction from Cd2+ exposure (18).
In the body, Cd2+ circulates mainly as a complex with the 6-kDa protein metallothionein (MT) or with the tripeptide glutathione (GSH), which are easily filtered through the glomerulus. Cd-MT is translocated from the primary urinary filtrate into renal PT cells by receptor-mediated endocytosis (31, 54). After degradation of GSH by -glutamyltranspeptidase bound to the apical plasma membrane of PT cells (10), Cd2+ may also be taken up as a free ion through yet unknown apical transport pathways. In a cell line derived from the S1 segment of rat PT, we have previously demonstrated that Cd2+ (10 μM) is taken up by apical pathways, which operate with slow kinetics and saturate at 24 h (16).
When Cd2+ enters the cytosol of PT cells, it may lose its toxic potency by forming a complex with small molecules (e.g., amino acids) or with peptides and proteins that contain SH groups (such as GSH or MTs) and protect the intracellular milieu from oxidative stress. Depletion of these protective factors, as well as the consequence of Cd2+-induced displacement of endogenous redox active metals (Fe2+, Cu2+) (42), results in the formation of reactive oxygen species (ROS) and subsequent damage to critical organelles. If ROS-mediated stress events are not sufficiently balanced by repair processes, PT cells affected by Cd2+ may be induced to undergo cell death via apoptosis or necrosis in vivo (37, 49) and in vitro (51, 52). Indeed, we have previously shown in an S1 segment of a rat PT cell line that low micromolar (5–10 μM) Cd2+ concentrations induce apoptosis, but not necrosis, of PT cells by a process involving ROS (51, 52).
Although a number of stimuli appear to trigger apoptosis, the current dogma is that apoptosis occurs by activation of two major signaling pathways: the death receptor pathway and the death receptor-independent (mitochondrial) pathway, which may also cross talk under certain conditions (26, 60). Despite the differing upstream apoptosis-mediating pathways, there appears to be some degree of convergence at the mitochondrion (12). After a cell death signal, a number of proapoptotic proteins are liberated from the intermembrane space into the surrounding cytosol, which activate downstream pathways. Factors that are normally sequestered in the mitochondria, but are released upon apoptotic signals, include cytochrome c (Cyt c) (38), apoptosis-inducing factor (AIF) (48), and Smac/Diablo (14, 53), as well as a number of procaspases, which appear to be key elements in the cascade of biochemical events leading to apoptosis.
Accordingly, it has been suggested that Cd2+ induces the release of Cyt c by modifying the interaction of members of the Bcl-2 protein family with mitochondrial membranes (27, 28, 35) and/or by activation of the permeability transition pore (PTP) (13, 36). However, there is accumulating evidence that cytotoxic compounds may alter mitochondrial membrane permeability and the release of Cyt c by processes that are independent of these classic mechanisms (1, 2, 20, 45). Thus the molecular processes underlying Cd2+-induced mitochondrial damage and dysfunction that are associated with Cyt c release still await clarification.
Here, we demonstrate that 10 μM Cd2+ induces apoptosis of rat kidney PT cells at 6 and 24 h whereas Cyt c release is observed after 24 h only, when intracellular Cd2+ concentrations may be high enough to interact directly with mitochondria (16). Cyt c release may be partially mediated by direct effects of Cd2+ on mitochondrial permeability pathways; Cd2+ induces swelling (EC50 9 μM) and release of Cyt c from energized mitochondria and mitoplasts isolated from rat kidney cortex. These processes are independent of opening of the PTP. Rather, Cd2+ enters the inner mitochondrial compartment through the mitochondrial Ca2+ uniporter (MCU) to exert its effects, which may include activation of aquaporin H2O channels, resulting in osmotic swelling and release of Cyt c. We therefore propose that direct effects of cytosolic Cd2+ on inner mitochondrial membrane (IMM) permeability pathways may cause, at least in part, mitochondrial dysfunction and therefore contribute to the development of apoptosis in kidney PT cells.
EXPERIMENTAL PROCEDURES
Materials
DMEM:nutrient mixture F-12 (1:1), FBS, penicillin, and streptomycin were all purchased from GIBCO (Carlsbad, CA). Alamethicin, apo-transferrin, bongkrekic acid (BKA), cyclosporin A (CsA), dexamethasone, epidermal growth factor, ethidium bromide (EtBr), insulin, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT), quinine hydrochloride, rotenone, and ruthenium red (RR) were all purchased from Sigma (St. Louis, MO). CdCl2 was from Merck (Darmstadt, Germany). Digitonin, Hoechst 33342 (H-33342) and Ru360 were from Calbiochem (San Diego, CA). The fluorescent indicator FluoZin-1 was obtained from Molecular Probes (Eugene, OR). The mouse monoclonal antibody against Cyt c (7H8.2C12) was from BD Pharmingen (Erembodegem, Belgium), mouse monoclonal antibody raised against human AIF was from Santa Cruz Biotechnology, (Santa Cruz, CA), the rabbit polyclonal antibody against rat aquaporin-8 (AQP8) was from Alpha Diagnostics (San Antonio, TX), the donkey anti-rabbit IgG and the sheep anti-mouse IgG conjugated with horseradish peroxidase were from Amersham Life Sciences (Bucks, UK), and the donkey anti-mouse indocarbocyanin (Cy3)-coupled IgG was from Dianova (Hamburg, Germany). Inhibitors and drugs were dissolved in either water, ethanol, or dimethyl sulfoxide, and in control experiments, solvents were added to PT cells or isolated mitochondria at a concentration not exceeding 0.2%.
Animals
All experiments conducted were in accordance with the Guiding Principles for Research Involving Animals and Human Beings. Male Sprague-Dawley rats (250–300 g) were obtained from Charles River Laboratories (Sulzfeld, Germany) and were given access to food and water ad libitum.
Methods
Cell culture. An immortalized cell line from the S1 segment of rat PT (WKPT-0293 Cl.2) (57) was cultured in DMEM:nutrient mixture F-12 (1:1) supplemented with 10% FBS, 100 U/ml penicillin G and 100 μg/ml streptomycin sulfate, 1.2 mg/ml NaHCO3, 5 μg/ml insulin, 4 μg/ml dexamethasone, 0.01 μg/ml epidermal growth factor, and 5 μg/ml apo-transferrin in 75-cm2 standard tissue culture flasks (Nalge Nunc International, Rochester, NY) at 37°C in a humidified incubator with 5% CO2. Cells were passaged twice a week on reaching confluency.
Detection of apoptosis and necrosis with H-33342 and EtBr by fluorescence imaging. Staining and experiments were conducted as previously described (52). Briefly, control and Cd2+-treated cells were stained with 2 μg/ml H-33342 followed by 5 μg/ml EtBr. After being washed with 0.2 M HEPES, pH 7.4, cells were visualized under ex/em of 350/460 and 518/605 nm for H-33342 and EtBr, respectively, with the Visichrome High Speed Monochromator system (Visitron Systems, Puchheim, Germany), which was connected to a Zeiss Axiovert 200M microscope (Carl Zeiss, Jena, Germany) equipped with a x20 objective. Images were captured using a digital CoolSPAN ES CCD camera (Roper Scientific, Tucson, AZ) and acquired, processed, and analyzed with MetaMorph software (Universal Imaging, Downingtown, PA). Cells from five random microscopic fields of view at x200 magnification were counted per dish, and the average %apoptotic and necrotic cells were calculated.
MTT assay to detect apoptosis and necrosis. The MTT assay is a measurement of cell viability, and hence an indicator of cell death but does not distinguish between apoptosis and necrosis as well as the inhibition of cell growth (40). MTT is a tetrazolium salt that is catalyzed by active succinate dehydrogenase in the mitochondria of living cells into a blue formazan product. Cells (5 x 103) were seeded into each well of six-well plates and given 48 h before treatment with CdCl2. The original method by Mosmann (40) was modified according to Denizot and Lang (11). This entailed the incubation of 1 mg/ml MTT in serum-free medium without phenol red, aspiration of MTT-containing medium, and dissolving of the formazan product with pure isopropanol without HCl. The product was measured at = 560 and 690 nm using the fixed-wavelength program on a Beckman Coulter DU640 spectrophotometer (Beckman Instruments, Fullerton, CA). Reference wavelength values were subtracted from the test wavelength values. The values were normalized to the control, which was equivalent to 0% cell death.
Detection of cytochrome c or AIF release and apoptosis by immuno- and H-33342 staining using fluorescence imaging. WKPT-0293 Cl.2 cells were grown on glass coverslips and fixed in 4% paraformaldehyde/PBS for 30 min at room temperature. All subsequent steps were also carried out at room temperature. Cells were rinsed three times for 5 min with PBS and permeabilized by incubation in PBS containing 1% SDS for 5 min. After three rinses in PBS, coverslips were inverted onto anti-Cyt c (1:600) or anti-AIF antibodies (1:50) and incubated for 2 h in a moist chamber. After a further washing with PBS, coverslips were incubated on drops of Cy3-conjugated IgG (1:600) for 1 h in the dark. Antibodies were diluted in PBS. After three rinses of 5 min in PBS, nuclei of cells were counterstained with 2 μg/ml H-33342 in 0.2 M HEPES, pH 7.4, for 20 min at 37°C, washed once, and subsequently mounted onto glass slides with Vectashield HardSet Mounting Medium (Vector Laboratories, Peterborough, UK). The cells were visualized under ex/em of 550/630 and 350/460 nm for Cy3 and H-33342, respectively, using the Visichrome High Speed Monochromator system connected to a Zeiss Axiovert 200M microscope equipped with a x100 oil-immersion objective. Images were captured using a digital CoolSPAN ES CCD camera and acquired, processed, and analyzed with MetaMorph software. Using the Color Combine function in MetaMorph, the red component of the original Cy3 16-bit images and the blue component of the original H-33342 16-bit images were merged to produce 24-bit images.
Isolation of mitochondria and mitoplasts. Rat kidney cortex mitochondria were isolated by differential centrifugation essentially as described by Ott et al. (43). Mitoplasts were prepared from mitochondria as described by Greenawalt (21) with slight modifications. A portion of freshly isolated mitochondria (100 mg/ml) was incubated on ice with an equal volume of 12 mg/ml digitonin for 15 min with constant vigorous stirring. The digitonin-containing mitochondrial suspension was diluted 1:3 with isolation buffer before centrifugation at 12,000 g for 10 min at 4°C. The pellet was resuspended in isolation buffer to one-half its volume immediately before the first centrifugation, apportioned into aliquots, and centrifuged again at 12,000 g for 10 min at 4°C. The mitoplast pellets were kept on ice and resuspended in hyperosmotic buffer (1.5 M sucrose, 100 mM HEPES, pH 7.4, with Tris) immediately before use. The purity of the mitoplast preparations was determined by measuring the degree of contamination with Cyt c of the fractions obtained by immunoblotting (data not shown). Protein concentration was determined by the method of Bradford (7).
Measurements of mitochondrial oxygen consumption. Only mitochondria with a respiratory control ratio 4 were used for experiments. Oxygen consumption of isolated mitochondria was measured at 30°C with a PC-supported Oroboros high-resolution oxygraph (Anton Parr, Graz, Austria), which comprised an airtight chamber containing an oxygen electrode and a magnetic stirrer (24). The standard incubation medium was composed of 110 mM mannitol, 60 mM KCl, 60 mM Tris, and 10 mM KH2PO4, pH 7.4, with HCl, which was prewarmed to 30°C, and bubbled with air for at least 15 min before measurements. MgCl2 (5 mM), 10 mM sodium succinate, and 1 μM rotenone were added to 1.5 ml of buffer, followed by 0.35 mg/ml mitochondria, and a steady state of endogenous respiration was allowed to be reached. To induce state 3 respiration, 250 μM K+-ADP was added. Once the ADP was exhausted, respiration reverted back to state 4.
Osmotic swelling assay of mitochondria and mitoplasts. Mitochondria and mitoplasts were used for osmotic swelling experiments within 4 h of isolation. For these, 0.7–1.1 mg/ml mitochondria or 0.3–0.5 mg/ml mitoplasts were incubated in 2 ml of either MSH buffer (210 mM mannitol, 70 mM sucrose, 3 mM HEPES, 10 mM sodium succinate, 1 μM rotenone, pH 7.4, with Tris) or KCl buffer (140 mM KCl, 3 mM HEPES, 10 mM sodium succinate, 1 μM rotenone, pH 7.4, with Tris). Inhibitors, when used, were added before mitochondria/mitoplasts. Changes in volume of energized mitochondria/mitoplasts due to colloid-osmotic effects of solute flux into the mitochondrial matrix after addition of salts and modulators were monitored by the change in absorbance at = 540 nm (3). Kinetic measurements were carried out at 25°C in a Beckman DU-640 spectrophotometer equipped with a Peltier constant-temperature chamber and an automatic six-unit sampler (Beckman Instruments). Data were captured and converted using DU-WinConnection Suite software and analyzed using Microsoft Excel and SigmaPlot 8.0 (SPSS, Chicago, IL). Rates of swelling (absorbance540 nm/min) were calculated from the initial linear portion of mitochondrial absorbance curves. Swelling curves were normalized to maximal mitochondrial swelling. To determine maximal swelling, mitochondria were suspended in KCl buffer, and the maximal absorbance changes induced by the monovalent cation ionophore alamethicin were defined as 100% swelling. Alamethicin-induced absorbance changes amounted to 53 ± 1.4% of initial absorbance. To simplify calculations, maximal swelling was assumed to be 50% of initial absorbance, and normalized values were obtained from the formula {1 – [absorbance/(initial absorbance/2)]} x 100.
Immunoblotting of mitochondrial proteins for Cyt c and AQP8. For immunoblotting, mitochondrial suspensions were collected after swelling measurements and centrifuged at 10,000 g for 5 min at 4°C. Both pellets and supernatants were sonicated on ice for 3 x 5 s at 10 A. Proteins were separated by SDS-PAGE on 15% acrylamide Laemmli minigels and transferred onto polyvinylidene difluoride membranes overnight at 4°C. Blots were blocked with 3% nonfat dry milk and incubated overnight at 4°C with primary anti-Cyt c (1:250) or anti-AQP8 (1:400) antibody. After incubation with horseradish peroxidase-conjugated secondary antibody (1:10,000) for 1 h at 4°C, blots were developed using Western Lighting Plus chemiluminescence reagents (PerkinElmer Life Sciences, Boston, MA), and signals were visualized on X-ray film. Blots were scanned digitally using a Biorad GS 700 Densitometer apparatus and analyzed with Molecular Analyst software (Bio-Rad Laboratories, Hercules, CA).
Cd2+ uptake measurements in mitoplasts with the fluorescence probe FluoZin-1. To measure Cd2+ uptake into mitoplasts, the probe FluoZin-1 (ex = 490 nm, em = 517 nm) was used, which is virtually nonfluorescent in the absence of metals but undergoes an up to 200-fold fluorescence increase upon saturation with micromolar concentrations of metals, such as Zn2+ or Cd2+, with minimal interference by Ca2+ (19). The fluorescence decreases again upon dissociation of Cd2+ from its binding site on the membrane-impermeant indicator. In preliminary experiments, a Kd for Cd2+ of 97 μM was determined (data not shown). Freshly isolated mitoplasts (0.3 mg/ml) were added to 3 ml of buffer containing 110 mM mannitol, 60 mM Tris, 60 mM KCl, pH 7.4, with HCl, as well as 2 μM FluoZin-1 and energized with 10 mM sodium succinate and 1 μM rotenone. Measurements were performed in an LS50B luminescence spectrophotometer (PerkinElmer, Wellesley, MA), and data were collected using the PerkinElmer FL Data Manager program. After stabilization of the fluorescence signal, 20 μM Cd2+ was added every 400 s. As a control, the experiments were repeated in the absence of mitoplasts.
Statistical Analyses
Unless otherwise indicated, all experiments were repeated at least three times with different preparations of mitochondria. Representative data or means ± SE are shown. Statistical analysis using unpaired Student's t-test was carried out with the Sigma Plot 8.0 spreadsheet program. For more than two groups, statistical differences were compared using one-way ANOVA assuming equality of variance with Levene's test and Tukey's post hoc test for pairwise comparison. Statistical analysis was carried out with the SPSS 11.0 program. Results with P 0.05 were considered to be statistically significant. To obtain EC50 values, dose-response curves of Cd2+-induced mitochondrial swelling were fitted using the Sigma Plot 8.0 spreadsheet program assuming a sigmoidal dose response (variable slope).
RESULTS
Cd2+-Induced Apoptosis of PT Cells Is Associated with Cyt c Release at 24 h But Not at 6 h
When cultured PT cells are incubated with the toxic metal Cd2+ at low micromolar concentrations, cell death occurs. To distinguish between apoptosis and necrosis, cell death was determined using the lipophilic DNA dye H-33342, which permeates intact plasma membranes and can therefore label all cells including apoptotic cells (Fig. 1A), whereas the charged DNA dye EtBr can only permeate necrotic cells that have lost their plasma membrane integrity. Control cells at both 6 and 24 h exhibited round, flat, pale blue nuclei (Fig. 1A, left) and negligible apoptosis or necrosis (Fig. 1B). In contrast, 10 μM Cd2+ treatment caused a significant increase of apoptosis but no change in necrosis (Fig. 1B). Apoptosis was defined by fragmentation of the nucleus (asterisk in Fig. 1A) or by condensation (arrow in Fig. 1A). Condensed nuclei have increased fluorescence intensity and are smaller compared with controls. Moreover, they display sharper nucleic outer edges. Quantification of cell death shows that 10 μM Cd2+ induces a significantly higher incidence of apoptosis at 6 or 24 h but not of necrosis (Fig. 1B), which is in accordance with previous data of Cd2+-induced apoptosis in PT cells (16, 52). At 6 h, 10 μM Cd2+ induced 4.2 ± 0.5% apoptosis compared with controls (0.7 ± 0.1%, n = 6). After 24-h Cd2+ treatment, there was 4.9 ± 0.7% apoptosis, which was significantly higher than in the controls (0.8 ± 0.2%, n = 6). The slight increase in apoptosis by Cd2+ after 24 h compared with 6 h was not significant (P = 0.43). Cd2+ (50 μM) further increased the rate of apoptosis at both 6 (7.9 ± 1.2%, n = 6) and 24 h (13.6 ± 2.0%, n = 6). With 50 μM Cd2+, the percentage of necrosis was 1.8 ± 1.3% (n = 6) at 6 h and 8.5 ± 2.5% (n = 6) at 24 h. Necrosis was significantly increased by 50 μM Cd2+ at 24 h compared with controls (P < 0.01). We then used an additional independent technique to measure cell death induced by 10 μM Cd2+, namely, the MTT assay. As shown in Fig. 1C, PT cells incubated with 10 μM Cd2+ for 6 h showed 7.1 ± 1.0% (n = 6) cell death, whereas incubation with 10 μM Cd2+ for 24 h developed 9.1 ± 1.6% (n = 6) cell death, which is of the same magnitude as that obtained with H-33342 and EtBr. The difference between 6 and 24 h was not significant. Moreover, the results also confirm previous published data obtained using annexin V-Alexa 568 staining (49).
Because apoptosis mediated by cytotoxic stimuli including Cd2+ has been associated with mitochondrial release of proapoptotic factors, such as Cyt c or AIF (12, 13, 27, 35), we investigated the release of these factors from mitochondria in PT cells at 6 and 24 h. In Fig. 1D, PT cells were treated with 10 μM Cd2+ for 6 or 24 h and subsequently stained for Cyt c or AIF using specific antibodies and for occurrence of apoptosis with H-33342. In the controls (Fig. 1D, left), the cells have round, pale blue nuclei and punctate Cyt c distribution, as demonstrated by the brighter particulate staining in the cytosolic compartment, indicating that Cyt c is localized to mitochondria. When the cells are treated with Cd2+ for 6 h (Fig. 1D, top right), there is the occurrence of apoptosis as exemplified by the fragmented nucleus, but the cytosolic bright particulate staining of Cyt c remains largely the same as in the control (Fig. 1D, top left). However, after 24-h Cd2+ treatment, apoptotic cells display a major change in Cyt c distribution (Fig. 1D, middle right), compared with the control (Fig. 1D, middle left), which has changed from punctate to diffuse, indicating that Cyt c has been released from mitochondria into the cytosol. Furthermore, the change in Cyt c distribution also seems to occur in cells without condensed or fragmented nuclei (Fig. 1D, middle right), suggesting that Cyt c release from mitochondria may take place before DNA condensation/fragmentation. To confirm the data obtained with Cyt c, we investigated AIF, which is another proapoptotic mitochondrial factor that is released into the cytosol as a result of the loss of integrity of the outer mitochondrial membrane and translocates directly to the nucleus to induce chromatin condensation and high-molecular-weight DNA fragmentation, culminating in apoptosis (48). In control cells, AIF had a well-defined punctate distribution, demonstrated by the brighter particulate staining in the cytosolic compartment (Fig. 1D, bottom left), indicating that AIF was localized to mitochondria. After 6-h exposure to 10 μM Cd2+, the pattern of AIF distribution remained unchanged (data not shown). In contrast, after 24-h incubation with 10 μM Cd2+, apoptotic cells displayed a diffuse AIF distribution, indicating that AIF has been released from mitochondria into the cytosol (Fig. 1D, bottom right).
Cd2+ Induces Swelling and Release of Cyt c in Isolated Mitochondria
Strikingly, recent work from our laboratory has demonstrated that cultured confluent PT cells exposed to 10 μM Cd2+ take up 109Cd2+ through apical transport pathways with slow kinetics, which appeared to have reached equilibrium at 24 h after an approximately fourfold increase in intracellular Cd2+ concentration compared with a 4-h incubation (16). Because the release of Cyt c and AIF observed at 24 h (Fig. 1D) correlated with the increase in intracellular Cd2+ concentration previously described (16), we speculated that when cytosolic Cd2+ exceeds a threshold concentration, it may directly damage mitochondria and induce Cyt c release, which is an integral step of the mitochondrial apoptotic pathway. To test this hypothesis, mitochondria were isolated from rat kidney cortex (rather than from cultured PT cells to obtain a higher yield) and incubated with relevant micromolar Cd2+ concentrations. Figure 2 shows the concentration-dependent swelling of rat kidney cortex mitochondria suspended in MSH buffer induced by Cd2+ with an EC50 of 9 μM (Fig. 2B), which is in accordance with Cd2+-induced apoptosis of PT cells (51, 52). Similar results were also obtained by using KCl buffer (EC50 = 8.4 ± 0.7 μM, n = 8) (data not shown). To investigate whether swelling of kidney cortex mitochondria induced by Cd2+ is associated with Cyt c release, samples were collected for separation of the mitochondrial pellet and supernatant after addition of Cd2+. Figure 3A demonstrates that Cd2+-induced swelling of isolated mitochondria suspended in MSH buffer is accompanied by the release of Cyt c into the supernatant. Mitochondrial preparations were only taken into account for analysis when the controls showed a release of 15% of total Cyt c. Under these conditions, Cd2+ concentrations between 5 and 50 μM significantly increased Cyt c release into the supernatant (Fig. 3B). Data obtained by using KCl buffer showed a consistently high basal Cyt c release in the controls (MSH: 13.1 ± 4.5%, n = 8 vs. KCl: 68.9 ± 3.9%, n = 4). Therefore, no significant difference between controls and Cd2+-exposed mitochondria were obtained (data not shown). The high basal Cyt c release in KCl buffer may be accounted for by the effect of the ionic strength of KCl on a pool of Cyt c that is attached to the IMM by a loose electrostatic interaction (8) and/or centrifugation artifacts in the context of the separation of the mitochondrial pellet and supernatant in KCl buffer.
Cd2+-Induced Mitochondrial Swelling and Cyt c Release Occur Independently of PTP Opening
Figure 4 shows the effect of 1 μM CsA, a potent inhibitor of the PTP (9), on mitochondrial swelling induced by 20 (Fig. 4A) or 50 μM Cd2+ (Table 1) in MSH or KCl buffers (Table 1). As can be seen from the data summarized in Table 1, there was no significant inhibition of the initial rates of swelling when 1 μM CsA was added before the addition of Cd2+. BKA (5 μM), another drug that blocks the PTP by binding to the adenine nucleotide translocator (ANT) (22), did not inhibit Cd2+-induced swelling either (Table 1), suggesting that mitochondrial swelling induced by Cd2+ is not mediated by PTP opening. In line with these observations, we showed that Cyt c release induced by 50 μM Cd2+ was not prevented by 1 μM CsA (Fig. 4B).
View this table:
Cd2+ Prevents Activation of the PTP and Subsequent Cyt c Release
Rather than induce PTP opening, Cd2+ actually inhibited mitochondrial swelling triggered by 5 mM PO43–, an inducer of the PTP (34; for a review, see Refs. 4 and 23) in a concentration-dependent manner (Fig. 5A). To ensure that the inhibition was not caused by an indirect effect of Cd2+ via interaction with the mitochondrial phosphate translocase (23), 2 mM H2O2+50 μM Ca2+, which also induce PTP opening (4, 23), were tested. Again, mitochondrial swelling induced by H2O2 was inhibited as a function of Cd2+ concentration (Fig. 5B). Mitochondrial swelling induced by 5 mM PO43– or 2 mM H2O2+50 μM Ca2+ was strongly inhibited by 1 μM CsA. Moreover, both the Cd2+- and CsA-mediated inhibition of PTP opening were not additive, indicating that both Cd2+ and CsA have the same target of inhibition (data not shown). The inhibitory effect of Cd2+ on PTP opening was also associated with an inhibition of Cyt c release. Mitochondria exposed to 5 mM PO43– released the majority of their Cyt c into the supernatant. In contrast, 50 μM Cd2+ reduced PO43–-induced Cyt c release into the supernatant (Fig. 5C).
Blockers of MCU Prevent Cd2+-Induced Swelling and Cyt c Release
Table 1 shows that the noncompetitive MCU inhibitors, RR and its active component Ru360, as well as the competitive inhibitor La3+ (for a review, see Ref. 4) strongly inhibited Cd2+-induced swelling of kidney cortex mitochondria. Dixon plots of the effects of RR and La3+ on Cd2+-induced mitochondrial swelling in MSH buffer (data not shown) demonstrated that inhibition is noncompetitive and competitive, respectively, as has already been reported for Ca2+ (59). Furthermore, a maximal inhibitory concentration of RR (1–5 μM) prevented the decrease in Cyt c contents in the mitochondrial pellet induced by 50 μM Cd2+ (Fig. 6, A and B). This indicates that Cd2+-induced mitochondrial swelling and Cyt c release are mediated by Cd2+ influx through the RR- and Ru360-sensitive MCU of the IMM. Moreover, Cd2+ inhibition of PTP opening induced by PO43– was released by coincubation with the MCU blocker RR. As shown in Fig. 6C, 1 and 5 μM RR partially restored mitochondrial swelling induced by 5 mM PO43– in the presence of Cd2+, suggesting that the Cd2+ concentration in the inner mitochondrial compartment rather than the outside Cd2+ concentration determines the extent of PTP inhibition.
Evidence for Cd2+Uptake by Mitoplasts from Rat Kidney Cortex
The fluorescent indicator FluoZin-1 is used as a probe to measure the divalent metal Zn2+ with a Kd of 8 μM (19) but is also used as a sensitive indicator of Cd2+ at micromolar concentrations (Molecular Probes). We used membrane-impermeant FluoZin-1 to monitor extramitochondrial changes in Cd2+ concentration. After addition of mitoplasts, the fluorescence intensity decreased slightly (possibly a dilution effect). Addition of 60 nmol Cd2+ induced a small increase in fluorescence signal (solid line in Fig. 7A), whereas a further addition of Cd2+ produced a much larger signal increase. The peak fluorescence signal decreased with time until a plateau was reached after 2–3 min. The decrease in peak fluorescence was observed only in the presence of mitoplasts (compare solid and dotted lines in Fig. 7A) and reflected uptake of Cd2+ from the medium into mitoplasts. With further additions of Cd2+, the rate of change of peak fluorescence diminished further (Fig. 7B), possibly due to saturation of the uptake process.
Cd2+-Induced Mitochondrial Swelling Involves Activation of an Ag+-Sensitive H2O Channel
The magnitude of osmotic swelling induced by Cd2+ cannot be explained by solute and H2O flux through an open PTP, because it is inhibited by Cd2+ (see Figs. 4A and 5 and Table 1). Furthermore, the osmolarity of micromolar Cd2+ entering the matrix space through the MCU is too small to account for the swelling effects. Therefore, we hypothesized that a hydraulic permeability distinct from the PTP may be activated by Cd2+ to mediate H2O flux into the osmotically active matrix space and induce osmotic swelling. The H2O channel protein AQP8 has been detected in hepatocyte mitochondria (17) and is also expressed intracellularly in rat kidney PT (15). Indeed, immunoblotting of membrane fractions of rat kidney cortex with an anti-rat AQP8 antibody revealed a protein band of 28 kDa that was enriched in mitochondria compared with homogenate and was enriched even further in mitoplasts (Fig. 8A). Moreover, the aquaporin inhibitor AgNO3 (5 μM) (41) nearly abolished Cd2+-induced osmotic swelling of mitoplasts (Fig. 8B).
DISCUSSION
What is the Mechanism of Cd2+-Induced Cyt c Release in PT Cells
In the present study, the release of Cyt c or AIF induced by 10 μM Cd2+ in cultured PT cells was observed at 24 h but not at 6 h (Fig. 1D), despite the occurrence of similar rates of apoptosis at both time points (Fig. 1, B and C). This suggested that different processes are responsible for Cd2+-induced apoptosis at 6 and 24 h, and thus we investigated the mechanisms of Cyt c release in more detail. We have previously demonstrated that uptake of Cd2+ by confluent cultured PT cells is slow and saturates at 24 h (16). Therefore, we wondered whether free cytosolic Cd2+ exceeding a threshold concentration may directly affect mitochondria to induce the release of Cyt c and AIF. Here, we provide evidence that Cd2+ induces Cyt c release by acting directly on mitochondrial permeability pathways in the IMM. The data obtained in Figs. 2 and 3 suggest that concentrations of Cd2+ >2 μM may contribute to Cyt c release and thus to apoptosis of cells exposed to 10 μM Cd2+, which may become relevant when intra- and extracellular Cd2+ concentrations are near equilibrium (16). However, other mechanisms may also contribute to apoptosis induced by Cd2+. ROS are generated during Cd2+ exposure of PT cells for 4–8 h (51, 52) and may therefore activate PTP opening (13, 36). Interestingly, concentrations of 5–10 μM Cd2+ inhibit PTP opening induced by H2O2 (Fig. 5), which could explain why no major Cyt c release is observed at 6 h in cultured PT cells (Fig. 1C), even when ROS are elevated (51, 52). In addition, it has been shown that Cd2+ induces the release of Cyt c by modifying the interaction of members of the Bcl-2 protein family with mitochondrial membranes (27, 28, 35). It is therefore likely that several processes contribute to Cd2+-induced Cyt c release in PT cells, depending on time and Cd2+ concentrations. Moreover, recent data from our laboratory demonstrate that exposure of PT cells to 10 μM Cd2+ induces activation of caspases 3 and 9 at 24 h, but not at 6 h, which nicely corresponds with the observation of Cyt c release at 24 h. Caspase-independent apoptosis at early time points (3–8 h) turns out to be mediated by activation of the Ca2+-activated proteases calpains (W.-K. Lee and F. Thévenod, unpublished observations).
How Does Cd2+ Affect Permeability of the IMM
In general terms, temporary or permanent alterations of the integrity of the IMM and/or outer mitochondrial membrane induce the release of proapoptotic proteins (6, 33, 39). Various factors have been implicated in alterations of mitochondrial membrane permeability, such as members of the Bcl-2 protein family (46, 47, 55) and/or ROS-triggered opening of the PTP, which may consist of both IMM and outer mitochondrial membrane proteins, but whose molecular identity is still elusive (32).
Several major conclusions can be drawn from this study. First, Cd2+ does not induce opening of the PTP. Typical inhibitors of PTP, such as CsA and BKA, had no significant inhibitory effect on Cd2+-induced mitochondrial swelling and Cyt c release (Fig. 4, Table 1). Moreover, rather than open the PTP, Cd2+ concentrations 5 μM inhibited opening of the PTP induced by Ca2++H2O2 or PO43– (Fig. 5), which are characteristically operative during hypoxia or oxidative stress. If the PTP is not involved in Cd2+-induced osmotic swelling of mitochondria suspended in MSH buffer, the only likely alternative mechanism that can account for Cd2+-induced swelling must be the activation of an hydraulic conductance because the osmolality of Cd2+ entering the mitochondrial matrix through the MCU is too small to explain the observed magnitude of mitochondrial swelling (15–20% absorbance change). As a matter of fact, we found AQP8 to be expressed in the IMM (Fig. 8A), which confirms recent data in hepatocytes (17). The transmitochondrial membrane driving force for swelling is induced by the osmotic gradient provided by the mitochondrial matrix that consists of anionic proteins, monovalent cations, intermediates of the Krebs cycle, and other small organic molecules (3). With the use of AgNO3, a more potent inhibitor of aquaporins than Hg2+ (41), Cd2+-induced swelling of mitoplasts was abolished (Fig. 8B). Possibly, other hydraulic permeability pathways in the IMM may also be affected by Cd2+. Thus we show that Cd2+ can alter IMM permeability pathways and induce the release of Cyt c by processes that are independent of the PTP, which confirms that PTP opening is not the only mechanism underlying mitochondrial swelling (1, 2, 20, 45). At the present stage, we can only speculate about the mechanism of Cd2+-induced activation of AQP8, but cysteine residues are important for AQP channel function and inhibition by Hg2+ (44) and could be involved in the alteration of the AQP8-associated permeability of the membrane by Cd2+.
Which Pathway Mediates Cd2+ Entry into the Inner Mitochondrial Compartment
The key element to an understanding of how Cd2+ affects IMM permeability and function is the MCU. Blockers of the MCU prevented mitochondrial swelling and Cyt c release (Fig. 6, Table 1). Furthermore, all inhibitors of the MCU, when tested on Cd2+-induced swelling, displayed the same modes of inhibition (competitive for La3+, noncompetitive for RR) as has already been described for Ca2+ (4, 59), suggesting that Ca2+ and Cd2+ are transported into the mitochondria by the same mechanism, that is, the MCU. The inhibitory effects of Cd2+ on the PTP require Cd2+ entry through the MCU and are dependent on the Cd2+ concentration within the mitochondrial matrix and not the extramitochondrial Cd2+ concentration. This was shown by the attenuation of PTP inhibition by Cd2+ with MCU blockers even in the continuous presence of an extramitochondrial Cd2+ concentration of 50 μM (Fig. 6C). A recent report has demonstrated that the mitochondrial MCU behaves as a highly selective Ca2+- and Sr2+-permeable ion channel (29). Unfortunately, its permeability to Cd2+ was not tested. Here, we have used an indirect approach by monitoring the uptake of Cd2+ from the medium into mitoplasts using the membrane-impermeant fluorescence indicator FluoZin-1 (Fig. 7). All the changes of IMM permeability observed appear to be a consequence of Cd2+ influx through the MCU rather than a direct interaction of extramitochondrial Cd2+ with the outer surface of the IMM.
The critical role of the MCU in the regulation of mitochondrial function in health and disease must once more be emphasized in the context of Cd2+ toxicity. Although the targets of Ca2+ and Cd2+ on changes in mitochondrial membrane permeability and toxicity clearly differ [i.e., activation of the PTP by Ca2+ (4, 23) compared with activation of a H2O permeability and inhibition of PTP by Cd2+], the rate-limiting step for these alterations of IMM permeability and subsequent release of Cyt c is entry of these divalent cations through the MCU.
What are the Consequences of Changes in Membrane Permeability on Mitochondrial Function
An important event associated with Cd2+ toxicity of PT cells is mitochondrial swelling-associated release of Cyt c. In MSH buffer, Cyt c release amounted to 20–25% of total mitochondrial Cyt c (Fig. 3). This is consistent with functional estimates of Cyt c present in the intermembrane space (5). Additional Cyt c that is attached to the IMM by electrostatic interaction could be released due to the mere presence of salt (KCl) in the buffer as has already been described by others (8, 20, 43). Once released into the cytosol, Cyt c will typically form an apoptosome with other cytosolic components and trigger activation of a cellular cascade of proteases that ultimately lead to apoptosis (12, 26, 38, 60). It is likely that these processes are operative in Cd2+-induced apoptosis of PT cells (50–52) and are currently under investigation.
In summary, the present study shows that Cd2+ may induce Cyt c release and apoptosis of cultured PT cells after prolonged exposure times. This may occur only when intracellular Cd2+ concentrations are high enough to directly damage mitochondria. Cd2+ concentrations >2 μM induce an increase in IMM permeability and osmotic swelling by entering the matrix space through the MCU and activating H2O channels, possibly AQP8. Swelling is associated with an increase in Cyt c release from the intermembrane space, ultimately leading to the characteristic feature of Cd2+ toxicity of PT cells: apoptosis.
GRANTS
This study was supported by start-up funds from the University of Witten/Herdecke (F. Thévenod), the Deutsche Forschungsgemeinschaft (TH 345/8–1 and 8–2 to F. Thévenod), and grants from the National Kidney Research Fund, UK (F. Thévenod and W.-K. Lee).
ACKNOWLEDGMENTS
We thank Dr. P. Schnfeld (Institute of Biochemistry, Otto-von-Guericke University, Magdeburg, Germany) for the kind gift of alamethicin and advice, Dr. U. Pfüller (Institute of Phytochemistry, University of Witten/Herdecke) for access to the PerkinElmer LS50B luminescence spectrophotometer, Dr. W. S. Kunz (Department of Epileptology, Bonn University Medical Center, Bonn, Germany) for valuable discussions and the use of the Oroboros high-resolution oxygraph, and Dr. U. Hopfer (Department of Physiology and Biophysics, Case Western Reserve University, Cleveland, OH) for providing rat proximal tubule cell lines.
FOOTNOTES
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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School of Biological Sciences, University of Manchester, Manchester, United Kingdom
ABSTRACT
Cd2+ induces apoptosis of kidney proximal tubule (PT) cells. Mitochondria play a pivotal role in toxic compound-induced apoptosis by releasing cytochrome c. Our objective was to investigate the mechanisms underlying Cd2+-induced cytochrome c release from mitochondria in rat PT cells. Using Hoechst 33342 or MTT assay, 10 μM Cd2+ induced 5–10% apoptosis in PT cells at 6 and 24 h, which was associated with cytochrome c and apoptosis-inducing factor release at 24 h only. This correlated with previously described maximal intracellular Cd2+ concentrations at 24 h, suggesting that elevated Cd2+ may directly induce mitochondrial liberation of proapoptotic factors. Indeed, Cd2+ caused swelling of energized isolated kidney cortex mitochondria (EC50 9 μM) and cytochrome c release, which were independent of permeability transition pore (PTP) opening since PTP inhibitors cyclosporin A or bongkrekic acid had no effect. On the contrary, Cd2+ inhibited swelling and cytochrome c release induced by PTP openers (PO43– or H2O2+Ca2+). The mitochondrial Ca2+ uniporter (MCU) played a key role in mitochondrial damage: 1) MCU inhibitors (La3+, ruthenium red, Ru360) prevented swelling and cytochrome c release; and 2) ruthenium red attenuated Cd2+ inhibition of PO43–-induced swelling. Using the Cd2+-sensitive fluorescent indicator FluoZin-1, Cd2+ was also taken up by mitoplasts. The aquaporin inhibitor AgNO3 abolished Cd2+-induced swelling of mitoplasts. This could be partially mediated by activation of the mitoplast-enriched water channel aquaporin-8. Thus cytosolic Cd2+ concentrations exceeding a certain threshold may directly cause mitochondrial damage and apoptotic development by interacting with MCU and water channels in the inner mitochondrial membrane.
cadmium; aquaporin; apoptosis-inducing factor; cyclosporin A
CD2+ REPRESENTS AN INCREASINGLY prevalent occupational and environmental hazard. Every year, >15,000 tons of Cd2+ are used to manufacture nickel-cadmium batteries, paints, chemical stabilizers, plating, or alloys (25). The wide environmental distribution of Cd2+ has led to an increased interest in its biological effects and toxicity (58). Cd2+ has no known physiological function but shares some toxic properties with other metals (18, 58).
A focal point for chronic Cd2+ nephrotoxicity is the proximal tubule (PT), in particular its S1 segment (18), which is associated with a general transport defect of the PT that mimics the de Toni-Debré-Fanconi syndrome. Renal dysfunction in the form of proteinuria, aminoaciduria, glucosuria, and phosphaturia has been demonstrated in persons occupationally exposed to Cd2+ (for a review, see Refs. 30 and 56). In fact, human studies indicate that up to 7% of the general population may develop renal dysfunction from Cd2+ exposure (18).
In the body, Cd2+ circulates mainly as a complex with the 6-kDa protein metallothionein (MT) or with the tripeptide glutathione (GSH), which are easily filtered through the glomerulus. Cd-MT is translocated from the primary urinary filtrate into renal PT cells by receptor-mediated endocytosis (31, 54). After degradation of GSH by -glutamyltranspeptidase bound to the apical plasma membrane of PT cells (10), Cd2+ may also be taken up as a free ion through yet unknown apical transport pathways. In a cell line derived from the S1 segment of rat PT, we have previously demonstrated that Cd2+ (10 μM) is taken up by apical pathways, which operate with slow kinetics and saturate at 24 h (16).
When Cd2+ enters the cytosol of PT cells, it may lose its toxic potency by forming a complex with small molecules (e.g., amino acids) or with peptides and proteins that contain SH groups (such as GSH or MTs) and protect the intracellular milieu from oxidative stress. Depletion of these protective factors, as well as the consequence of Cd2+-induced displacement of endogenous redox active metals (Fe2+, Cu2+) (42), results in the formation of reactive oxygen species (ROS) and subsequent damage to critical organelles. If ROS-mediated stress events are not sufficiently balanced by repair processes, PT cells affected by Cd2+ may be induced to undergo cell death via apoptosis or necrosis in vivo (37, 49) and in vitro (51, 52). Indeed, we have previously shown in an S1 segment of a rat PT cell line that low micromolar (5–10 μM) Cd2+ concentrations induce apoptosis, but not necrosis, of PT cells by a process involving ROS (51, 52).
Although a number of stimuli appear to trigger apoptosis, the current dogma is that apoptosis occurs by activation of two major signaling pathways: the death receptor pathway and the death receptor-independent (mitochondrial) pathway, which may also cross talk under certain conditions (26, 60). Despite the differing upstream apoptosis-mediating pathways, there appears to be some degree of convergence at the mitochondrion (12). After a cell death signal, a number of proapoptotic proteins are liberated from the intermembrane space into the surrounding cytosol, which activate downstream pathways. Factors that are normally sequestered in the mitochondria, but are released upon apoptotic signals, include cytochrome c (Cyt c) (38), apoptosis-inducing factor (AIF) (48), and Smac/Diablo (14, 53), as well as a number of procaspases, which appear to be key elements in the cascade of biochemical events leading to apoptosis.
Accordingly, it has been suggested that Cd2+ induces the release of Cyt c by modifying the interaction of members of the Bcl-2 protein family with mitochondrial membranes (27, 28, 35) and/or by activation of the permeability transition pore (PTP) (13, 36). However, there is accumulating evidence that cytotoxic compounds may alter mitochondrial membrane permeability and the release of Cyt c by processes that are independent of these classic mechanisms (1, 2, 20, 45). Thus the molecular processes underlying Cd2+-induced mitochondrial damage and dysfunction that are associated with Cyt c release still await clarification.
Here, we demonstrate that 10 μM Cd2+ induces apoptosis of rat kidney PT cells at 6 and 24 h whereas Cyt c release is observed after 24 h only, when intracellular Cd2+ concentrations may be high enough to interact directly with mitochondria (16). Cyt c release may be partially mediated by direct effects of Cd2+ on mitochondrial permeability pathways; Cd2+ induces swelling (EC50 9 μM) and release of Cyt c from energized mitochondria and mitoplasts isolated from rat kidney cortex. These processes are independent of opening of the PTP. Rather, Cd2+ enters the inner mitochondrial compartment through the mitochondrial Ca2+ uniporter (MCU) to exert its effects, which may include activation of aquaporin H2O channels, resulting in osmotic swelling and release of Cyt c. We therefore propose that direct effects of cytosolic Cd2+ on inner mitochondrial membrane (IMM) permeability pathways may cause, at least in part, mitochondrial dysfunction and therefore contribute to the development of apoptosis in kidney PT cells.
EXPERIMENTAL PROCEDURES
Materials
DMEM:nutrient mixture F-12 (1:1), FBS, penicillin, and streptomycin were all purchased from GIBCO (Carlsbad, CA). Alamethicin, apo-transferrin, bongkrekic acid (BKA), cyclosporin A (CsA), dexamethasone, epidermal growth factor, ethidium bromide (EtBr), insulin, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT), quinine hydrochloride, rotenone, and ruthenium red (RR) were all purchased from Sigma (St. Louis, MO). CdCl2 was from Merck (Darmstadt, Germany). Digitonin, Hoechst 33342 (H-33342) and Ru360 were from Calbiochem (San Diego, CA). The fluorescent indicator FluoZin-1 was obtained from Molecular Probes (Eugene, OR). The mouse monoclonal antibody against Cyt c (7H8.2C12) was from BD Pharmingen (Erembodegem, Belgium), mouse monoclonal antibody raised against human AIF was from Santa Cruz Biotechnology, (Santa Cruz, CA), the rabbit polyclonal antibody against rat aquaporin-8 (AQP8) was from Alpha Diagnostics (San Antonio, TX), the donkey anti-rabbit IgG and the sheep anti-mouse IgG conjugated with horseradish peroxidase were from Amersham Life Sciences (Bucks, UK), and the donkey anti-mouse indocarbocyanin (Cy3)-coupled IgG was from Dianova (Hamburg, Germany). Inhibitors and drugs were dissolved in either water, ethanol, or dimethyl sulfoxide, and in control experiments, solvents were added to PT cells or isolated mitochondria at a concentration not exceeding 0.2%.
Animals
All experiments conducted were in accordance with the Guiding Principles for Research Involving Animals and Human Beings. Male Sprague-Dawley rats (250–300 g) were obtained from Charles River Laboratories (Sulzfeld, Germany) and were given access to food and water ad libitum.
Methods
Cell culture. An immortalized cell line from the S1 segment of rat PT (WKPT-0293 Cl.2) (57) was cultured in DMEM:nutrient mixture F-12 (1:1) supplemented with 10% FBS, 100 U/ml penicillin G and 100 μg/ml streptomycin sulfate, 1.2 mg/ml NaHCO3, 5 μg/ml insulin, 4 μg/ml dexamethasone, 0.01 μg/ml epidermal growth factor, and 5 μg/ml apo-transferrin in 75-cm2 standard tissue culture flasks (Nalge Nunc International, Rochester, NY) at 37°C in a humidified incubator with 5% CO2. Cells were passaged twice a week on reaching confluency.
Detection of apoptosis and necrosis with H-33342 and EtBr by fluorescence imaging. Staining and experiments were conducted as previously described (52). Briefly, control and Cd2+-treated cells were stained with 2 μg/ml H-33342 followed by 5 μg/ml EtBr. After being washed with 0.2 M HEPES, pH 7.4, cells were visualized under ex/em of 350/460 and 518/605 nm for H-33342 and EtBr, respectively, with the Visichrome High Speed Monochromator system (Visitron Systems, Puchheim, Germany), which was connected to a Zeiss Axiovert 200M microscope (Carl Zeiss, Jena, Germany) equipped with a x20 objective. Images were captured using a digital CoolSPAN ES CCD camera (Roper Scientific, Tucson, AZ) and acquired, processed, and analyzed with MetaMorph software (Universal Imaging, Downingtown, PA). Cells from five random microscopic fields of view at x200 magnification were counted per dish, and the average %apoptotic and necrotic cells were calculated.
MTT assay to detect apoptosis and necrosis. The MTT assay is a measurement of cell viability, and hence an indicator of cell death but does not distinguish between apoptosis and necrosis as well as the inhibition of cell growth (40). MTT is a tetrazolium salt that is catalyzed by active succinate dehydrogenase in the mitochondria of living cells into a blue formazan product. Cells (5 x 103) were seeded into each well of six-well plates and given 48 h before treatment with CdCl2. The original method by Mosmann (40) was modified according to Denizot and Lang (11). This entailed the incubation of 1 mg/ml MTT in serum-free medium without phenol red, aspiration of MTT-containing medium, and dissolving of the formazan product with pure isopropanol without HCl. The product was measured at = 560 and 690 nm using the fixed-wavelength program on a Beckman Coulter DU640 spectrophotometer (Beckman Instruments, Fullerton, CA). Reference wavelength values were subtracted from the test wavelength values. The values were normalized to the control, which was equivalent to 0% cell death.
Detection of cytochrome c or AIF release and apoptosis by immuno- and H-33342 staining using fluorescence imaging. WKPT-0293 Cl.2 cells were grown on glass coverslips and fixed in 4% paraformaldehyde/PBS for 30 min at room temperature. All subsequent steps were also carried out at room temperature. Cells were rinsed three times for 5 min with PBS and permeabilized by incubation in PBS containing 1% SDS for 5 min. After three rinses in PBS, coverslips were inverted onto anti-Cyt c (1:600) or anti-AIF antibodies (1:50) and incubated for 2 h in a moist chamber. After a further washing with PBS, coverslips were incubated on drops of Cy3-conjugated IgG (1:600) for 1 h in the dark. Antibodies were diluted in PBS. After three rinses of 5 min in PBS, nuclei of cells were counterstained with 2 μg/ml H-33342 in 0.2 M HEPES, pH 7.4, for 20 min at 37°C, washed once, and subsequently mounted onto glass slides with Vectashield HardSet Mounting Medium (Vector Laboratories, Peterborough, UK). The cells were visualized under ex/em of 550/630 and 350/460 nm for Cy3 and H-33342, respectively, using the Visichrome High Speed Monochromator system connected to a Zeiss Axiovert 200M microscope equipped with a x100 oil-immersion objective. Images were captured using a digital CoolSPAN ES CCD camera and acquired, processed, and analyzed with MetaMorph software. Using the Color Combine function in MetaMorph, the red component of the original Cy3 16-bit images and the blue component of the original H-33342 16-bit images were merged to produce 24-bit images.
Isolation of mitochondria and mitoplasts. Rat kidney cortex mitochondria were isolated by differential centrifugation essentially as described by Ott et al. (43). Mitoplasts were prepared from mitochondria as described by Greenawalt (21) with slight modifications. A portion of freshly isolated mitochondria (100 mg/ml) was incubated on ice with an equal volume of 12 mg/ml digitonin for 15 min with constant vigorous stirring. The digitonin-containing mitochondrial suspension was diluted 1:3 with isolation buffer before centrifugation at 12,000 g for 10 min at 4°C. The pellet was resuspended in isolation buffer to one-half its volume immediately before the first centrifugation, apportioned into aliquots, and centrifuged again at 12,000 g for 10 min at 4°C. The mitoplast pellets were kept on ice and resuspended in hyperosmotic buffer (1.5 M sucrose, 100 mM HEPES, pH 7.4, with Tris) immediately before use. The purity of the mitoplast preparations was determined by measuring the degree of contamination with Cyt c of the fractions obtained by immunoblotting (data not shown). Protein concentration was determined by the method of Bradford (7).
Measurements of mitochondrial oxygen consumption. Only mitochondria with a respiratory control ratio 4 were used for experiments. Oxygen consumption of isolated mitochondria was measured at 30°C with a PC-supported Oroboros high-resolution oxygraph (Anton Parr, Graz, Austria), which comprised an airtight chamber containing an oxygen electrode and a magnetic stirrer (24). The standard incubation medium was composed of 110 mM mannitol, 60 mM KCl, 60 mM Tris, and 10 mM KH2PO4, pH 7.4, with HCl, which was prewarmed to 30°C, and bubbled with air for at least 15 min before measurements. MgCl2 (5 mM), 10 mM sodium succinate, and 1 μM rotenone were added to 1.5 ml of buffer, followed by 0.35 mg/ml mitochondria, and a steady state of endogenous respiration was allowed to be reached. To induce state 3 respiration, 250 μM K+-ADP was added. Once the ADP was exhausted, respiration reverted back to state 4.
Osmotic swelling assay of mitochondria and mitoplasts. Mitochondria and mitoplasts were used for osmotic swelling experiments within 4 h of isolation. For these, 0.7–1.1 mg/ml mitochondria or 0.3–0.5 mg/ml mitoplasts were incubated in 2 ml of either MSH buffer (210 mM mannitol, 70 mM sucrose, 3 mM HEPES, 10 mM sodium succinate, 1 μM rotenone, pH 7.4, with Tris) or KCl buffer (140 mM KCl, 3 mM HEPES, 10 mM sodium succinate, 1 μM rotenone, pH 7.4, with Tris). Inhibitors, when used, were added before mitochondria/mitoplasts. Changes in volume of energized mitochondria/mitoplasts due to colloid-osmotic effects of solute flux into the mitochondrial matrix after addition of salts and modulators were monitored by the change in absorbance at = 540 nm (3). Kinetic measurements were carried out at 25°C in a Beckman DU-640 spectrophotometer equipped with a Peltier constant-temperature chamber and an automatic six-unit sampler (Beckman Instruments). Data were captured and converted using DU-WinConnection Suite software and analyzed using Microsoft Excel and SigmaPlot 8.0 (SPSS, Chicago, IL). Rates of swelling (absorbance540 nm/min) were calculated from the initial linear portion of mitochondrial absorbance curves. Swelling curves were normalized to maximal mitochondrial swelling. To determine maximal swelling, mitochondria were suspended in KCl buffer, and the maximal absorbance changes induced by the monovalent cation ionophore alamethicin were defined as 100% swelling. Alamethicin-induced absorbance changes amounted to 53 ± 1.4% of initial absorbance. To simplify calculations, maximal swelling was assumed to be 50% of initial absorbance, and normalized values were obtained from the formula {1 – [absorbance/(initial absorbance/2)]} x 100.
Immunoblotting of mitochondrial proteins for Cyt c and AQP8. For immunoblotting, mitochondrial suspensions were collected after swelling measurements and centrifuged at 10,000 g for 5 min at 4°C. Both pellets and supernatants were sonicated on ice for 3 x 5 s at 10 A. Proteins were separated by SDS-PAGE on 15% acrylamide Laemmli minigels and transferred onto polyvinylidene difluoride membranes overnight at 4°C. Blots were blocked with 3% nonfat dry milk and incubated overnight at 4°C with primary anti-Cyt c (1:250) or anti-AQP8 (1:400) antibody. After incubation with horseradish peroxidase-conjugated secondary antibody (1:10,000) for 1 h at 4°C, blots were developed using Western Lighting Plus chemiluminescence reagents (PerkinElmer Life Sciences, Boston, MA), and signals were visualized on X-ray film. Blots were scanned digitally using a Biorad GS 700 Densitometer apparatus and analyzed with Molecular Analyst software (Bio-Rad Laboratories, Hercules, CA).
Cd2+ uptake measurements in mitoplasts with the fluorescence probe FluoZin-1. To measure Cd2+ uptake into mitoplasts, the probe FluoZin-1 (ex = 490 nm, em = 517 nm) was used, which is virtually nonfluorescent in the absence of metals but undergoes an up to 200-fold fluorescence increase upon saturation with micromolar concentrations of metals, such as Zn2+ or Cd2+, with minimal interference by Ca2+ (19). The fluorescence decreases again upon dissociation of Cd2+ from its binding site on the membrane-impermeant indicator. In preliminary experiments, a Kd for Cd2+ of 97 μM was determined (data not shown). Freshly isolated mitoplasts (0.3 mg/ml) were added to 3 ml of buffer containing 110 mM mannitol, 60 mM Tris, 60 mM KCl, pH 7.4, with HCl, as well as 2 μM FluoZin-1 and energized with 10 mM sodium succinate and 1 μM rotenone. Measurements were performed in an LS50B luminescence spectrophotometer (PerkinElmer, Wellesley, MA), and data were collected using the PerkinElmer FL Data Manager program. After stabilization of the fluorescence signal, 20 μM Cd2+ was added every 400 s. As a control, the experiments were repeated in the absence of mitoplasts.
Statistical Analyses
Unless otherwise indicated, all experiments were repeated at least three times with different preparations of mitochondria. Representative data or means ± SE are shown. Statistical analysis using unpaired Student's t-test was carried out with the Sigma Plot 8.0 spreadsheet program. For more than two groups, statistical differences were compared using one-way ANOVA assuming equality of variance with Levene's test and Tukey's post hoc test for pairwise comparison. Statistical analysis was carried out with the SPSS 11.0 program. Results with P 0.05 were considered to be statistically significant. To obtain EC50 values, dose-response curves of Cd2+-induced mitochondrial swelling were fitted using the Sigma Plot 8.0 spreadsheet program assuming a sigmoidal dose response (variable slope).
RESULTS
Cd2+-Induced Apoptosis of PT Cells Is Associated with Cyt c Release at 24 h But Not at 6 h
When cultured PT cells are incubated with the toxic metal Cd2+ at low micromolar concentrations, cell death occurs. To distinguish between apoptosis and necrosis, cell death was determined using the lipophilic DNA dye H-33342, which permeates intact plasma membranes and can therefore label all cells including apoptotic cells (Fig. 1A), whereas the charged DNA dye EtBr can only permeate necrotic cells that have lost their plasma membrane integrity. Control cells at both 6 and 24 h exhibited round, flat, pale blue nuclei (Fig. 1A, left) and negligible apoptosis or necrosis (Fig. 1B). In contrast, 10 μM Cd2+ treatment caused a significant increase of apoptosis but no change in necrosis (Fig. 1B). Apoptosis was defined by fragmentation of the nucleus (asterisk in Fig. 1A) or by condensation (arrow in Fig. 1A). Condensed nuclei have increased fluorescence intensity and are smaller compared with controls. Moreover, they display sharper nucleic outer edges. Quantification of cell death shows that 10 μM Cd2+ induces a significantly higher incidence of apoptosis at 6 or 24 h but not of necrosis (Fig. 1B), which is in accordance with previous data of Cd2+-induced apoptosis in PT cells (16, 52). At 6 h, 10 μM Cd2+ induced 4.2 ± 0.5% apoptosis compared with controls (0.7 ± 0.1%, n = 6). After 24-h Cd2+ treatment, there was 4.9 ± 0.7% apoptosis, which was significantly higher than in the controls (0.8 ± 0.2%, n = 6). The slight increase in apoptosis by Cd2+ after 24 h compared with 6 h was not significant (P = 0.43). Cd2+ (50 μM) further increased the rate of apoptosis at both 6 (7.9 ± 1.2%, n = 6) and 24 h (13.6 ± 2.0%, n = 6). With 50 μM Cd2+, the percentage of necrosis was 1.8 ± 1.3% (n = 6) at 6 h and 8.5 ± 2.5% (n = 6) at 24 h. Necrosis was significantly increased by 50 μM Cd2+ at 24 h compared with controls (P < 0.01). We then used an additional independent technique to measure cell death induced by 10 μM Cd2+, namely, the MTT assay. As shown in Fig. 1C, PT cells incubated with 10 μM Cd2+ for 6 h showed 7.1 ± 1.0% (n = 6) cell death, whereas incubation with 10 μM Cd2+ for 24 h developed 9.1 ± 1.6% (n = 6) cell death, which is of the same magnitude as that obtained with H-33342 and EtBr. The difference between 6 and 24 h was not significant. Moreover, the results also confirm previous published data obtained using annexin V-Alexa 568 staining (49).
Because apoptosis mediated by cytotoxic stimuli including Cd2+ has been associated with mitochondrial release of proapoptotic factors, such as Cyt c or AIF (12, 13, 27, 35), we investigated the release of these factors from mitochondria in PT cells at 6 and 24 h. In Fig. 1D, PT cells were treated with 10 μM Cd2+ for 6 or 24 h and subsequently stained for Cyt c or AIF using specific antibodies and for occurrence of apoptosis with H-33342. In the controls (Fig. 1D, left), the cells have round, pale blue nuclei and punctate Cyt c distribution, as demonstrated by the brighter particulate staining in the cytosolic compartment, indicating that Cyt c is localized to mitochondria. When the cells are treated with Cd2+ for 6 h (Fig. 1D, top right), there is the occurrence of apoptosis as exemplified by the fragmented nucleus, but the cytosolic bright particulate staining of Cyt c remains largely the same as in the control (Fig. 1D, top left). However, after 24-h Cd2+ treatment, apoptotic cells display a major change in Cyt c distribution (Fig. 1D, middle right), compared with the control (Fig. 1D, middle left), which has changed from punctate to diffuse, indicating that Cyt c has been released from mitochondria into the cytosol. Furthermore, the change in Cyt c distribution also seems to occur in cells without condensed or fragmented nuclei (Fig. 1D, middle right), suggesting that Cyt c release from mitochondria may take place before DNA condensation/fragmentation. To confirm the data obtained with Cyt c, we investigated AIF, which is another proapoptotic mitochondrial factor that is released into the cytosol as a result of the loss of integrity of the outer mitochondrial membrane and translocates directly to the nucleus to induce chromatin condensation and high-molecular-weight DNA fragmentation, culminating in apoptosis (48). In control cells, AIF had a well-defined punctate distribution, demonstrated by the brighter particulate staining in the cytosolic compartment (Fig. 1D, bottom left), indicating that AIF was localized to mitochondria. After 6-h exposure to 10 μM Cd2+, the pattern of AIF distribution remained unchanged (data not shown). In contrast, after 24-h incubation with 10 μM Cd2+, apoptotic cells displayed a diffuse AIF distribution, indicating that AIF has been released from mitochondria into the cytosol (Fig. 1D, bottom right).
Cd2+ Induces Swelling and Release of Cyt c in Isolated Mitochondria
Strikingly, recent work from our laboratory has demonstrated that cultured confluent PT cells exposed to 10 μM Cd2+ take up 109Cd2+ through apical transport pathways with slow kinetics, which appeared to have reached equilibrium at 24 h after an approximately fourfold increase in intracellular Cd2+ concentration compared with a 4-h incubation (16). Because the release of Cyt c and AIF observed at 24 h (Fig. 1D) correlated with the increase in intracellular Cd2+ concentration previously described (16), we speculated that when cytosolic Cd2+ exceeds a threshold concentration, it may directly damage mitochondria and induce Cyt c release, which is an integral step of the mitochondrial apoptotic pathway. To test this hypothesis, mitochondria were isolated from rat kidney cortex (rather than from cultured PT cells to obtain a higher yield) and incubated with relevant micromolar Cd2+ concentrations. Figure 2 shows the concentration-dependent swelling of rat kidney cortex mitochondria suspended in MSH buffer induced by Cd2+ with an EC50 of 9 μM (Fig. 2B), which is in accordance with Cd2+-induced apoptosis of PT cells (51, 52). Similar results were also obtained by using KCl buffer (EC50 = 8.4 ± 0.7 μM, n = 8) (data not shown). To investigate whether swelling of kidney cortex mitochondria induced by Cd2+ is associated with Cyt c release, samples were collected for separation of the mitochondrial pellet and supernatant after addition of Cd2+. Figure 3A demonstrates that Cd2+-induced swelling of isolated mitochondria suspended in MSH buffer is accompanied by the release of Cyt c into the supernatant. Mitochondrial preparations were only taken into account for analysis when the controls showed a release of 15% of total Cyt c. Under these conditions, Cd2+ concentrations between 5 and 50 μM significantly increased Cyt c release into the supernatant (Fig. 3B). Data obtained by using KCl buffer showed a consistently high basal Cyt c release in the controls (MSH: 13.1 ± 4.5%, n = 8 vs. KCl: 68.9 ± 3.9%, n = 4). Therefore, no significant difference between controls and Cd2+-exposed mitochondria were obtained (data not shown). The high basal Cyt c release in KCl buffer may be accounted for by the effect of the ionic strength of KCl on a pool of Cyt c that is attached to the IMM by a loose electrostatic interaction (8) and/or centrifugation artifacts in the context of the separation of the mitochondrial pellet and supernatant in KCl buffer.
Cd2+-Induced Mitochondrial Swelling and Cyt c Release Occur Independently of PTP Opening
Figure 4 shows the effect of 1 μM CsA, a potent inhibitor of the PTP (9), on mitochondrial swelling induced by 20 (Fig. 4A) or 50 μM Cd2+ (Table 1) in MSH or KCl buffers (Table 1). As can be seen from the data summarized in Table 1, there was no significant inhibition of the initial rates of swelling when 1 μM CsA was added before the addition of Cd2+. BKA (5 μM), another drug that blocks the PTP by binding to the adenine nucleotide translocator (ANT) (22), did not inhibit Cd2+-induced swelling either (Table 1), suggesting that mitochondrial swelling induced by Cd2+ is not mediated by PTP opening. In line with these observations, we showed that Cyt c release induced by 50 μM Cd2+ was not prevented by 1 μM CsA (Fig. 4B).
View this table:
Cd2+ Prevents Activation of the PTP and Subsequent Cyt c Release
Rather than induce PTP opening, Cd2+ actually inhibited mitochondrial swelling triggered by 5 mM PO43–, an inducer of the PTP (34; for a review, see Refs. 4 and 23) in a concentration-dependent manner (Fig. 5A). To ensure that the inhibition was not caused by an indirect effect of Cd2+ via interaction with the mitochondrial phosphate translocase (23), 2 mM H2O2+50 μM Ca2+, which also induce PTP opening (4, 23), were tested. Again, mitochondrial swelling induced by H2O2 was inhibited as a function of Cd2+ concentration (Fig. 5B). Mitochondrial swelling induced by 5 mM PO43– or 2 mM H2O2+50 μM Ca2+ was strongly inhibited by 1 μM CsA. Moreover, both the Cd2+- and CsA-mediated inhibition of PTP opening were not additive, indicating that both Cd2+ and CsA have the same target of inhibition (data not shown). The inhibitory effect of Cd2+ on PTP opening was also associated with an inhibition of Cyt c release. Mitochondria exposed to 5 mM PO43– released the majority of their Cyt c into the supernatant. In contrast, 50 μM Cd2+ reduced PO43–-induced Cyt c release into the supernatant (Fig. 5C).
Blockers of MCU Prevent Cd2+-Induced Swelling and Cyt c Release
Table 1 shows that the noncompetitive MCU inhibitors, RR and its active component Ru360, as well as the competitive inhibitor La3+ (for a review, see Ref. 4) strongly inhibited Cd2+-induced swelling of kidney cortex mitochondria. Dixon plots of the effects of RR and La3+ on Cd2+-induced mitochondrial swelling in MSH buffer (data not shown) demonstrated that inhibition is noncompetitive and competitive, respectively, as has already been reported for Ca2+ (59). Furthermore, a maximal inhibitory concentration of RR (1–5 μM) prevented the decrease in Cyt c contents in the mitochondrial pellet induced by 50 μM Cd2+ (Fig. 6, A and B). This indicates that Cd2+-induced mitochondrial swelling and Cyt c release are mediated by Cd2+ influx through the RR- and Ru360-sensitive MCU of the IMM. Moreover, Cd2+ inhibition of PTP opening induced by PO43– was released by coincubation with the MCU blocker RR. As shown in Fig. 6C, 1 and 5 μM RR partially restored mitochondrial swelling induced by 5 mM PO43– in the presence of Cd2+, suggesting that the Cd2+ concentration in the inner mitochondrial compartment rather than the outside Cd2+ concentration determines the extent of PTP inhibition.
Evidence for Cd2+Uptake by Mitoplasts from Rat Kidney Cortex
The fluorescent indicator FluoZin-1 is used as a probe to measure the divalent metal Zn2+ with a Kd of 8 μM (19) but is also used as a sensitive indicator of Cd2+ at micromolar concentrations (Molecular Probes). We used membrane-impermeant FluoZin-1 to monitor extramitochondrial changes in Cd2+ concentration. After addition of mitoplasts, the fluorescence intensity decreased slightly (possibly a dilution effect). Addition of 60 nmol Cd2+ induced a small increase in fluorescence signal (solid line in Fig. 7A), whereas a further addition of Cd2+ produced a much larger signal increase. The peak fluorescence signal decreased with time until a plateau was reached after 2–3 min. The decrease in peak fluorescence was observed only in the presence of mitoplasts (compare solid and dotted lines in Fig. 7A) and reflected uptake of Cd2+ from the medium into mitoplasts. With further additions of Cd2+, the rate of change of peak fluorescence diminished further (Fig. 7B), possibly due to saturation of the uptake process.
Cd2+-Induced Mitochondrial Swelling Involves Activation of an Ag+-Sensitive H2O Channel
The magnitude of osmotic swelling induced by Cd2+ cannot be explained by solute and H2O flux through an open PTP, because it is inhibited by Cd2+ (see Figs. 4A and 5 and Table 1). Furthermore, the osmolarity of micromolar Cd2+ entering the matrix space through the MCU is too small to account for the swelling effects. Therefore, we hypothesized that a hydraulic permeability distinct from the PTP may be activated by Cd2+ to mediate H2O flux into the osmotically active matrix space and induce osmotic swelling. The H2O channel protein AQP8 has been detected in hepatocyte mitochondria (17) and is also expressed intracellularly in rat kidney PT (15). Indeed, immunoblotting of membrane fractions of rat kidney cortex with an anti-rat AQP8 antibody revealed a protein band of 28 kDa that was enriched in mitochondria compared with homogenate and was enriched even further in mitoplasts (Fig. 8A). Moreover, the aquaporin inhibitor AgNO3 (5 μM) (41) nearly abolished Cd2+-induced osmotic swelling of mitoplasts (Fig. 8B).
DISCUSSION
What is the Mechanism of Cd2+-Induced Cyt c Release in PT Cells
In the present study, the release of Cyt c or AIF induced by 10 μM Cd2+ in cultured PT cells was observed at 24 h but not at 6 h (Fig. 1D), despite the occurrence of similar rates of apoptosis at both time points (Fig. 1, B and C). This suggested that different processes are responsible for Cd2+-induced apoptosis at 6 and 24 h, and thus we investigated the mechanisms of Cyt c release in more detail. We have previously demonstrated that uptake of Cd2+ by confluent cultured PT cells is slow and saturates at 24 h (16). Therefore, we wondered whether free cytosolic Cd2+ exceeding a threshold concentration may directly affect mitochondria to induce the release of Cyt c and AIF. Here, we provide evidence that Cd2+ induces Cyt c release by acting directly on mitochondrial permeability pathways in the IMM. The data obtained in Figs. 2 and 3 suggest that concentrations of Cd2+ >2 μM may contribute to Cyt c release and thus to apoptosis of cells exposed to 10 μM Cd2+, which may become relevant when intra- and extracellular Cd2+ concentrations are near equilibrium (16). However, other mechanisms may also contribute to apoptosis induced by Cd2+. ROS are generated during Cd2+ exposure of PT cells for 4–8 h (51, 52) and may therefore activate PTP opening (13, 36). Interestingly, concentrations of 5–10 μM Cd2+ inhibit PTP opening induced by H2O2 (Fig. 5), which could explain why no major Cyt c release is observed at 6 h in cultured PT cells (Fig. 1C), even when ROS are elevated (51, 52). In addition, it has been shown that Cd2+ induces the release of Cyt c by modifying the interaction of members of the Bcl-2 protein family with mitochondrial membranes (27, 28, 35). It is therefore likely that several processes contribute to Cd2+-induced Cyt c release in PT cells, depending on time and Cd2+ concentrations. Moreover, recent data from our laboratory demonstrate that exposure of PT cells to 10 μM Cd2+ induces activation of caspases 3 and 9 at 24 h, but not at 6 h, which nicely corresponds with the observation of Cyt c release at 24 h. Caspase-independent apoptosis at early time points (3–8 h) turns out to be mediated by activation of the Ca2+-activated proteases calpains (W.-K. Lee and F. Thévenod, unpublished observations).
How Does Cd2+ Affect Permeability of the IMM
In general terms, temporary or permanent alterations of the integrity of the IMM and/or outer mitochondrial membrane induce the release of proapoptotic proteins (6, 33, 39). Various factors have been implicated in alterations of mitochondrial membrane permeability, such as members of the Bcl-2 protein family (46, 47, 55) and/or ROS-triggered opening of the PTP, which may consist of both IMM and outer mitochondrial membrane proteins, but whose molecular identity is still elusive (32).
Several major conclusions can be drawn from this study. First, Cd2+ does not induce opening of the PTP. Typical inhibitors of PTP, such as CsA and BKA, had no significant inhibitory effect on Cd2+-induced mitochondrial swelling and Cyt c release (Fig. 4, Table 1). Moreover, rather than open the PTP, Cd2+ concentrations 5 μM inhibited opening of the PTP induced by Ca2++H2O2 or PO43– (Fig. 5), which are characteristically operative during hypoxia or oxidative stress. If the PTP is not involved in Cd2+-induced osmotic swelling of mitochondria suspended in MSH buffer, the only likely alternative mechanism that can account for Cd2+-induced swelling must be the activation of an hydraulic conductance because the osmolality of Cd2+ entering the mitochondrial matrix through the MCU is too small to explain the observed magnitude of mitochondrial swelling (15–20% absorbance change). As a matter of fact, we found AQP8 to be expressed in the IMM (Fig. 8A), which confirms recent data in hepatocytes (17). The transmitochondrial membrane driving force for swelling is induced by the osmotic gradient provided by the mitochondrial matrix that consists of anionic proteins, monovalent cations, intermediates of the Krebs cycle, and other small organic molecules (3). With the use of AgNO3, a more potent inhibitor of aquaporins than Hg2+ (41), Cd2+-induced swelling of mitoplasts was abolished (Fig. 8B). Possibly, other hydraulic permeability pathways in the IMM may also be affected by Cd2+. Thus we show that Cd2+ can alter IMM permeability pathways and induce the release of Cyt c by processes that are independent of the PTP, which confirms that PTP opening is not the only mechanism underlying mitochondrial swelling (1, 2, 20, 45). At the present stage, we can only speculate about the mechanism of Cd2+-induced activation of AQP8, but cysteine residues are important for AQP channel function and inhibition by Hg2+ (44) and could be involved in the alteration of the AQP8-associated permeability of the membrane by Cd2+.
Which Pathway Mediates Cd2+ Entry into the Inner Mitochondrial Compartment
The key element to an understanding of how Cd2+ affects IMM permeability and function is the MCU. Blockers of the MCU prevented mitochondrial swelling and Cyt c release (Fig. 6, Table 1). Furthermore, all inhibitors of the MCU, when tested on Cd2+-induced swelling, displayed the same modes of inhibition (competitive for La3+, noncompetitive for RR) as has already been described for Ca2+ (4, 59), suggesting that Ca2+ and Cd2+ are transported into the mitochondria by the same mechanism, that is, the MCU. The inhibitory effects of Cd2+ on the PTP require Cd2+ entry through the MCU and are dependent on the Cd2+ concentration within the mitochondrial matrix and not the extramitochondrial Cd2+ concentration. This was shown by the attenuation of PTP inhibition by Cd2+ with MCU blockers even in the continuous presence of an extramitochondrial Cd2+ concentration of 50 μM (Fig. 6C). A recent report has demonstrated that the mitochondrial MCU behaves as a highly selective Ca2+- and Sr2+-permeable ion channel (29). Unfortunately, its permeability to Cd2+ was not tested. Here, we have used an indirect approach by monitoring the uptake of Cd2+ from the medium into mitoplasts using the membrane-impermeant fluorescence indicator FluoZin-1 (Fig. 7). All the changes of IMM permeability observed appear to be a consequence of Cd2+ influx through the MCU rather than a direct interaction of extramitochondrial Cd2+ with the outer surface of the IMM.
The critical role of the MCU in the regulation of mitochondrial function in health and disease must once more be emphasized in the context of Cd2+ toxicity. Although the targets of Ca2+ and Cd2+ on changes in mitochondrial membrane permeability and toxicity clearly differ [i.e., activation of the PTP by Ca2+ (4, 23) compared with activation of a H2O permeability and inhibition of PTP by Cd2+], the rate-limiting step for these alterations of IMM permeability and subsequent release of Cyt c is entry of these divalent cations through the MCU.
What are the Consequences of Changes in Membrane Permeability on Mitochondrial Function
An important event associated with Cd2+ toxicity of PT cells is mitochondrial swelling-associated release of Cyt c. In MSH buffer, Cyt c release amounted to 20–25% of total mitochondrial Cyt c (Fig. 3). This is consistent with functional estimates of Cyt c present in the intermembrane space (5). Additional Cyt c that is attached to the IMM by electrostatic interaction could be released due to the mere presence of salt (KCl) in the buffer as has already been described by others (8, 20, 43). Once released into the cytosol, Cyt c will typically form an apoptosome with other cytosolic components and trigger activation of a cellular cascade of proteases that ultimately lead to apoptosis (12, 26, 38, 60). It is likely that these processes are operative in Cd2+-induced apoptosis of PT cells (50–52) and are currently under investigation.
In summary, the present study shows that Cd2+ may induce Cyt c release and apoptosis of cultured PT cells after prolonged exposure times. This may occur only when intracellular Cd2+ concentrations are high enough to directly damage mitochondria. Cd2+ concentrations >2 μM induce an increase in IMM permeability and osmotic swelling by entering the matrix space through the MCU and activating H2O channels, possibly AQP8. Swelling is associated with an increase in Cyt c release from the intermembrane space, ultimately leading to the characteristic feature of Cd2+ toxicity of PT cells: apoptosis.
GRANTS
This study was supported by start-up funds from the University of Witten/Herdecke (F. Thévenod), the Deutsche Forschungsgemeinschaft (TH 345/8–1 and 8–2 to F. Thévenod), and grants from the National Kidney Research Fund, UK (F. Thévenod and W.-K. Lee).
ACKNOWLEDGMENTS
We thank Dr. P. Schnfeld (Institute of Biochemistry, Otto-von-Guericke University, Magdeburg, Germany) for the kind gift of alamethicin and advice, Dr. U. Pfüller (Institute of Phytochemistry, University of Witten/Herdecke) for access to the PerkinElmer LS50B luminescence spectrophotometer, Dr. W. S. Kunz (Department of Epileptology, Bonn University Medical Center, Bonn, Germany) for valuable discussions and the use of the Oroboros high-resolution oxygraph, and Dr. U. Hopfer (Department of Physiology and Biophysics, Case Western Reserve University, Cleveland, OH) for providing rat proximal tubule cell lines.
FOOTNOTES
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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