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Inhibition of Ca²⁺ Influx Is Required for Mitochondrial Reactive Oxygen Species-Induced Endoplasmic Reticulum Ca²⁺ Depletion and Cell Death in Leukemia Cells

Arthritis and Immune Disorder Research Centre, University Health Network and Department of Immunology, University of Toronto; Toronto, Ontario, Canada

Received March 9, 2006; accepted July 18, 2006

	Abstract

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Disturbances of endoplasmic reticulum (ER) Ca²⁺ homeostasis or protein processing can lead to ER stress-induced cell death. Increasing evidence suggests that oxidative stress (OS) plays an important role in a variety of cell death mechanisms. To investigate the role of OS in ER stress, we measured OS in response to three ER stress agents: econazole (Ec), which stimulates ER Ca²⁺ release and blocks Ca²⁺ influx; thapsigargin (Tg), a sarco(endo)plasmic reticulum Ca²⁺ ATPase inhibitor that releases ER Ca²⁺ and stimulates Ca²⁺ influx; and tunicamycin (Tu), a glycosylation inhibitor that causes protein accumulation in the ER. Ec, but not Tg or Tu, caused a rapid increase in OS. Reactive oxygen species (ROS) generation was observed within mitochondria immediately after exposure to Ec. Furthermore, Ec hyperpolarized the mitochondrial membrane and inhibited adenine nucleotide transport in cell-free mitochondria, suggesting a mitochondrial target. Antimycin A, an inhibitor of complex III in electron transport, reversed mitochondrial hyperpolarization, OS generation, ER Ca²⁺ depletion, and cell death by Ec, suggesting complex III dependence for these effects. Antioxidants butylated hydroxytoluene and N-Acetyl-L-cysteine prevented ER Ca²⁺ depletion and cell death by Ec. However, inhibition of Ca²⁺ influx by Ec was unaffected by either antimycin A or the antioxidants, suggesting that this target is distinct from the mitochondrial target of Ec. Atractyloside, an adenine nucleotide transport inhibitor, generated ROS and stimulated ER Ca²⁺ release, but it did not block Ca²⁺ influx, deplete the ER or induce cell death. Taken together, these results demonstrate that combined mitochondrial ROS generation and Ca²⁺ influx blockade by Ec is required for cell death.

Disruption of Ca²⁺ homeostasis in the ER, such as after treatment with the SERCA inhibitor thapsigargin (Tg) (Thastrup et al., 1990), can stimulate sustained Ca²⁺ influx from the extracellular milieu. In some cell types, this leads to both cytosolic and mitochondrial Ca²⁺ overload (Babich et al., 1994; Soboloff and Berger, 2002), triggering apoptosis (Orrenius et al., 2003). Econazole (Ec), an imidazole antifungal, also stimulates depletion of the Tg-sensitive ER calcium store. However unlike Tg, Ec additionally blocks Ca²⁺ influx, resulting in sustained ER Ca²⁺ depletion (Franzius et al., 1994; Jan et al., 1999; Soboloff and Berger, 2002). In previous studies, we have shown that the Ca²⁺ depletion caused by Ec induces activation enhanced cell death in leukemic cells, breast cancer cells, and murine bone marrow-derived mast cells through sustained inhibition of protein synthesis (Gommerman and Berger, 1998; Soboloff and Berger, 2002; Soboloff et al., 2002; Zhang et al., 2002; Zhang and Berger, 2004). However, the mechanism of how this compound affects the intracellular ER store and the subsequent fate of the cell remains unknown.

Increasing evidence suggests that reactive oxygen species (ROS) and the oxidation-reduction (redox) state play important roles in a variety of cell death mechanisms induced by widely used antitumor drugs or by environmental toxic substances (Orrenius, 1985; Orrenius and Nicotera, 1987; Slater et al., 1995) (Feinendegen, 2002; Ueda et al., 2002). Because oxidative damage to the ER has been implicated in some forms of cell death (Hayashi et al., 2003; Lai et al., 2003; Watanabe et al., 2003), we investigated the possible role of oxidative stress (OS) in the induction of ER stress by different agents. Here, we show that ER Ca²⁺ depletion and cell death induced by Ec but not Tg or tunicamycin (Tu) is dependent on ROS production at the mitochondria, thus identifying a mediating role for ROS in communicating mitochondrial disruption to the ER.

	Materials and Methods

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Cell Culture, Cell Death, and Caspase Assays. Human promyelocytic leukemia HL-60 cells were grown in RPMI 1640 medium containing 10% heat-inactivated fetal bovine serum (FBS) and 2.5 mM L-glutamine. HL-60 cells were treated with 15 µM Ec in the presence or absence of 125 nM AA, 5 µM BHT, or 1 mM Nac at 37°C for 2 h, in medium containing 1% FBS. Drugs were washed out, and the cells were recovered in 10% FBS drug-free RPMI 1640 medium for 6 h. Samples were costained with Annexin V-Cy5 (BioVision, Mountain View, CA) and propidium iodide (PI), and cell death was determined by flow cytometry. Early (Annexin V-positive, PI-negative) and late (Annexin V-positive, PI-positive) events were scored as dead cells. Caspase-3/7 activation was measured using the Vybrant 5-carboxyfluorescein caspase assay kit from Invitrogen (Carlsbad, CA) as described by the manufacturer.

Detection of Oxidative Stress and Mitochondrial ROS. To examine the generation of OS, HL-60 cells were incubated with the indicated concentrations of Ec, Tu, or Tg in RPMI 1640 containing 2% FBS at 37°C for 2 h and then loaded with the OS indicator 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate (8 µM CM-H₂DCFDA; Invitrogen, Carlsbad, CA) at 37°C for 30 min. Fluorescence was measured by flow cytometry using excitation at 488 nm and emission at 530 nm at indicated intervals after treatment. Mitochondrial-specific ROS generation was measured using the MitoSox Red fluorescent dye (Invitrogen) as described by the manufacturer. This dye accumulates in the mitochondria, is oxidized by superoxide, and emits at 580 nm.

Measurement of Cytosolic Ca²⁺ Concentration. Cytosolic Ca²⁺ measurements were performed by flow cytometry. HL-60 cells were serum-deprived for 2 h in Tyrode's buffer (10 mM HEPES, 100 mM NaCl, 5 mM KCl, 1.4 mM CaCl₂, 1 mM MgCl₂, 5.6 mM glucose, and 0.05% bovine serum albumin (0.05%) containing Ec, atractyloside (Atra), antioxidants, and/or hydrogen peroxide where indicated. Cells were then incubated in loading buffer (5 µM indo-1AM and 0.03% Pluronic F-127 in Tyrode's buffer; both from Invitrogen) at 37°C for 30 min and washed and incubated (15 min at room temperature) to allow for the complete conversion of indo-1AM to Ca²⁺-sensitive indo-1 through hydrolysis. Measurements were performed using a laser tuned to 338 nm while monitoring emissions at 405 and 450 nm. FloJo software (Treestar Software, Ashland, OR) was used to analyze the flow cytometric data. The concentration of intracellular free Ca²⁺ was calculated according to the following formula (Grynkiewcz et al., 1985): [Ca²⁺]_i = K_d x (F_min/F_max) x (R - R_min)/(R_max - R), where R is the ratio of the fluorescence intensities measured at 405 and 450 nm during the experiments, and F is the fluorescence intensity measured at 450 nm. R_min, R_max, F_min, and F_max were determined from in situ calibration of viable cells using 4 µM ionomycin in the absence (R_min and F_min; 10 mM EGTA) and presence of (R_max and F_max) of Ca²⁺. K_d (250 nM) is the dissociation constant for indo-1 at 37°C. R_min, R_max, F_min, and F_max varied depending upon settings and were determined at the beginning of each experimental procedure and experimental condition.

Potentiometric Measurement of Mitochondria Membrane Potential. MMP measurements were performed by flow cytometry. Cells (5 x 10⁵ cells/ml) were growth factor-deprived for 2hin Tyrode's buffer and then incubated with 5 µg/ml JC-1 (Invitrogen) for 15 min at room temperature in Tyrode's buffer. Then, cells were washed (three times) to remove extracellular JC-1. Measurements were performed using a laser tuned to 488 nm while monitoring the emissions of JC-1 monomers at 530 nm and JC-1 aggregates at 585 nm. MMP measurements were normalized using a modification of a formula from Rottenberg and Wu (1998): MMP = (R - R_Fccp)/(R₀ - R_Fccp) x 100, where R is the ratio of the fluorescence intensities measured at 530 and 585 nm during the experiments, R₀ is the fluorescence ratio of untreated cells, and R_Fccp is the fluorescence intensity measured after the addition of 2 µM carbonyl cyanide p-(trifluomethoxy) phenylhydrazone (FCCP; Sigma-Aldrich, St. Louis, MO), a procedure that collapses the mitochondrial membrane potential and was performed at the beginning of each experiment.

Adenosine Nucleotide Transfer-Dependent ADP Import Assay. Adenosine nucleotide transfer (ANT) across the mitochondrial membrane in ER-stressed cells was measured as described by Vander Heiden et al. (1999). In brief, 5 x 10⁷ HL-60 cells were disrupted by mechanical lysis through homogenization in a mitochondria isolation buffer (200 mM mannitol, 70 mM sucrose, 10 mM HEPES, pH 7.4, and 1 mM EGTA). After centrifugation of the supernatant (750g for 10 min to remove debris followed by 10,000g for 10 min), mitochondrial pellets were resuspended in ADP import buffer (250 mM sucrose, 20 mM HEPES, pH 7.2, 10 mM KCl, 5 mM succinate, 3 mM KH₂PO₄, 1.5 mM MgCl₂, 1 mM EGTA, and 5 µM rotenone) with or without the adenine nucleotide transporter inhibitor atractyloside (50 µM; Sigma-Aldrich). [¹⁴C]ADP (1 µCi; PerkinElmer Life and Analytical Sciences, Boston, MA) was added to the mitochondrial suspension and incubated for 10 min on ice. After washing two times in ADP import buffer, the samples were resuspended in scintillant (PerkinElmer) and quantified using a beta counter (PerkinElmer Wallac, Gaithersburg, MD). ANT-dependent ADP transport activity was determined by calculating the difference in counts between samples that were or were not preincubated with atractyloside.

ATP Bioluminescence Assay. ATP levels were measured by using ATP bioluminescence assay kit CLS II (Roche Diagnostics, Indianapolis, IN). In brief, cells (5 x 10⁵ cells/ml) were treated with 15 µM Ec, 125 nM antimycin A, or both at 37°C for 2 or6hin RPMI 1640 medium with 0.05% bovine serum albumin. After washing in chilled PBS, cells were resuspended in 100 µl of dilution buffer, added to 900 µl of boiling buffer (100 mM Tris and 4 mM EDTA, pH 7.5), boiled for 2 min, and centrifuged at 1000g for 1 min. The supernatants were transferred to a fresh tube and kept on ice until measurements. ATP concentration was measured with a luminometer (Luminoskan; Thermo Electron Corporation, Waltham, MA) by adding 100 µl of luciferase reagent to 100 µl of sample or ATP standards (10^-9-10^-4 M). ATP values were calculated from a log-log plot of the standard curve.

View larger version (28K): [in this window] [in a new window]   Fig. 1. Generation of OS in ER stress agent-treated cells. HL-60 cells were incubated with ER stress agents Ec (15 µM) (A), Tg (1 µM) (B), or Tu (1 µg/ml) (C) for 2 h and then loaded with 8 µMH2DCFDA for 30 min at 37°C and rinsed with PBS. Fluorescence was monitored by flow cytometry at FL-1. The gray tracing is the untreated control. The data depicted are a representative experiment. D, comparison of DCF fluorescence in arbitrary units for Ec-, Tg-, or Tutreated cells compared with control. Reported values are averages of three independent determinations and are reported ± S.E.M. **, p < 0.01 compared with control. E, time course of DCF fluorescence in response to 15 µM Ec.

	Results

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Oxidative Stress Generation by ER Stress Agents. To investigate the possibility that ER stress inducers generate ROS, we exposed HL-60 cells to the agents Ec, Tg, and Tu for 2 h and measured OS levels in the cells by flow cytometry using the generalized OS-sensitive fluorescent dye CM-H₂DCFDA (Halliwell and Whiteman, 2004). As shown in Fig. 1, A to D, increased fluorescence levels, indicating increases in OS levels, was observed for cells treated with Ec but not Tg or Tu. We have previously demonstrated that all three agents induce ER stress within the chosen 2-h time period, as indicated by induction of eukaryotic translation initiation factor 2 subunit phosphorylation and BiP expression, along with suppression of protein synthesis (Soboloff and Berger, 2002; Zhang and Berger, 2004). However, this is achieved differently for each agent. Thus, Tu is an inhibitor of glycosylation (Kuo and Lampen, 1974) and causes ER stress through the accumulation of protein in the ER (Dorner et al., 1990). Tg inhibits the SERCA pump responsible for refilling ER Ca²⁺ stores (Thastrup et al., 1990), leading to rapid depletion of ER Ca²⁺ stores, followed by influx of Ca²⁺ from the extracellular milieu. Although Ec is best known as an inhibitor of Ca²⁺ influx, it also promotes ER Ca²⁺ depletion (Jan et al., 1999), although the mechanism of this Ca²⁺ release has not previously been addressed.

View larger version (31K): [in this window] [in a new window]   Fig. 2. Ec hyperpolarizes MMP, generates mitochondrial ROS, and inhibits ADP transport. A, JC-1 loaded HL-60 cells were exposed to 15 µM Ec, 1 µM Tg, 1 µg/ml Tu, or 2 µM FCCP, respectively, and the membrane potential was monitored by flow cytometry as described under Materials and Methods. B, HL-60 cells were exposed to 15 µM Ec, 2 µM PMA, or 50 µM atractyloside for 2 h and then loaded with 8 µMH2DCFDA for 30 min at 37°C and rinsed with PBS. The mean fluorescence intensity was measured by flow cytometry. C, HL-60 cells were treated with ROS-generating agents described as in B but loaded with 5 µM MitoSox for 30 min at 37°C and rinsed with PBS. The mean fluorescence intensity was measured by flow cytometry. D, time course of MitoSox-sensitive ROS in response to 15 µM Ec. E, cell-free mitochondria were prepared as described under Materials and Methods. Samples were incubated with ER stress agents for 10 min in the presence or absence of atractyloside followed by the addition of [14C]ADP. ANT-dependent ADP transport activity was determined by calculating the atractyloside inhabitable counts. ANT-dependent ADP transport was inhibited by 15 µM Ec but not by 1 µMTgor1 µg/ml Tu. Values are reported as mean ± S.E.M.

Hyperpolarization of Mitochondrial Membrane Potential by Ec. The most common mechanism of OS generation by toxic agents is the disruption of electron transport in mitochondria (Ueda et al., 2002; Eckert et al., 2003; Inoue et al., 2003; Kadenbach, 2003; Rego and Oliveira, 2003). To investigate the effect of Ec on mitochondrial function, we first measured mitochondrial membrane potential, a useful indicator of mitochondrial activity and status, after exposure to Ec. As shown in Fig. 2A, the addition of Ec to cells caused an immediate hyperpolarization of the mitochondrial membrane. In contrast, Tg and Tu had no effect, whereas the protonophore Fccp caused an immediate collapse of the potential. These observations suggested that Ec, but not Tg or Tu, had a mitochondrial target.

To further investigate the possibility of a mitochondrial target for Ec-induced ROS generation, we used MitoSox Red dye (Invitrogen), a mitochondrial-specific superoxide indicator. To confirm that MitoSox measures ROS generated at the mitochondria, we used the phorbol ester PMA, which stimulates oxidase activity at the plasma membrane (Teufelhofer et al., 2003; Dooley et al., 2004) and atractyloside, which generates mitochondrial ROS via inhibition of adenine nucleotide activity (see below) as controls. As shown in Fig. 2B, the CM-H₂DCFDA dye detects similar levels of fluorescence by Ec, PMA, and atractyloside over a 2-h time period. In contrast, Ec and atractyloside, but not PMA, generate the superoxide anion, as detected with MitoSox (Fig. 2C). It is noteworthy that a nearly 3.5-fold increase in superoxide accumulation was observed in the presence of Ec, a far greater change than observed when DCF fluorescence was assessed. Cells were then stained with MitoSox, and the effect of Ec addition was followed over time. As shown in Fig. 2D, Ec stimulated a rapid and sustained increase in MitoSox fluorescence in contrast to the somewhat slower change in total DCF fluorescence (Fig. 1E). These observations are consistent with DCF, indicating ROS generation and mitochondria as the source of Ec-induced ROS.

View larger version (32K): [in this window] [in a new window]   Fig. 3. Antimycin A reverses mitochondrial membrane hyperpolarization by Ec. A, illustration of the sites of action of inhibitors of mitochondrial electron transport. B, JC-1-loaded cells were pretreated for 20 min with 10 µM rotenone, 10 µM 4,4,4-trifluoro-1-(2-thienyl)-1,3-butanedione, or 400 µM KCN. The arrow indicates where 15 µMEcor2 µM FCCP was added to the cells. C, effect of antimycin A on membrane hyperpolarization. Cells stimulated with antimycin A (125 nM), Ec, or FCCP. Mitochondrial membrane potential was recorded for 2 h.

Antimycin A Reverses Mitochondrial Hyperpolarization by Ec. Because adenine nucleotide transport is coupled to electron transport, it is possible that a member of the electron transport chain is the direct target of Ec, leading to ROS production and suppression of ANT function. To further investigate this possibility, we determined the effect of various inhibitors of electron transport on the ability of Ec to hyperpolarize the mitochondrial membrane. The inhibitors used are illustrated in Fig. 3A. As shown in Fig. 3B, preexposure of cells to 4,4,4-trifluoro-1-(2-thienyl)-1,3-butanedione, rotenone, or KCN, inhibitors of electron transport complexes I, II, and IV, respectively, had no effect on the ability of Ec to hyperpolarize the mitochondrial membrane. In contrast, the complex III inhibitor antimycin A both partially reduced mitochondrial membrane potential on its own and completely neutralized mitochondrial membrane hyperpolarization by Ec (Fig. 3C). Furthermore, the ability of antimycin A to neutralize Ec-induced hyperpolarization persisted for 2 h. These results therefore suggest that Ec is targeting the electron transport chain in a complex III-dependent manner, resulting in ROS generation and inhibition of ANT function.

Antimycin A Reverses ROS Generation by Ec. The fact that antimycin A neutralizes Ec-induced hyperpolarization seems somewhat paradoxical given that antimycin A on its own has been reported to generate reactive oxygen. To further investigate the interaction between Ec and antimycin A, we measured ROS production after treatment with Ec, antimycin A, or both compounds together. Cells were incubated with the two compounds for 2 h, and ROS content was measured as described above. As shown in Fig. 4A, antimycin A alone, at the concentration used in this study, did not have any effects on ROS levels. However, it clearly suppressed ROS generation by Ec. This result therefore correlates Ec-induced hyperpolarization with ROS production and provides further evidence that the ability of Ec to generate ROS is dependent on complex III activity.

View larger version (16K): [in this window] [in a new window]   Fig. 4. Antimycin A inhibits and reverses OS generation induced by Ec and partially restores ATP production. A, cells were incubated with 15 µM Ec, 125 nM antimycin A, or both at 37°C for 2 h, and DCF fluorescence was measured as described under Materials and Methods. Data are shown as mean ± S.E.M. *, p < 0.05; **, p < 0.01. B and C, intracellular ATP was measured in cells (0.5-1.0 x 106 cells/ml) treated with 15 µM Ec, 125 mM antimycin A, or both, at 37°C for 2 h (B) or 6 h (C). Values shown are reported as mean ± S.E.M. *, p < 0.05; **, p < 0.01.

Antioxidants Do Not Reverse Mitochondrial Membrane Hyperpolarization by Ec. The results described above show that Ec hyperpolarizes the mitochondrial membrane and generates ROS. Moreover, both of these endpoints could be reversed by the complex III inhibitor antimycin A. These results, however, do not address the causal nature of ROS production. In particular, are increased ROS levels required for Ec-induced changes in membrane potential? To address this question, we pretreated cells with the antioxidants BHT or Nac and then followed mitochondrial membrane potential after exposure to Ec. Control experiments demonstrated that both antioxidants suppressed ROS generation by Ec (data not shown). As shown in Fig. 5, Nac slightly lowered the resting mitochondrial membrane potential compared with untreated cells. However, neither Nac nor BHT prevented Ec from hyperpolarizing the membrane above the resting level. Furthermore, the hyperpolarizing effect of Ec persisted for more than 2 h. Therefore, it is likely that the generation of ROS occurs either in parallel with or down-stream of changes in mitochondrial membrane potential.

View larger version (14K): [in this window] [in a new window]   Fig. 5. Antioxidants do not reverse MMP hyperpolarization by Ec. Cells were pretreated with or without 5 µM BHT or 1 mM Nac at 37°C for 20 min followed by JC-1 loading and flow cytometry procedures described under Materials and Methods. Mitochondrial potential was followed for 2 h after 15 µMEcor2 µM FCCP was added.

ER Ca²⁺ Depletion by Ec Is Inhibited by Antimycin A and Antioxidants. We have previously documented that Ec, in addition to blocking Ca²⁺ influx, also causes Ca²⁺ depletion from the ER (Soboloff and Berger, 2002; Zhang and Berger, 2004). To determine the role of the mitochondrial activity of Ec in ER depletion, we treated cells with Ec or Ec plus antimycin A and then measured the amount of Ca²⁺ present in the ER by releasing it into the cytoplasm with Tg while blocking the influx of Ca²⁺ with the nonspecific Ca²⁺ channel blocker Ni²⁺. As shown in Fig. 6A, no Ca²⁺ was released into the cytoplasm by Tg after pretreatment of cells with Ec, demonstrating that Ec depletes the ER. Antimycin A alone had little effect on Tg-releasable Ca²⁺. However, Ec-induced depletion of the Tg-sensitive store was reversed by antimycin A, revealing a key role for the mitochondrial target of Ec in ER Ca²⁺ depletion. To assess the role of mitochondria on Ca²⁺ influx, experiments were then performed in the absence of Ni²⁺. The difference between the magnitude of Tg-induced Ca²⁺ influx in the presence of Ni²⁺ versus the absence of Ni²⁺ reflects Ca²⁺ influx. Because Ec blocks Ca²⁺ influx in addition to depleting the ER of Ca²⁺, no Tg-induced changes in Ca²⁺ concentration occur after Ec pretreatment, even when Ni²⁺ is not present (Fig. 6B). Antimycin A alone had no effect on Tg-induced Ca²⁺ release or influx. However, the Tg-induced Ca²⁺ concentration observed with both anti-mycin A and Ec present was comparable with that observed in experiments when Ni²⁺ was present (Fig. 6A). Hence, antimycin A neutralized Ec-induced ER Ca²⁺ depletion, but it had little or no effect on Ec-induced inhibition of Ca²⁺ influx.

View larger version (33K): [in this window] [in a new window]   Fig. 6. Ec-generated OS targets ER Ca2+ stores. Cells were loaded with indo-1AM in Tyrode's buffer as described under Materials and Methods. Ca2+ concentration was monitored by flow cytometry. A, cells were preincubated in the absence (Ctl) or presence of 15 µM Ec or Ec + 125 nM AA for 2h.Ni2+ (5 mM) was added to the cells to nonspecifically prevent influx. Ca2+ released by 3 µM Tg into the cytoplasm was used as an indicator of ER Ca2+ content. B, as in A, however, Ni2+ was not added to cells before Tg. C, as in A, except 5 µM BHT was used. D, as in C, however, Ni2+ was not added before Tg. E, as in A, except 1 mM Nac was used. F, as in E, however, Ni2+ was not added before Tg.

Antimycin A and Antioxidants Protect Cells from Ec-Induced Cell Death. Because antimycin A and the antioxidants BHT and Nac prevent ER Ca²⁺ depletion by Ec, we tested its effect on Ec-induced toxicity. Cells were exposed to Ec plus antimycin A or the antioxidants for 2 h. The agents were then removed, and the cells were incubated in medium for an additional 6 h. Antimycin A, BHT, and Nac were nontoxic at the concentrations used under these conditions (data not shown). The cells were then evaluated by flow cytometry using Annexin V staining. As shown in Fig. 7A, antimycin A, BHT, and Nac provided significant protection from Ec-induced cell death, revealing a key role for mitochondrially generated ROS and the release of ER Ca²⁺ in Ec toxicity.

View larger version (23K): [in this window] [in a new window]   Fig. 7. Ec-induced cell death and caspase activation are suppressed by antimycin A, BHT, and Nac. Cells were treated with 15 µMEcinthe presence or absence of 125 nM AA, 5 µM BHT, or 1 mM Nac at 37°C for 2 h, in medium containing 1% FBS. Drugs were washed out, and the cells were recovered in RPMI 1640 medium plus 10% FBS for 6 h (A). Samples were costained with Annexin V-Cy5 and propidium iodide, and cell death was determined by flow cytometry. Values include both early- and late-stage apoptotic cells. **, p < 0.01; ***, p < 0.001 compared with Ec. B, cells were treated as described above, except the recovery time was extended to18 h. Cells were then incubated with the fluorochrome inhibitor of caspases reagent (Invitrogen) to detect active caspases-3 or-7 and analyzed by flow cytometry. Reported values are percentage of cells that are positive for activated caspase-3/7 and are averages of two repeat measurements with standard errors.

Hydrogen Peroxide and Atractyloside Mobilize Ca²⁺ from the ER without Depleting the Store. Other studies have shown that ROS stimulate Ca²⁺ release from the ER (Downey, 1990). Consistent with these observations, we observed that H₂O₂ caused a slow, but significant Ca²⁺ transient in HL-60 cells (Fig. 8A). When cells were exposed to Ec, followed by H₂O₂, we observed that H₂O₂ failed to stimulate additional release of Ca²⁺ (Fig. 8B). This observation suggests that Ca²⁺ released by Ec and H₂O₂ are probably from the same intracellular store. It is noteworthy that when cells were treated for 2 h with H₂O₂, we observed that significant levels of Ca²⁺ could still be mobilized from the ER with Tg (Fig. 8C). This observation suggests that, unlike Ec (Fig. 6), ROS alone stimulate Ca²⁺ release without fully depleting the ER of Ca²⁺, presumably because they do not block the refilling process.

View larger version (13K): [in this window] [in a new window]   Fig. 8. Ec and H2O2 both target the Tg-releasable Ca2+ store. A, 200 µM H2O2-induced changes in cytoplasmic Ca2+ concentration. Indo-1AM-loaded cells were exposed to H2O2, and cytoplasmic Ca2+ was followed over time. B, as described in A, except cells were preincubated in 15 µM Ec. C, Tg-induced release of ER Ca2+ content can still be detected in cells pretreated with 200 µM H2O2 for 2 h. Ni2+ (5 mM) was used to demonstrate that this was release of Ca2+ from intracellular stores and not Ca2+ entry.

View larger version (17K): [in this window] [in a new window]   Fig. 9. Atractyloside-generated ROS targets ER Ca2+ stores. A, HL-60 cells were incubated with 50 µM atractyloside for 2 h and then loaded with 8 µMH2DCFDA for 30 min at 37°C. H2DCFDA fluorescence was measured by flow cytometry. B, indo-1-loaded HL-60 cells were treated with 50 µM atractyloside and cytoplasmic Ca2+ followed by flow cytometry. C, cells were pretreated with 50 µM atractyloside for 2 h. ER Ca2+ content was measured by releasing ER Ca2+ into the cytoplasm with 3 µM Tg in the presence of Ni2+ to prevent Ca2+ influx.

	Discussion

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High levels of ROS have multiple effects on proteins, lipids, and nucleic acids in key organelles such as the mitochondria, ER, and nucleus. Emerging evidence also supports the concept that subtoxic levels of ROS act as endogenous signaling messengers, especially in calcium signaling and protein phosphorylation (Nathan, 2003). In the course of investigating the role of OS in ER stress, we observed that Ec, an imidazole that blocks Ca²⁺ influx and stimulates ER Ca²⁺ depletion, increased OS levels in cells. In contrast, Tg or Tu, two other agents that also induce ER stress-related cell death, did not generate increased OS. Ec was also observed to cause rapid hyperpolarization of the mitochondrial membrane. This hyperpolarization was associated with inhibition of adenine nucleotide transport, observed in cell-free mitochondria. The complex III inhibitor antimycin A neutralized Ec-induced hyperpolarization of the mitochondrial membrane, suppressed ROS increases by Ec, partially reversed ATP depletion by Ec, prevented ER Ca²⁺ depletion, and protected cells from Ec-induced cell death. The antioxidants BHT and Nac also blocked Ca²⁺ mobilization from the ER by Ec and protected from Ec-induced cell death, consistent with a role for ROS in ER depletion. Taken together, these results suggest that Ec decouples complex III from downstream steps in electron transport, resulting in relatively rapid ROS generation, inhibition of adenine nucleotide transport, and ATP depletion. The reactive oxygen produced is responsible for stimulating Ca²⁺ release from the ER. The combination of ROS generation stimulating ER Ca²⁺ mobilization with Ca²⁺ influx blockade results in the sustained ER Ca²⁺ depletion associated with cell death. This model is illustrated in Fig. 10.

View larger version (35K): [in this window] [in a new window]   Fig. 10. Model for how Ec-stimulated mitochondrial ROS contributes to ER stress. 1) Ec targets complex III in the electron transport chain resulting in hyperpolarization of the mitochondrial membrane; 2) decoupling of electron transport from adenine nucleotide transport and ATP production; 3) inhibition of ANT and ROS generation; 4) ROS stimulates ER Ca2+ release; and 5) Ec also blocks Ca2+ influx, resulting in an ER stress response.

Because antimycin A reverses Ec-induced hyperpolarization, ROS generation, ER Ca²⁺ depletion, and cell death, complex III activity is probably required for ROS generation by Ec. Furthermore, the inhibition of ANT activity by Ec suggests that electron transport does not continue down the chain to the transporter. Taken together, these observations suggest that Ec decouples complex III from complex IV, presumably by providing an alternative outlet for complex III electrons (directly or indirectly). Although further work will be required to identify how Ec targets complex III causing ROS generation, our observations show that the mitochondrial target is critical for Ec-induced cell death. We have demonstrated that cancer and leukemia cells are highly sensitive to Ec-induced cell death (Gommerman and Berger, 1998; Soboloff et al., 2002; Zhang et al., 2002). Our identification of a mitochondrial target for Ec may prove to be useful in further identifying the cellular factors associated with sensitivity to this compound.

In addition to its inhibitory effects on complex III function, antimycin A has also been reported to bind to and inhibit bcl-2 and related family members (Tzung et al., 2001). However, we suspect that this activity is unrelated to the ability of antimycin A to inhibit Ec function, because the concentration of the drug required for bcl-2 inhibition is much higher than that used in the current study. As well, we previously reported that cell death induced by Ec was unaffected by high levels of bcl-2 in cells (Gommerman and Berger, 1998), suggesting that bcl-2 does not affect the mechanism of action of Ec. Nevertheless, given the importance of bcl-2 family members and their effects on mitochondrial function, it would be of interest to further investigate a potential role for these proteins in ROS generation and Ca²⁺ depletion.

We previously demonstrated that exposure to Ec for 2 h was sufficient to initiate an irreversible cell death process. It is of interest to note that after 2 h of exposure, the mitochondrial membrane remains hyperpolarized and ATP content is reduced, but only by approximately 50%. Antimycin A exposure also results in a similarly reduced ATP level, whereas the combination of the two agents results in a slightly increased ATP content. This increase may be due to reduced ATP consumption associated with decreased ROS levels rather than a restoration of ATP production.

Cells exposed to lethal agents generally undergo variations of necrotic or apoptotic cell death (Orrenius et al., 2003). After exposure to Ec, HL-60 cells exhibit clear endpoints associated with apoptosis, such as membrane blebbing, Annexin positivity, and nuclear condensation (Zhang and Berger, 2004); however, these endpoints occur late in the cell death process. In the present study, caspase activation was observed at 18 h after Ec exposure (Fig. 7B), but not at 6 h. The inability to detect caspase activation early may be partly due to inactivation of caspases by ROS as has been documented in other cell systems (Hampton et al., 1998; Samali et al., 1999). On the other hand, apoptosis may be a secondary or indirect consequence of the ER stress process and is probably not absolutely required for cell death. We have observed, in a variety of cell systems, that 2-h exposure to Ec optimizes the difference in sensitivity between caner or leukemia cells and normal cells (Soboloff et al., 2002; Zhang et al., 2002). We have also found that ER Ca²⁺ release is stimulated at lower Ec concentrations than those required for blocking influx (Soboloff and Berger, 2002). Taken together, these observations suggest that longer term exposure to lower doses of Ec may result in a qualitatively different form of cell death that is more dependent on mitochondrial dysfunction than ER Ca²⁺ depletion.

The ER Ca²⁺ store is a dynamic entity requiring constant energy and SERCA activity to maintain the large concentration gradient across the ER membrane. It is probable that as cells undergo various forms of cell death, depleted energy levels of the cell will be insufficient to maintain the Ca²⁺ gradient resulting in indirect ER Ca²⁺ depletion (Nicotera and Orrenius, 1998). In contrast, agents such as the ANT inhibitor atractyloside and others (Le Bras et al., 2005) generate ROS and cause Ca²⁺ release from the ER rapidly, well before energy levels are depleted. If the cell can maintain sufficient energy levels, then these agents will not be toxic. However, if the oxidative burden is too high, eventually, as energy levels deplete, the cell will undergo cell death (Richter et al., 1996).

Our observations are consistent with a general role for ROS generated at the mitochondria in stimulating ER Ca²⁺ release. The full spectrum of ROS targets in the ER remains unknown. However, one likely target is the SERCA, because ROS have been reported to inhibit SERCA function (Barnes et al., 2000). Our observations that Ec and H₂O₂ both targeted the Tg-sensitive store are consistent with the concept that SERCA is the ROS target responsible for ER Ca²⁺ release.

Both antimycin A and the antioxidants neutralized the ability of Ec to deplete the ER with no discernible effect on Ec-induced inhibition of Ca²⁺ influx. Furthermore, antimycin A and the antioxidants provided significant protection from cell death and caspase activation induced by Ec. Likewise, although atractyloside caused Ca²⁺ release from the ER, this did not lead to ER Ca²⁺ depletion or significant cell death. These observations are in agreement with our model that sustained Ca²⁺ depletion of the ER is required for lethality (Soboloff and Berger, 2002; Zhang and Berger, 2004). This would seem to suggest that although Ec releases ER Ca²⁺ content via ROS just like atractyloside, blocking store-operated Ca²⁺ is required for its toxicity and conversely, blocking Ca²⁺ entry is only fully toxic when ER Ca²⁺ content is depleted. This is consistent with our previous observation in mast cells and breast cancer cells, where Ec-induced cell death was strongly enhanced by the addition of agents that stimulate Ca²⁺ release (Gommerman and Berger, 1998; Soboloff and Berger, 2002; Zhang et al., 2002).

Large increases in cytoplasmic Ca²⁺ caused by ER depletion coupled with influx can result in rapid increases in mitochondrial Ca²⁺. Excessive mitochondrial Ca²⁺ has been linked to activation of the mitochondrial permeability transition and induction of apoptosis (Orrenius et al., 2003). Our results suggest that ROS can perform the opposite function, namely, communicating mitochondrial dysfunction to the ER through the stimulation of Ca²⁺ release.

	Footnotes

 
This work was supported by grants from the National Cancer Institute of Canada (to S.A.B.).

ABBREVIATIONS: ER, endoplasmic reticulum; SERCA, sarco(endo)plasmic reticulum Ca²⁺ ATPase; Tg, thapsigargin; Ec, econazole; Atra, atractyloside; ROS, reactive oxygen species; OS, oxidative stress; Tu, tunicamycin; FBS, fetal bovine serum; AA, antimycin A; BHT, butylated hydroxytoluene; Nac, N-Acetyl-L-cysteine; PI, propidium iodide; CM-H₂DCFDA, 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate; AM, acetoxymethyl ester; MMP, mitochondrial membrane potential; JC-1, 5,5'6,6'-tetraethylbenzimidazolcarbocyanine iodide; FCCP, carbonyl cyanide p-(trifluomethoxy) phenylhydrazone; ANT, adenine nucleotide transporter; PBS, phosphate-buffered saline; PMA, phorbol 12-myristate 13-acetate; DCF, 2',7'-dichlorofluorescein.

¹ Current affiliation: Department of Hematology, Tongji Hospital, Tongji Medical College, and Huazhong University of Science and Technology, Wuhan China, 430030.

² Current affiliation: Department of Biochemistry and Molecular Biology, University of Maryland School of Medicine, Baltimore, MD 21201.

Address correspondence to: Dr. Stuart A. Berger, Arthritis and Immune Disorder Research Centre, University Health Network, Toronto Medical Discovery Tower, 8th Floor, Rm 8-356, 101 College St., Toronto, ON, Canada M5G 1L7. E-mail: berger{at}uhnres.utoronto.ca

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