* Division of Pharmacology and Toxicology, College of Pharmacy, University of Texas at Austin, Austin, Texas 78712; Department of Pharmacology and Toxicology, College of Pharmacy, University of Arizona Health Science Center, Tucson, Arizona 85721
1 To whom correspondence should be addressed at Department of Pharmacology and Toxicology, College of Pharmacy, University of Arizona Health Sciences Center, 1703 E. Mabel St., Tucson, AZ 85721. E-mail: scouser{at}pharmacy.arizona.edu.
Received January 10, 2005; accepted March 18, 2005
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ABSTRACT |
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INTRODUCTION |
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By regulating mitochondrial function, the Bcl-2 family members are key players in stress/toxicantinduced apoptosis. The Bcl-2 family members are divided into two subgroups, based on their ability to either promote or antagonize apoptosis (Chao and Korsmeyer, 1998; Gross et al., 1999
; Wei et al., 2001
). In general, the anti-apoptotic Bcl-2 family members, such as Bcl-2, Bcl-xL, and Bcl-w, are integrated into the mitochondrial membrane, whereas the pro-apoptotic Bcl-2 family members, such as Bad and Bax, are localized in the cytosol in the absence of a death signal. The pro-apoptotic proteins are upregulated in response to death signals, and they translocate into mitochondrial membranes, where they interact with the anti-apoptotic members, contributing to apoptosis. Cell survival signals can also influence the Bcl-2 family of proteins. For example, IL-3 activates the Akt survival signaling pathway, resulting in phosphorylation of pro-apoptotic Bad, which subsequently binds to, and is sequestered by 1433 (Datta et al., 2000
; Zha et al., 1996
; Zhou et al., 2000
). Thus, the combination of post-translational modifications and the upregulation or downregulation of pro-apoptotic and anti-apoptotic protein expression determines the anti-apoptotic or pro-apoptotic function of the Bcl-2 family members.
Benzene is an industrial solvent used to produce plastics, nylon and other synthetic fibers, lubricants, and dyes; it is also a natural component of gasoline and cigarette smoke (Imbriani et al., 1995). Benzene causes bone marrow suppression in rodents and is both hematotoxic and leukemogenic in humans, leading to hematological disorders, such as aplastic anemia and acute myelogenous leukemia (Golding and Watson, 1999
; Rinsky et al., 1981
; Tunek et al., 1981
). Benzene must be metabolized and bioactivated to mediate its toxic effects. A number of hydroquinone-thioether metabolites have been identified in the bone marrow of rats and mice exposed to a combination of hydroquinone and phenol, or benzene (Bratton et al., 1997
). In particular, 2,3,5-tris(glutathion-S-yl)hydroquinone (TGHQ) causes hematotoxicity in rats, and it induces apoptosis in human promyelocytic leukemia (HL-60) cells (Bratton et al., 1997
, 2000
). TGHQ likely induces toxicity either by the generation of reactive oxygen species (ROS) or via the covalent binding of reactive metabolites to critical tissue macromolecules, or both (Bratton et al., 1997
). Prior to the onset of apoptosis, TGHQ depletes cellular glutathione (GSH) levels and stimulates sphingomyelin turnover (Bratton et al., 2000
). However, the mechanisms by which TGHQ induces apoptosis have not been fully elucidated.
In the present study we demonstrate that TGHQ facilitates ROS production, an essential element for inducing apoptosis, and stimulates cytochrome c release from mitochondria, leading to caspase activation, in the absence of a disruption in the mitochondrial membrane potential. Mitochondrial cytochrome c release therefore appears to be mediated by the ability of TGHQ to stimulate the subcellular relocation of pro-apoptotic Bax protein, which translocates to mitochondria, and in combination with the dephosphorylation of Bcl-2 protein, promotes cytochrome c release, contributing to TGHQ-induced apoptosis of HL-60 cells.
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MATERIALS AND METHODS |
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Cell lines and culture conditions.
HL-60 cells were maintained in Roswell Park Memorial Institute medium (RPMI 1640; Gibco BRL, Grand Island, NY) containing 20% fetal bovine serum (FBS) in a 37°C, 5% CO2-regulated incubator. Cells were routinely cultured at a density of 1.0 x 106 cells/ml. Immediately prior to all experiments, cells were washed and resuspended in RPMI 1640 containing 25 mM HEPES and 10% fetal bovine serum (FBS).
Annexin V-FITC/propidium iodide apoptosis assay.
The percentage of apoptotic cells was determined according to the manufacturer's protocol using an annexin V FITC kit and an EPICS XL-MCL (Coulter, Miami, FL) flow cytometer.
ROS measurement using carboxy-H2DCFDA.
Cells were loaded with 20 µM 5-( and-6)-carboxy-2',7'-dichlorodihydrofluorescein diacetate (Carboxy-H2DCFDA) (Molecular Probes) for 30 min. After TGHQ (200 µM) treatment, cells were pelleted, washed, and resuspended in PBS. Then, 5-(and-6)-carboxy-2',7'-dichlorofluorescein (carboxy-DCF) fluorescence was determined on an EPICS XL-MCL flow cytometer, with excitation at 495 nm and emission at 525 nm.
Measurement of mitochondrial inner transmembrane potential (m).
Cells were treated with TGHQ (200 µM) and collected at each time point. Cells were resuspended in 500 µl of PBS and labeled with TMRM (final concentration 150 nM). As a positive control, aliquots of cells were stained in the presence of 100 µM CCCP. Cells were subsequently incubated at 37°C for 30 min and returned to ice. Fluorescence was determined on an EPICS XL-MCL flow cytometer, with excitation at 495 nm and emission at 575 nm.
Preparation of nuclear and cytosolic extracts.
After treatment of cells with TGHQ, nuclear and cytosolic extracts were prepared as described (Read et al., 1994), with slight modifications. Cells were harvested, washed with cold PBS, and resuspended in 100 µl of buffer A (10 mM HEPES [pH 8.0], 1.5 mM MgCl2, 10 mM KCl, 0.5 mM dithiothreitol [DTT], 200 mM sucrose, 0.5 mM phenylmethylsulfonyl fluoride [PMSF], 1 mM Na3VO4, 10 mM sodium glycerophosphate, 50 mM NaF, 5 mM sodium pyrophosphate and 0.5% Nonidet P-40) with protease inhibitors. Suspended cells were incubated for 5 min at 4°C. The lysed cells were microcentrifuged at 16,000 x g for 15 s at 4°C. The supernatants (cytosolic fraction) were saved, and the pellet (nuclear fraction) was rinsed once in buffer A. The nuclear extracts were resuspended in 40 µl of buffer B (20 mM HEPES [pH 7.9], 1.5 mM MgCl2, 420 mM NaCl, 0.2 mM EDTA, 0.5 mM PMSF, 1 mM DTT, 1 mM Na3VO4, 10 mM sodium glycerophosphate, 50 mM NaF, 5 mM sodium pyrophosphate) with protease inhibitors. The nuclear extracts were incubated at 4°C for 30 min and then microcentrifuged at 16,000 x g for 10 min. The resulting supernatants were diluted 1:1 with buffer C (20 mM HEPES [pH 7.9], 100 mM KCl, 0.2 mM EDTA, 20% glycerol, 1 mM DTT and 0.5 mM PMSF) with protease inhibitors. The cytosolic extracts were clarified by microcentrifugation at 14,000 x g for 30 min. Protein concentration was determined by the modified Lowry method (Biorad protein assay, Bio-Rad).
Preparation of mitochondria-enriched and cytosolic fractions.
The mitochondria-enriched and cytosolic extracts were obtained as described elsewhere (Ganju and Eastman, 2002), with slight modifications. Briefly, 6 x 106 cells were incubated in ice-cold lysis buffer (75 mM NaCl, 1 mM NaH2PO4, 8 mM Na2HPO4, 250 mM sucrose, 1 mM Na3VO4, 10 mM sodium glycerophosphate, 50 mM NaF, 5 mM sodium pyrophosphate and protease inhibitors) with 26.25 µg digitonin in 100 µl on ice for 20 min, followed by centrifugation at 12,000 x g for 1 min. The supernatants were saved as the cytosolic fraction, and the pellet (mitochondria-enriched fraction) was resuspended in the same volume of lysis buffer without digitonin. The pellet was solubilized by sonication for 10 s at 4°C and stored as the mitochondria-enriched fraction.
Preparation of total cell extracts.
RIPA buffer (10 mM Tris [pH 7.5], 150 mM NaCl, 5 mM ethylene diamine tetraacetic acid [EDTA], 0.1% sodium dodecyl sulfate [SDS], 0.5% deoxycholate, 1% Nonidet P-40 containing 0.5 mM PMSF, 1 mM Na3VO4, 10 mM sodium glycerophosphate, 50 mM NaF, 5 mM sodium pyrophosphate and protease inhibitors) was used to obtain total cell extracts. The cells were incubated for 30 min at 4°C in RIPA buffer followed by centrifugation for 20 min at 14,000 x g. The supernatants were used as total cell extracts.
Western blotting.
Samples were mixed with sample buffer (Laemmli sample buffer, Bio-Rad) and then heated for 5 min at 100°C and loaded onto 10% or 12% SDS-polyacrylamide gels. After they were electroblotted onto PVDF membranes, the sample blots were blocked for 1 h with 5% nonfat milk powder in TBS-T solution (25 mM Tris-HCl [pH 7.6], 0.2 M NaCl, and 0.1% v/v Tween 20) at room temperature. Membranes were incubated with the antibodies of interest overnight at 4°C, unless stated otherwise. After washing three times in TBS-T for 5 min, membranes were incubated with horseradish-conjugated secondary antibodies. Bound antibodies were detected by enhanced chemiluminescence (ECL kit, Amersham, Arlington Heights, IL).
Caspase 9 activity.
HL-60 cells were treated with 200 µM TGHQ for each time period (0, 0.25, 0.5, 1, and 2 h) and collected by centrifugation at 1000 x g for 3 min. Cells were washed with PBS twice. Cells were resuspended with extraction buffer (50 mM Tris, 150 mM NaCl, 0.5 mM EDTA, and 0.5% Nonidet P-40) and incubated on ice for 10 min. The cell lysates were obtained by centrifugation at 1000 x g for 5 min. Caspase 9 activity was measured according to the manufacturer's instructions provided in the fluorometric assay kit from Oncogene (San Diego, CA).
Statistical analysis.
Data are presented as the mean ± SD. Statistical significance was determined by analysis of variance (ANOVA), followed by Tukey's post hoc comparison.
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RESULTS |
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DISCUSSION |
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Caspases 9 and 3 are both activated in HL-60 cells exposed to TGHQ (Figs. 2 and 3). Caspase activation represents the "irreversible" or execution stage of apoptosis, because caspase-mediated proteolysis is irreversible (Thornberry and Lazebnik, 1998). Many proteins required for the maintenance of cell structure and function are substrates of active caspase 3. During TGHQ-induced apoptosis of HL-60 cells, lamin B, a nuclear structural protein, is cleaved by active effector caspases (Fig. 3B), leading to the disruption of nuclear architecture. Because lamin B is reported to be cleaved predominantly by caspase 6, we can assume that caspase 6 is activated in TGHQ-treated HL-60 cells. In addition, HPK 1, which is predominantly expressed in hematopoietic tissue, is also a substrate for active caspase 3 (Fig. 3C). This finding is consistent with the report that intact HPK 1 and its caspase 3cleaved form may support apoptosis of T lymphocytes (Schulze-Leuhrmann et al., 2002
). In particular, the caspase 3cleaved C terminal of HPK 1 blocks I
B-
degradation, inhibiting the anti-apoptotic function of NF-
B. Caspase 8, an initiator caspase in the death receptormediated signaling pathway, is also a substrate of active effector caspases during TGHQ-induced apoptosis (Fig. 4); the significance of this remains unclear, however. We assume that caspase 6 is activated during TGHQ-induced apoptosis, because lamin B cleavage is observed. In addition, caspase 6 can cleave caspase 3 directly (Cowling and Downward, 2002
). Thus, it is possible that activated caspase 6 cleaves caspase 8 directly during TGHQ-induced apoptosis of HL-60 cells. Another possibility is that active caspase 3 acts upstream of caspase 8. It has recently been suggested that caspase 3 and caspase 8 are associated with the Fas-associated death domain in lipid rafts, and that caspase 3 is required for complete caspase 8 activation during Fas-mediated cell death (Aouad et al., 2004
).
Mitochondria play an important role during stress/toxicantinduced apoptosis (Green and Reed, 1998; Wang, 2001
), because mitochondria contain many apoptosis-stimulating elements, including cytochrome c and Smac. In most cases of toxicant-induced apoptosis, cytochrome c is required for caspase 9 activation. Cytochrome c may be released from mitochondria into cytosol in several ways (Bratton and Cohen, 2001
; Hengartner, 2000
; Ly et al., 2003
). One possible mechanism involves changes in the PTP, in which a loss in
m occurs. The PTP is believed to consist of the mitochondrial outer membrane voltage-dependent anion channel, the inner membrane adenine nucleotide translocase, and the mitochondrial benzodiazepine receptor. The adenine nucleotide translocase is associated with cyclophilin D. According to this mechanism, the apoptosis-inducing agent causes an opening of the PTP, the dissipation of
m, and the release of cytochrome c. However, in TGHQ-treated HL-60 cells, cytochrome c is released from mitochondria 2 h after TGHQ treatment (Fig 2C), a time at which the
m is still maintained (Fig. 5). In addition, pretreatment of HL-60 cells with cyclosporine A had little effect on TGHQ-induced apoptosis (Fig. 6). Alternatively, Bcl-2 family members may form a channel through which cytochrome c and other molecules can escape from mitochondria. Recently, Bax was found to translocate, oligomerize, and form a protein-permeable pore (Antonsson et al., 2000
; De Giorgi et al., 2002
; Gross et al., 1998
). In addition, Bax may form a Bax/voltagedependent anion channel hybrid channel to release cytochrome c (Shimizu et al., 1999
). HL-60 cells exposed to TGHQ upregulate the pro-apoptotic Bax protein, which is subsequently translocated to mitochondria (Fig. 8). Moreover, TGHQ promotes the dephosphorylation of the anti-apoptotic S70 Bcl-2 protein (Fig. 7), suggesting that the redistribution of cytochrome c from the mitochondria to the cytosol during TGHQ-induced apoptosis of HL-60 cells may result not from the opening of PTP but via the formation of a conducting channel.
Debate continues over whether phosphorylation of Bcl-2 enhances or inhibits its anti-apoptotic function (Deng et al., 2004; Haldar et al., 1998
; Ruvolo et al., 2001
; Yamamoto et al., 1999
). Phosphorylation of Bcl-2 at Ser70 correlates with increases in cell survival in chemotherapeutic druginduced apoptosis (Ito et al., 1997
; May et al., 1994
). In contrast, paclitaxel induces Bcl-2 phosphorylation at Ser70 and causes cell death (Haldar et al., 1996
, 1998
). Korsmeyer's group identified the phosphorylation sites of Bcl-2 at Ser70, Ser87 and Thr69 and suggested that phosphorylation inactivates Bcl-2 (Yamamoto et al., 1999
). However, even in experiments in which a series of serine/threonine (S/T)
glutamate/alanine (E/A) mutants were created to mimic or abrogate phosphorylation, the phosphorylation of Bcl-2 had different effects on paclitaxel-induced apoptosis in different cell types. Such differences in the role of Bcl-2 phosphorylation on apoptosis are thus likely context specific and dependent on the initiating apoptotic insult. The role of the phosphorylation status of Bcl-2 must therefore be assessed in a context-specific fashion. In TGHQ treated HL-60 cells, dephosphorylation of S70 Bcl-2 may block the anti-apoptotic function of Bcl-2 and contribute to the apoptotic process. In support of this view, in TGHQ-treated HL-60 cells, GSH depletion induces sphingomyelin turnover and increases ceramide concentrations (Bratton et al., 2000
). Ceramide activates mitochondrial phosphatase 2A, which dephosphorylates Bcl-2, leading to apoptosis (Ruvolo et al., 1999
, 2002
). Collectively, TGHQ treatment may increase cellular ceramide, which subsequently activates phosphatase 2A and dephosphorylates anti-apoptotic Bcl-2 protein, which, in association with translocated Bax, contributes to the release of cytochrome c from mitochondria into the cytosol (Fig. 2C).
In conclusion, TGHQ-induced apoptosis of HL-60 cells requires the generation of ROS, and decreases in GSH concentrations concomitant with an increase in ceramide. Ceramide-mediated activation of phosphatase 2A may then promote the dephosphorylation of Bcl-2, which, in combination with Bax, can then support mitochondrial cytochrome c release and activation of the caspase cascade (Fig. 9).
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NOTES |
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ACKNOWLEDGMENTS |
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