4-Hydroxynonenal Induces Apoptosis via Caspase-3 Activation and

Jul 31, 2001 - Vanderbilt University School of Medicine, Nashville, Tennessee 37232. Chem. Res. Toxicol. , 2001, 14 (8), .... James D. West, Chuan Ji,...
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4-Hydroxynonenal Induces Apoptosis via Caspase-3 Activation and Cytochrome c Release Chuan Ji,† Ventkataraman Amarnath,‡ Jennifer A. Pietenpol,† and Lawrence J. Marnett*,† Departments of Biochemistry and Pathology, Center in Molecular Toxicology, and the Vanderbilt-Ingram Cancer Center. Vanderbilt University School of Medicine, Nashville, Tennessee 37232 Received August 23, 2000

We investigated the mechanism by which 4-hydroxynonenal (HNE), a major aldehydic product of lipid peroxidation, induces apoptosis in tumor cells. Treatment of human colorectal carcinoma (RKO) cells with HNE-induced poly-ADP-ribose-polymerase (PARP) cleavage and DNA fragmentation in a dose- and time-dependent manner. The induction of PARP cleavage and DNA fragmentation paralleled caspase-2, -3, -8, and -9 activation. Pretreatment of cells with an inhibitor of caspase-3, z-DEVD-fmk, or a broad spectrum caspase inhibitor, z-VAD-fmk, abolished caspase activation and subsequent PARP cleavage. Constitutive expression of high levels of Bcl-2 protected cells from HNE-mediated apoptosis. In addition, Bcl-2 overexpression inhibited cytochrome c release from mitochondria and subsequent caspase-2, -3, and -9 activation. These findings demonstrate that HNE triggers apoptotic cell death through a mitochondrion-dependent pathway involving cytochrome c release and caspase activation. Bcl-2 overexpression protected cells from HNE-induced apoptosis through inhibition of cytochrome c release.

Introduction 4-Hydroxynonenal (HNE)1 is a major aldehydic product of lipid peroxidation (1) and is believed to contribute significantly to the cytopathological effects observed during oxidative stress (1-4). HNE exerts these effects because of its high reactivity with biological molecules, particularly those containing sulfhydryl and amino groups (e.g., DNA polymerases and dehydrogenases) (1, 5, 6). HNE causes different cytohistopathological effects including apoptotic cell death in a wide range of cell types (7-10). Two major pathways of apoptosis have been identified in recent years. One pathway is Fas/Fadd-dependent (11, 12). Fas is a receptor protein that contains a protein interaction module known as the death domain, whereas Fadd is an adaptor protein that contains both a death domain and the death effector domain. Fas/Fadd interact with the receptor-associated death proteases (procaspases-8 and -10) leading to activation of downstream effector caspases (11, 12). The second pathway of apoptosis is mitochondrion-dependent and results from release of cytochrome c leading to caspase activation through the apoptotic protease-activating factor-1 (13-17). The Bcl-2 * To whom correspondence should be addressed. Phone: (615) 343-7329. Fax: (615) 343-7534. E-mail: marnett@ toxicology.mc.vanderbilt.edu. † Department of Biochemistry. ‡ Department of Pathology. 1 Abbreviations: HNE, 4-hydroxynonenal; PARP, poly-ADP-ribosepolymerase; PBS, phosphate-buffered saline; SDS-PAGE, sodium dodecyl sulfate-polyacryamide gel electrophoresis; z-VAD-fmk, carbobenzoxy-Val-Ala-Asp-fluoromethyl ketone; z-DEVD-fmk, carbobenzoxy-Asp-Glu-Val-Asp-fluoromethyl ketone; RKO, human colorectal carcinoma cells; RKO-φ cells, RKO cells transfected with empty vector, pCEP4; RKO-Bcl-2 cells, RKO cells transfected with pCEP4 vector containing a cDNA encoding human Bcl-2.

family of proteins plays a major role in regulating this mitochondrion-dependent pathway (18-21). Although studies have shown that HNE-induced apoptosis correlates with extracellular calcium uptake (22, 23), c-Jun amino-terminal kinase activation (24), glutathione deletion (8, 9), and a Fas-independent caspase cascade (10), the involvement of the major apoptotic pathway initiated by HNE remains unclear. Caspases are a family of cysteine proteases responsible for promoting cell death (25-27). Present as inactive procaspases, most caspases are activated following cleavage at a specific aspartate site by assembly of their active subunit forms (25). Caspase-9 activation requires the release of cytochrome c from mitochondria to cytosol where the caspase resides (26). Caspase-3 activation is effected through proteolytic cleavage by caspase-9 or caspase-8 (25). Caspase-3 may then cleave cellular target proteins including poly-ADP-ribose-polymerase (PARP) (25, 26). In the present study, we investigated the mechanism of HNE-induced apoptotic cell death in human colorectal carcinoma cells (RKO). The results demonstrate that HNE induces apoptosis by inducing alteration of mitochondrial function leading to the release of cytochrome c and subsequent activation of the caspase cascade. Overexpression of Bcl-2 inhibited HNE-induced release of mitochondrial cytochrome c and protected cells from HNE-induced apoptosis.

Material and Methods Cell Culture Conditions and Chemical Treatment. RKO cells were maintained in McCoy’s 5A medium (Gibco/BRL, Gaithersburg, MD), supplemented with 10% bovine serum (Hyclone, Logan, UT), 100 units/mL penicillin, and 100 µg/mL

10.1021/tx000186f CCC: $20.00 © 2001 American Chemical Society Published on Web 07/31/2001

4-Hydroxynonenal Induces Apoptosis streptomycin and incubated in a 5% CO2/95% air incubator at 37 °C. Cells were plated 24 h prior to chemical treatment and were 50-70% confluent at the time of treatment. HNE was dissolved in ethanol. Synthetic peptide inhibitors: Z-Val-AlaAsp fluoromethyl ketone (z-VAD-fmk) and Z-Asp-Glu-Val-Asp fluoromethyl ketone (z-EVD-fmk) were purchased from BioVision (Palo Alto, CA) and dissolved in ethanol. The final concentration of ethanol in the medium was e0.1%. Cell Transfection and Establishment of Cell Lines Ectopically Expressing Bcl-2. Lipofectamine (Gibco/BRL) was used to transfect cells with DNA. Lipofectamine (36 µg) dissolved in Opti-MEM (Gibco/BRL) in a total volume of 250 µL was mixed with an equal volume of Opti-MEM, containing either pCEP4 (Invitrogen, San Diego, CA), a vector containing a hygromycin resistance gene, or the same vector containing a cDNA encoding human Bcl-2 (28). The mixtures were incubated at room temperature for 20 min, then added to RKO cells, which were 30-50% confluent in 2.5 mL of supplement-free medium. The cells were incubated at 37 °C for 6 h prior to the addition of 3 mL of complete medium then incubated overnight. Hygromycin (150 µg/mL) (Calbiochem-Novabiochem Corp, LA Jolla, CA) selection was carried out after the cells recovered in complete medium for 24 h. The stable transfected colony pool was obtained following 2-3 weeks of hygromycin selection. Morphological Assessment of Necrotic and Apoptotic Cells. After HNE treatment, attached and floating cells were harvested, stained with trypan blue, and counted for trypan blue-excluding (viable) and trypan blue-stained cells (necrotic). For assessment of apoptotic cells, cells were grown on collagencoated glass coverslips in a 6-well plate 18-24 h prior to treatment. A thin overlay of 1% low-melting soft agar was placed on the monolayer immediately before adding HNE to prevent cells from detaching. After staining with DAPI (4,6-diamino-2phenylindole), apoptotic bodies and normal nuclei were visualized by fluorescence microscopy using excitation and emission filters of 380 and 460 nm, respectively, at a magnification of 250×. At least 300 cells in four high-power fields were counted, and apoptotic cells were expressed as percentage of total cells. Cellular GSH Level Assessment. The method for measuring GSH levels is based on the ability of GSH to react with monochlorobimane in the presence of glutathione S-transferase and to form a fluorescenct adduct that can be detected with a fluorometer (9). After HNE treatment, cells were washed twice with PBS and incubated with 2 mL of 200 µM monochlorobimane in PBS at 37 °C for 30 min. Cells were then washed once with PBS and lysed in a hypotonic solution (equal volume of distilled water and PBS) containing 0.5% Triton X-100. Cell lysates were washed once with dichloromethane and the aqueous solution was collected and assayed for GSH levels using excitation and emission wavelengths at 398 and 488 nm, respectively. Standard curves were prepared by incubating 0-32 nmol of GSH with 200 µM monochlorobimane in the presence of 32 nM glutathione S-transferase at 37 °C for 30 min. Preparation of Protein Lysates and Western Blotting Analysis. The preparation of cell lysates and Western blotting procedures were as previously described (29) with minor modifications. Briefly, cells were washed twice with ice-cold PBS and lysed in kinase lysis buffer [50 mM Tris buffer (pH 7.5) 150 mM NaCl, 0.1% Triton X-100, 0.1% Nonidet P-40, 4 mM EDTA, 50 mM NaF, 0.1 mM sodium orthovanadate, 1 mM DTT, and protease inhibitors: antipain, leupeptin, pepstatin A and chymostatin (5 µg/mL), phenylmethanesulfonyl fluoride (50 µg/mL) and 4-(2-aminoethyl)-benzenesulfonylfluoride (100 µg/mL)] for 30 min at 4 °C. Cell lysates were cleared by centrifugation at 15000g for 15 min, and the resulting supernatant was collected. Cellular protein (30-50 µg) was mixed with an equal volume of 2× Laemmli sample buffer [125 mM Tris (pH 6.8), 10% β-mercaptoethanol, 20% glycerol, 4% SDS, and 0.05% bromophenol blue] and boiled for 5 min. The proteins were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and electrophoretically transferred onto polyvinylidene difluoride membranes (Millipore, Bedford, MA). The

Chem. Res. Toxicol., Vol. 14, No. 8, 2001 1091 membranes were blocked with 5% nonfat milk in Tris-buffered saline [Tris 50 mM (pH 7.5), NaCl 150 mM) containing 0.1% Tween 20, then incubated with anti-Bcl-2, anti-caspase-9, anticaspase-2, anti-caspase-8 (Santa Cruz, Santa Cruz, CA), antiCPP32, anti-cytochrome c, or anti-poly (ADP-ribose) polymerase (PharMingen, San Diego, CA) for 1-2 h. The primary antibody was then stained with either donkey anti-rabbit or goat antimouse horseradish peroxidase-conjugated secondary antibodies. Enhanced chemiluminescence was performed (ECL Western blotting detection system, Amersham, Arlington Heights, IL) and protein bands were detected by autoradiography. Determination of DNA Fragmentation. Cells were harvested by scraping, washed twice with ice-cold PBS, and lysed in lysis buffer [10 mM Tris-HCl (pH 8.0), 10 mM EDTA, and 0.2% Triton X-100] on ice for 30 min. After centrifugation at 13000g for 10 min, the resulting supernatant containing soluble DNA fragments was collected. The supernatant was incubated with RNase A (200 µg/mL) at 37 °C for 1 h, then incubated with proteinase K (1 mg/mL) with 1% SDS solution at 50 °C for 2 h. The soluble DNA was cleared by phenol extraction and ethanol precipitation. DNA pellets were dissolved in TE buffer, and resolved on 1.5% agarose gel. DNA fragments were visualized by staining with ethidium bromide. Preparation of Cytosol and Mitochondria. Cells were washed twice with ice-cold PBS and resuspended in buffer A [20 mM HEPES (pH 7.5) 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, and protease inhibitors] containing 250 mM sucrose. The cells were broken with 8 strokes of a Teflon homogenizer, and the homogenates were centrifuged at 750g for 10 min at 4 °C. The supernatants were further centrifuged at 15000g for 20 min. The pellets (mitochondria) were redissolved in buffer A containing 250 mM sucrose and the resulting supernatants were reserved as cytosol. Caspase-3 Colorimetric Assay. Cells were washed twice with ice-cold PBS and lysed in a lysis buffer (BioVision) on ice for 10 min, followed by centrifugation at 15000g for 10 min. Caspase-3 activity in the supernatant was determined by a colorimeric assay kit (BioVision) using the p-nitroanilide-labeled peptide, DEVD-pNA, as substrate. Caspase-3 activity was monitored by the release of p-nitroanilide from the substrate at 405 nm. The fold increase in caspase-3 activity was calculated by comparing the absorbance of p-nitroanilide from untreated controls with HNE-treated apoptotic samples.

Results HNE Induces RKO Cell Death in a Dose- and Time-Dependent Manner. The ability of HNE to induce apoptosis in RKO cells was analyzed by assessment of plasma membrane integrity, morphological changes, DNA fragmentation, and PARP cleavage assays. During necrosis, the plasma membrane transport function fails, resulting in cells that cannot exclude trypan blue or other dyes (30). Recognizable features of apoptotic cells include chromatin condensation, nuclear disintegration and fragmented nuclei. HNE treatment (30-60 µM) of RKO cells, led to a dose-dependent increase in apoptotic and necrotic cells (Figure 1A). At the higher concentration of 75 µM HNE, there was a decline in the number of apoptotic cells and an increase in necrotic cells in the culture. DNA fragmentation was detectable after 30 µM HNE treatment and reached a maximum at 4560 µM HNE (Figure 1B). Protein extracts were prepared from RKO cells treated with 60 µM HNE and PARP cleavage analyzed by Western blotting. An 85 kDa, cleaved PARP species was detectable by 15 h after HNE treatment, which increased in levels through 25 h, and declined thereafter (Figure 1C). To determine if HNEinduced changes in cellular glutathione (GSH) levels that correlated with the onset of cell death, GSH levels were

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Figure 1. HNE induces RKO cell death in a dose- and time-dependent manner. (A) RKO cells were treated with the indicated concentrations of HNE for 24 h. Cells were stained with trypan blue or DAPI for assessment of necrotic and apoptotic cell phenotypes. Cells were considered necrotic, if trypan blue positive, and apoptotic, if chromatin condensation and nuclear fragmentation were evident. In each case, at least 300 cells were counted. (B) Cells were treated with the increasing doses of HNE for 24 h and DNA fragmentation was analyzed by 1.5% agarose gel electrophoresis. DNA bands were visualized by staining with ethidium bromide. (C) Cells were treated with 60 µM HNE for the indicated times and PARP cleavage in total cell lysates (50 µg/lane) was analyzed by Western blotting. The results are representative of three independent experiments. (D) The cells were treated with 30 µM or 60 µM HNE for the indicated times. After reacting with monochlorobimane, GSH levels were evaluated by a fluorometric method described in the Materials and Methods. The results are representative of the mean ( SD from four experiments.

also measured after HNE exposure. After treatment of cells with 60 µM HNE, a concentration that induced maximal apoptosis, there was a ∼75% reduction in GSH levels by 2 h that returned to control levels by 4 h, a time preceding the onset of apoptosis (Figure 1D). HNE Induces Dose- and Time-Dependent Activation of Caspase-3. The induction of PARP cleavage and DNA fragmentation suggests that HNE treatment causes caspase-3 activation. Therefore, RKO cells were treated with increasing doses of HNE for 24 h, and caspase-3 activity was measured using a specific substrate of caspase-3 (DEVD-pNA) in an in vitro assay. HNE treatment resulted in a 2- and 6-fold increase in caspase-3 activity at 45 and 75 µM doses, respectively (Figure 2A). A similar pattern of caspase-3 activation was observed when cells were treated with 60 µM HNE and incubated for various time periods (Figure 2B). The activity of caspase-3 gradually increased with time reaching a maximum at 20-25 h. These results are in agreement with the findings shown in Figure 1, suggesting that caspase-3 activation is a key event in the induction of apoptosis by HNE.

Caspase Inhibitors Inhibit Caspase-3 Activation Induced by HNE. To confirm that caspase-3 is responsible for HNE-induced apoptosis, two alternate approaches were investigated. First, we analyzed cell lysates for the presence of the active form of caspase-3 (e.g., the 17 kDa form resulting from the cleavage of procaspase-3) following HNE treatment, then we determined the effect of co-administration of caspase inhibitors with HNE on the appearance of this species. RKO cells were treated with either HNE alone or HNE together with z-DEVD-fmk (a relatively specific caspase-3 inhibitor) or z-VAD-fmk (a general caspase inhibitor). After 24 h, PARP cleavage was seen only in the cells treated with HNE alone. PARP cleavage correlated with the conversion of the inactive 32 kDa pro-caspase-3 into the active 17 kDa caspase. Treatment of cells with caspase inhibitors abolished the activation of pro-caspase-3 and subsequent PARP cleavage (Figure 3). These findings suggest that caspase-3 is responsible for the apoptotic events in these cells. Overexpression of Bcl-2 Protein Blocks Caspase-3 Activation and Subsequent PARP Cleavage In-

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Figure 3. Caspase inhibitors block caspase-3 activation and subsequent PARP cleavage. RKO cells were treated with HNE (60 µM) or HNE containing Z-DEVD-fmk or Z-VAD-fmk (20 µM) for 24 h. Total cell lysates (50 µg/lane) were analyzed for the cleaved bands of PARP and pro-caspase-3 by Western blot. Results are representative of two independent experiments.

Figure 2. HNE induces dose- and time-dependent caspase-3 activation. (A) RKO cells were treated with the indicated concentrations of HNE for 24 h. (B) The cells were treated with 60 µM HNE for the indicated times. Caspase-3 activities in the cell lysates (75-100 µg) were evaluated in vitro using DEVDpNA as substrate. The fold increase was calculated by comparing the absorbance of p-nitroaniline at 405 nm from untreated controls with the values from HNE-treated samples. The results are representative of the mean of duplicate determinations from one of three independent experiments.

duced by HNE. Bcl-2 overexpression can block apoptosis induced by multiple cancer chemotherapeutic agents in a variety of cell types. To determine if overexpression of Bcl-2 could prevent HNE-induced apoptosis, RKO cells were stably transfected with an expression vector containing a full-length human Bcl-2 (RKO-Bcl-2). Parallel cultures of cells containing an empty vector were generated for use as controls (RKO-φ). Following treatment of cells with HNE (60 µM) for various time periods, total cellular protein was analyzed for PARP cleavage, Bcl-2 expression, and caspase-3 activation by immunoblot analysis (Figure 4). In RKO-φ cells, PARP cleavage occurred within 8 h and reached a maximum at 24 h. This paralleled the conversion of the inactive 32 kDa procaspase-3 into the active 17 kDa caspase. Of note, the level of endogenous Bcl-2 protein appeared to decline after HNE treatment of these cells. In contrast, PARP was not significantly cleaved in the RKO-Bcl-2 cells, an observation consistent with the absence of the active 17 kDa caspase-3 (Figure 4). Similar results were obtained at different doses of HNE (data not shown). As described above for RKO cells, treatment of the control RKO-φ cells with HNE resulted in the activation of caspase-3 in a dose-dependent manner. In contrast, no active caspase-3 or PARP cleavage was detected in RKO-Bcl-2 cells. These

Figure 4. Overexpression of Bcl2 protein blocks caspase-3 activation and subsequent PARP cleavage. RKO cells were transfected with a pCEP4 empty vector (RKO-φ) or the same vector containing cDNA encoding human Bcl-2 (RKO-Bcl2). RKO-φ or RKO-Bcl2 cells were treated with 60 µM HNE for the indicated times. Total cell lysates (50 µg/lane) were analyzed for the cleaved bands of PARP and pro-caspase-3, and the expression of Bcl-2 protein by Western blotting. The loading of protein on the SDS-PAGE gel was evaluated by staining the membrane with ink. Results are representative of three independent experiments.

results suggest that Bcl-2 prevents cells from undergoing apoptosis by inhibiting caspase-3 activation and subsequent PARP cleavage. Overexpression of Bcl-2 Protein Blocks Caspase-2 but Not Caspase-8 Activation Induced by HNE. Recent studies have shown that caspase-2, -6, -8, and -10 are downstream targets of caspase-3 and can be activated upon caspase-3 activation in a cell-free system in the presence of cytochrome c (27). We tested if caspase-2 and -8 are cleaved after caspase-3 activation in RKO-φ cells and if so, whether cleavage is blocked in RKO-Bcl-2 cells. Following treatment of cells with various concentrations

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Figure 5. Bcl-2 blocks HNE-induced caspase-2 but not caspase-8 activation. RKO-φ or RKO-Bcl2 cells were treated with indicated concentrations of HNE for 24 h. Total cell lysates (50 µg/lane) were analyzed for the cleaved bands of PARP, the active form p20 of caspase-2, and the expression of procaspase-2 and -8 by Western blotting. Results are representative of two independent experiments.

of HNE for 24 h, total cellular proteins were analyzed for PARP cleavage and caspase-2 and -8 activation by immunoblot analysis. In RKO-φ cells, PARP cleavage occurred with 45 µM HNE and reached a maximum at 75 µM. This paralleled the conversion of the inactive 50 kDa pro-caspase-2 into the active 20 kDa caspase. In contrast, PARP was not significantly cleaved in the RKOBcl-2 cells, and this observation was consistent with the absence of the active 20 kDa caspase-2 (Figure 5). Interestingly, in RKO-φ cells, PARP cleavage paralleled decreases in the levels of the 55 kDa procaspase-8, but cleavage of the latter was not inhibited by overexpression of Bcl-2. Cells were treated with 60 µM HNE and incubated for various periods then assayed for PARP cleavage and caspase-2 and -8 activation. Treatment of RKO-φ cells with HNE resulted in caspase-2 and -8 activation in a time-dependent manner (data not shown). Similarly, overexpression of Bcl-2 diminished caspase-2 but not caspase-8 activation. These results suggest that HNE-induced caspase-2 and -8 activation in RKO cells occurs by different mechanisms and only the former is Bcl-2 inhibitable. Overexpression of Bcl-2 Blocks Cytochrome c Translocation and Subsequent Caspase-9 Activation by HNE. Caspase-3 activation is a consequence of proteolysis by the active 37 kDa caspase-9 (16, 25). The activation of caspase-9 is promoted by cytochrome c in the presence of apoptotic protease activating factor-1 and dATP (15, 16). Cytochrome c is normally located on the outer surface of the inner mitochondrial membrane and may be released from the membrane to the cytosol following treatment of cells with an apoptotic stimulus (14). Overexpression of Bcl-2 may prevent the release of cytochrome c and subsequent caspase-9 activation (15). To test this possibility, we evaluated the levels of cytochrome c in cytosol and mitochondria and probed for the presence of the active 37 kDa caspase-9 in both RKO-φ and RKO-Bcl-2 cells following HNE treatment. In RKO-φ cells, the levels of cytochrome c in mitochondria decreased as the HNE doses increased, whereas the levels of cytochrome c in the cytosol increased. This

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Figure 6. Overexpression of Bcl-2 protein blocks the release of cytochrome c from mitochondria to cytosol and subsequent caspase-9 activation. RKO-φ or RKO-Bcl2 cells were treated with the indicated concentrations of HNE for 24 h. Cytosol and mitochondria were isolated from one-half of the cells as described in the Materials and Methods. The total cell lysates were isolated from the remaining cells. The cytosolic proteins (50 µg/ lane) and the mitochondrial proteins (20 µg/lane) were analyzed for cytochrome c content and the total cell lysates (50 µg/lane) were analyzed for the cleavage of pro-caspase-9 by Western blotting. The loading of protein on the SDS-PAGE gel was evaluated by staining the membrane with ink. Results are representative of three independent experiments.

coincided with the conversion of the inactive 48 kDa procaspase-9 into the active 37 kDa caspase-9 (Figure 6). However, in Bcl-2-expressing cells, the levels of cytochrome c in mitochondria were only slightly altered by treatment with HNE. Interestingly, the levels of cytochrome c in mitochondria were higher in RKO-φ cells than in RKO-Bcl-2 cells. In addition, the release of cytochrome c from mitochondria was substantially inhibited in Bcl-2 cells. Consequently, caspase-9 activation was blocked (Figure 6). These findings demonstrate that HNE triggers apoptotic cell death through promotion of cytochrome c release and activation of caspase cascades. Bcl-2 prevention of apoptosis results from inhibition of cytochrome c release and subsequent caspase activation.

Discussion Our experimental data demonstrate that the mitochondrion-dependent apoptotic mechanism plays a major role in HNE-induced cell death in RKO cells. In response to a variety of apoptosis-inducing agents, cytochrome c is released from mitochondria to cytosol (31, 32). Cytochrome c initiates apoptosis by inducing the formation of the apoptotic protease activating factor-1/caspase-9 complex in the presence of dATP (17). This processing leads to the cleavage of caspase-9, converting it to an active protease. Active caspase-9 then cleaves and activates caspase-3 leading to DNA fragmentation and cell death (14, 15). We observed similar apoptotic processes in cells undergoing HNE-induced apoptosis. The release of cytochrome c into cytosol may provide a potential target for regulation of HNE-induced apoptosis.

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Cytochrome c is bound to the outer surface of the inner mitochondrial membrane, whereas Bcl-2 is localized mainly on the outer mitochondrial membrane (33). Thus, a natural connection exists between cytochrome c and Bcl-2. In fact, recent studies have indicated that Bcl-2 expression affects cation transport in mitochondria and protects against organelle dysfunction induced by several apoptotic stimuli (31-33). In cells overexpressing Bcl-2, or the closely related protein, Bcl-xL, cytochrome c release and the subsequent apoptotic response are blocked (33). Our results are consistent with these findings and demonstrate that HNE-induced caspase activation and subsequent PARP cleavage are suppressed in Bcl-2 transfected RKO cells. This suppression parallels the inhibition of cytochrome c release from mitochondria indicating that mitochondrial integrity and function are protected by overexpression of Bcl-2. The mechanism of HNE-induced cytochrome c release is undefined. One possibility is that depletion of cellular glutathione (GSH), a critical component of the cellular defense system against oxidative stress (34), plays a role in HNE-induced cytochrome c release. Depletion of GSH has been considered to be an early event in the apoptotic process, and may increase sensitivity to pro-apoptotic stimuli (9, 10). Since HNE readily reacts with sulfhydryl groups of proteins (34) and its detoxification is via conjugation with GSH (34, 35), attention has been paid to its role in oxidative damage, especially on its effect on GSH concentrations. It has recently reported that buthionine sulfoximine (BSO)-induced GSH depletion is associated with a decrease in Bcl-2 protein levels and an increase in apoptosis in cholangiocytes (9). Pretreatment of cells with glutathione ethyl ester prevented BSOinduced GSH depletion, Bcl-2 degradation, and apoptosis (9). Bcl-2 expression results in an elevation of the basal levels of hydrogen peroxide but dampens the excessive production of hydrogen peroxide induced by apoptotic stimuli (36). PC12 cells overexpressing Bcl-2 exhibit higher levels of GSH and lower levels of HNE after oxidative stress (8). HNE-induced apoptosis in human T lymphoma Jurkat cells occurs through depletion of intracellular GSH and activation of the caspase cascade (10). Our experiments have shown that treatment of RKO cells with HNE-induced GSH depletion within 15 min, triggered a maximal level of depletion at 1-2 h and gradually recovered thereafter. These observations suggest that apoptosis is associated with the depletion of GSH levels in RKO cells treated with HNEs. Hydroxyalkenals were discovered as toxic products present in extracts of oxidized lipid (37, 38). Indeed, hydroxyalkenals and their cysteine-conjugates demonstrate in vivo antitumor activity (39). The toxicity of HNE at low concentration is due to its ability to apoptosis. We demonstrate here that a key event in this process is the HNE-induced release of cytochrome c. Thus, alteration of mitochondrial function is critical to the induction of apoptosis by a major product of lipid peroxidation. This may contribute to the induction of apoptosis by a broad range of agents that induce oxidative stress in tumor cells, neurons, and other cell types.

Acknowledgment. We thank Kevin Kozak for his kind assistance in preparation of the manuscript. This work was supported by research Grants CA47479 (L.J.M.) and CA70856 (J.A.P.) and center grants (CA68485 and ES00267) from the National Institutes of Health.

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