Endogenously Generated Hydrogen Peroxide Is Required for

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Endogenously Generated Hydrogen Peroxide Is Required for Execution of Melphalan-Induced Apoptosis as Well as Oxidation and Externalization of Phosphatidylserine Tatsuya Matsura,*,† Masachika Kai,† Jianfei Jiang,‡ Hareesh Babu,‡ Vidisha Kini,‡ Chiaki Kusumoto,† Kazuo Yamada,† and Valerian E. Kagan*,‡ Division of Medical Biochemistry, Department of Pathophysiological and Therapeutic Science, Faculty of Medicine, Tottori University, 86 Nishi-cho, Yonago 683-8503, Japan, and Department of Environmental and Occupational Health, University of Pittsburgh, 3343 Forbes Avenue, Pittsburgh, Pennsylvania 15260 Received October 15, 2003

Hydrogen peroxide (H2O2) is generated endogenously during execution of both intrinsic as well as extrinsic apoptotic programs suggesting that it may function as a secondary messenger in apoptotic pathways. In the present study, we investigated the role of endogenously generated H2O2 by using two cell linessHL-60 cells and its subclone, H2O2 resistant HP100 cells overexpressing catalase (CAT). With the exception of CAT, we found no differences in the expression of other primary antioxidant enzymes (Cu/Zn-superoxide dismutase, Mn-superoxide dismutase, and glutathione peroxidase) or apoptosis-related proteins (Bcl-2 and Bax) in HP100 cells as compared with the parental HL-60 cells. Production of H2O2 was readily detectable as early as 1 h after melphalan (Mel) exposure of HL-60 cells but not HP-100 cells. Biomarkers of apoptosis, such as release of cytochrome c, disruption of mitochondrial transmembrane potential, caspase-3 activation, and chromatin condensation, became apparent much later, 3 h and onward after Mel treatment of HL-60 cells. The emergence of essentially all biomarkers of apoptosis was dramatically delayed in HP100 cells as compared with HL-60 cells. A relatively minor phospholipid species, phosphatidylserine (PS), was markedly oxidized 3 h after Mel treatment in HL-60 cells (but not in HP100 cells) where it was significantly inhibited by exogenously added CAT. The two most abundant classes of membrane phospholipids, phosphatidylcholine and phosphatidyletanolamine, did not undergo any significant oxidation. PS oxidation took place 3 h after exposure of HL-60 cells to Mel and paralleled the appearance of cytochrome c in the cytosol. Neither cytochrome c release nor PS oxidation occurred in Meltreated HP100 cells, indicating that both endogenous H2O2 and cytochrome c were essential for selective PS oxidation detected in HL-60 cells. Mel-induced PS oxidation was also associated with externalization of PS on the surface of HL-60 cells. Given that 3-amino-1,2,4-triazole, a CAT inhibitor, suppressed the resistance of HP100 cells to apoptosis, production of reactive oxygen species, PS oxidation, and PS externalization induced by Mel, the results from the present study suggest that H2O2 is critical for triggering the Mel-induced apoptotic program as well as PS oxidation and externalization.

Introduction It became a banality that exogenously generated ROS1 are potent inducers of cell damage and apoptosis (1-4). More importantly, ROS have been identified as intermediates formed endogenously during execution of both intrinsic as well as extrinsic apoptotic programs triggered by a variety of stimuli (5-7). Thus, these observations imply that a potential signaling role for ROS in apoptotic pathways may exist (reviewed in 7 and 8). Several reports indicate that H2O2 may fulfill a messenger function by interacting with various physiological stimuli including * To whom correspondence should be addressed. (T.M.) Tel: +81-859-34-8010. Fax: +81-859-34-8077. E-mail: tmatsura@ grape.med.tottori-u.ac.jp. (V.E.K.) Tel: +1-412-383-2136. Fax: +1-412383-2123. E-mail: [email protected]. † Tottori University. ‡ University of Pittsburgh.

cytokines, growth factors, or transforming factors (9-13) and serve as a modulator of endothelial cell proliferation (14). However, specific role(s) for H2O2 in apoptosis remains to be established. We have reported that selective oxidation of PS facilitates its externalization on the surface of the plasma membrane during apoptosis and that oxidized PS acts 1 Abbreviations: ROS, reactive oxygen species; H O , hydrogen 2 2 peroxide; PS, phosphatidylserine; PC, phosphatidylcholine; CAT, catalase; Mel, melphalan; 3-AT, 3-amino-1,2,4-triazole; CCCP, carbonyl cyanide m-chlorophenyl-hydrazone; FCS, fetal calf serum; AMC, 7-amino-4-methyl-coumarin; DCFH2-DA, 2′,7′-dichlorodihydrofluorescein diacetate; CMXRos, chloromethyl-X-rosamine; PnA, cis-parinaric acid; PMSF, phenylmethylsulfonyl fluoride; GPX, glutathione peroxidase; SOD, superoxide dismutase; PVDF, polyvinylidine difluoride; RTPCR, reverse transcriptase-polymerase chain reaction; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; DCFH2, 2′,7′-dichlorodihydrofluorescein; DCF, 2′,7′-dichlorofluorescein; FITC, fluorescein isothiocyanate; ∆Ψm, mitochondrial transmembrane potential; PE, phosphatidylethanolamine; O2-, superoxide; NF-kB, nuclear factor-kB.

10.1021/tx030050s CCC: $27.50 © 2004 American Chemical Society Published on Web 04/28/2004

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as an important signal contributing to recognition of apoptotic cells by macrophages (4, 15-18). We further established that antioxidant enzymes and low molecular weight antioxidants capable of blocking PS oxidation inhibited these PS-dependent signaling pathways (4, 18, 19). These effects were specific to PS oxidation as other oxidized phospholipids (e.g., PC) were neither selectively oxidized nor recognized by macrophages (4, 18, 19). We have put forward a hypothesis that selectivity of PS oxidation is achieved through specific catalytic oxidation of negatively charged PS on the cytosolic side of plasma membrane by positively charged cytochrome c released into the cytosol from mitochondria during apoptosis (19, 20). However, the source of oxidizing equivalents required for redox catalysis of PS oxidation has not been identified. Disruption of mitochondrial electron transport and subsequent massive production of ROS (especially H2O2) is likely a candidate for activation of cytochrome c catalytic properties. Yet, experimental evidence for their role in oxidative signaling in apoptosis is lacking. Intracellularly generated H2O2 can diffuse from cells; hence, it becomes available for extracellular CAT, one of the major H2O2-decomposing enzymes, that is not readily internalized by cells. Therefore, employment of exogenous CAT for accurate characterization of the site of PS oxidation (inner or outer leaflet of plasma membrane) as it relates to endogenous sources of ROS production can hardly provide any valuable information. One approach to address the issue is to study apoptotic mechanisms in two comparable cell lines differing in their handling of H2O2. To this end, we chose to use two cell lines, HL-60 cells and its subclone, H2O2 resistant HP100 cells developed by repeated exposure of HL-60 cells to 100 µM H2O2 (21). The resistance of HP100 cells to H2O2 has been characterized as being mostly due to overexpression of CAT (22-24). Because major drawbacks associated with limited intracellular bioavailability of exogenously added CAT could be overcome in HP100 cells, we opted to use them along with HL-60 cells in comparative studies of a messenger role for H2O2 in apoptotic signaling. In the current study, we employed an alkylating agent, Mel, as a nonoxidant proapoptotic stimulus and found that Melinduced H2O2 indeed plays a pivotal role in implementation of apoptosis as a required messenger.

Experimental Procedures Chemicals. Mel, CAT, SDS, 3-AT, CCCP, and GSH were purchased from Sigma Chemical Co. (St. Louis, MO). H2O2 was purchased from Mitsubishi Gas Chemical Company Inc. (Tokyo, Japan). A 123 bp DNA ladder was purchased from Invitrogen Corporation (Carlsbad, CA). Me2SO was purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). FCS was purchased from PAA Laboratories GmbH (Linz, Austria). HPLC solvents (methanol, chloroform, and hexane) were obtained from Aldrich Chemical Co. (Milwaukee, WI). AMC and acetyl-AspGlu-Val-Asp-AMC (Ac-DEVD-AMC) were purchased from Peptide Institute, Inc. (Osaka, Japan). ThioGlo-1 maleimide reagent was obtained from Calbiochem-Novabiochem Corporation (San Diego, CA). RPMI 1640 medium without phenol red was obtained from Cosmo Bio Co., Ltd. (Tokyo, Japan). RPMI 1640 medium with phenol red was obtained from Nissui Pharmaceutical Co., Ltd. (Tokyo, Japan). Hoechst 33342, DCFH2-DA, CMXRos, and PnA were purchased from Molecular Probes (Eugene, OR). The purity of PnA was determined by UV spectrophotometry at 304 nm in ethanol ( ) 80 mM-1 cm-1). All other chemicals used were of analytical grade. Cell Cultures. The human promyelocytic leukemia HL-60 cell line and HP100 cell line, an HL-60-derived H2O2 resistant

Matsura et al. cell line were obtained from Riken Cell Bank (Tsukuba, Japan). Both cell lines were grown in RPMI 1640 medium supplemented with 10% heat-inactivated FCS, 2 mM glutamine, and antibiotics (100 units/mL penicillin and 100 µg/mL streptomycin) at 37 °C in a humidified incubator with 5% CO2. All of the experiments were performed using RPMI 1640 medium containing 10% FCS unless specified otherwise. Cells were seeded at a concentration of 2 × 105 cells/mL, and logarithmic growth was maintained by passaging every 2 or 3 days. Assay of CAT Activity. HL-60 cells and HP100 cells were harvested and washed with PBS twice. In some experiments, HP100 cells were treated with different concentrations of 3-AT, a specific inhibitor of CAT, for 1 or 2 h. Cells were resuspended in 50 mM Tris-HCl (pH 7.5), 1 mM EDTA, 0.5 mM DTT, 1 mM PMSF, 0.01% digitonin, and 0.25% sodium cholate. Samples were incubated for 30 min on ice and centrifuged at 12 000g for 15 min at 4 °C. CAT activity in the supernatant was determined by measuring the rate of H2O2 decomposition (25). The supernatant (20 µL) was added to 1.0 mL of 50 mM phosphate buffer (pH 7.0) containing 10 mM H2O2. The decrease in H2O2 absorbance at 240 nm was recorded against a blank containing no substrate at 20 °C and calculated using an extinction coefficient of 0.04 mM-1 cm-1. One unit of CAT activity was defined as the amount of enzyme required to decompose 1 µmol of H2O2 per minute at 20 °C. The protein concentration of samples was measured by the method of Bradford (26). Western Blot Analysis. Western blot analyses of CAT, GPX, Cu/Zn-SOD, Mn-SOD, Bcl-2, and Bax in HL-60 cells and HP100 cells were performed as previously described (3). Briefly, protein was extracted from cells with lysis buffer containing 50 mM Tris-HCl, pH 7.5, 5 mM EDTA, 150 mM NaCl, 0.5% NP40, and 1 mM PMSF. The protein content of each sample was quantified by the Bradford method (26). After the same volume of 2 × SDS sample buffer containing 125 mM Tris/HCl, pH 6.8, 20% glycerol, 4% SDS, 12% 2-mercaptoethanol, and 0.006% bromophenol blue was added, samples were incubated at 95 °C for 15 min to denature the proteins. The appropriate protein amount (10-30 µg) was separated by SDS-PAGE using 10 (for CAT and SOD) or 12% (for GPX, Bcl-2 and Bax) polyacrylamide gel and electroblotted onto a PVDF membrane (Millipore, Billerica, MA). After nonspecific binding sites were blocked with washing buffer containing 100 mM Tris/HCl, pH 7.5, 154 mM NaCl, and 0.05% Tween 20 supplemented with 5% skim milk, the PVDF membrane was incubated for 1 h at room temperature with rabbit anti-human CAT antibody (Calbiochem) (1:1000), anti-human GPX monoclonal antibody (Medical and Biological Laboratories Co., Ltd., Nagoya, Japan) (1:100), goat anti-human Cu/Zn-SOD antibody (Austral Biologicals, San Ramon, CA) (1: 1000), anti-human Mn-SOD monoclonal antibody (Chemicon International, Temecula, CA) (1:1000), anti-rat Bcl-2 monoclonal antibody (Medical and Biological Laboratories Co., Ltd.) (1:1000) or anti-human Bax monoclonal antibody (Medical and Biological Laboratories Co., Ltd.) (1:1000). It was then washed with washing buffer, incubated further with horseradish peroxidaseconjugated secondary IgG antibodies for 1 h at room temperature, and rewashed with washing buffer. The immunoblot was revealed using an enhanced chemiluminescence detection kit (Amersham Biosciences Corp., Piscataway, NJ). The densitometric quantitative analysis was performed using Multi-Analyst 1.1 computer software (Hercules, CA). RNA Preparation and RT-PCR. RT-PCR was performed to measure the level of CAT mRNA in HL-60 and HP100 cells. Total RNA was isolated from the cells using the acid guanidiumphenol-chloroform method. As a template for PCR, singlestranded cDNA was prepared from 0.6 µg of total RNA by MMLV reverse transcriptase (Invitrogen Corporation) and PCR was carried out with Taq DNA polymerase AmpliTaq Gold (Applied Biosystems, Foster City, CA) using a thermal circular Techne Genius [Techne (Cambridge) Ltd., Cambridge, U.K.]. The RT reaction was performed at 50 °C for 30 min. For PCR,

H2O2 Mediates Melphalan-Induced Apoptosis two sets of specific primers were designed from human CAT and human GAPDH cDNAs. PCR conditions were as follows: 21 cycles of denaturation at 94 °C for 1 min, annealing at 50 °C for 1 min, and extension at 72 °C for 1 min, followed by cooling to 4 °C. PCR products were electrophoresed on a 2% agarose gel, visualized under UV light after staining with ethidium bromide, and photographed. PCR primer sequences were as follows. CAT: sense, 5′GCCTGGGACCCAATTATCTT-3′; antisense, 5′-GAATCTCCGCACTTCTCCAG-3′. GAPDH: sense, 5′-CGACCACTTTGTCAAGCTCA-3′; antisense, 5′-AGGGGAGATTCAGTGTGGTG-3′. Detection of DNA Laddering. DNA ladder formation was detected as previously described (3). Briefly, 4 h after treatment with various concentrations of H2O2, HL-60 cells and HP100 cells were centrifuged, washed, pelleted, and incubated in lysis buffer containing 10 mM EDTA, 50 mM Tris/HCl, pH 8.0, 0.5 mg/mL proteinase K, and 0.5% SDS for 1 h at 50 °C. Then, the lysates were extracted with phenol, followed by chloroform, and precipitated by the addition of 1/10 volume of 3 M sodium acetate and 2 volumes of ethanol. The pellets were rinsed with 70% ethanol, air-dried, dissolved in TE buffer (pH 8.0) containing 10 mM Tris/HCl and 1 mM EDTA, and treated with DNasefree RNase for 2 h at 37 °C. After re-extraction and reprecipitation, samples were dissolved in TE buffer. Approximately 10 µg of DNA in each well was electrophoresed on a 2% agarose gel, visualized under UV light after staining with ethidium bromide, and photographed. Determination of Apoptotic Cells by Nuclear Morphology. Apoptotic cells were assessed by nuclear morphology as described previously (15). Cells were incubated with Mel at 37 °C, washed with PBS, and then fixed with 1% glutaraldehyde overnight. Samples were centrifuged, resuspended in PBS, stained with Hoechst 33342 (1 mM), and then mounted on a glass slide and observed under fluorescence microscope. Results were expressed as the percentage of the cells showing characteristic nuclear morphological features of apoptosis (nuclear condensation and fragmentation) relative to the total number of counted cells (g200 cells per each point). Determination of Caspase-3 Activity. The activity of caspase-3 was determined as described previously (3, 15, 27). Briefly, after treatment of HL-60 and HP100 cells (1 × 106 cells/ mL) with Mel, cells were collected, washed in PBS, and lysed for 20 min on ice in lysis buffer containing 10 mM HEPESKOH (pH 7.4), 2 mM EDTA, 0.1% CHAPS, 1 mM PMSF, and 5 mM DTT. The suspensions were centrifuged at 4 °C, and the supernatants were collected as lysates. For measurement of caspase activity, 10 µg of lysate diluted to 20 µL with lysis buffer was mixed with 20 µL 2 × ICE buffer containing 40 mM HEPES-KOH (pH 7.4), 20% (v/v) glycerol, 1 mM PMSF, and 4 mM DTT supplemented with 40 µM Ac-DEVD-AMC (a fluorogenic peptide substrate) and incubated for 60 min at 37 °C. After 60 min, 460 µL of distilled water was added and the fluorescence was measured in a CytoFluor II (Applied Biosystems) fluorescence microplate reader using excitation at 360 ( 40 nm and emission at 460 ( 40 nm. One unit of caspase activity was defined as the amount of enzyme required to release 1 pmol AMC per minute at 37 °C. The protein concentration of cell lysates was measured by the method of Bradford (26). Flow Cytometric Measurement of ROS Production with DCFH2-DA. To measure the production of ROS, cells were treated with 40 µg/mL Mel in RPMI 1640 without phenol red at 37 °C and incubated with 10 µM DCFH2-DA for the last 30 min of the treatment in the dark. Activity of cellular esterases cleaves DCFH2-DA into DCFH2. The DCF fluorescence resulting from the oxidation of DCFH2 was measured in 10 000 cells by using Epics cytofluorometer (Beckman Coulter, Inc., Fullerton, CA). Determination of Phospholipid Peroxidation in HL-60 Cells. PnA is a natural 18-carbon fatty acid with four conjugated double bonds. PnA is highly susceptible to peroxidation and also fluorescent. Fluorescence is irreversibly lost upon peroxidation thus providing a convenient approach to quantitate lipid per-

Chem. Res. Toxicol., Vol. 17, No. 5, 2004 687 oxidation (28, 29). PnA was metabolically incorporated into HL60 and HP100 cell phospholipids (1 × 106 cells/mL) by addition of PnA-human serum albumin complex to give a final concentration of 2.5 µg PnA/106 cells in FCS-free RPMI 1640 medium without phenol red. Cells were incubated for 2 h at 37 °C in the dark. Thereafter, PnA-labeled cells were resuspended in RPMI 1640 supplemented with 10% FCS and treated with Mel at 37 °C for indicated time periods. At the end of the incubation period, phospholipid oxidation was determined by fluorescence HPLC according to previously described methods (30). Determination of Intracellular GSH Contents. HL-60 cells and HP100 cells (1 × 106 cells/mL) were incubated in the absence or presence of Mel in RPMI 1640 medium for indicated time periods at 37 °C in 5% CO2. After treatments, cells were harvested and GSH content in the cells was determined fluorometrically using ThioGlo-1 as previously described (15). Briefly, the cells treated with Mel were harvested, washed with PBS, resuspended in PBS, and lysed by freezing and thawing once. Immediately after addition of 10 µM ThioGlo-1 to the cell lysates, fluorescence was measured in a CytoFluor II (Applied Biosystems) fluorescence microplate reader using excitation at 360 ( 40 nm and emission at 530 ( 25 nm. The protein concentration of cell lysates was measured by the method of Bradford (26). Flow Cytometry of PS Externalization. Annexin V binding to cells was determined using a commercially available Annexin V-FITC Apoptosis Detection Kit (Biovision Inc., Mountain View, CA) and flow cytometry as previously described (15). Briefly, after treatment of cells (1 × 106 cells/mL) with Mel, cells were recovered and washed once with PBS. Cells were incubated with FITC-conjugated annexin V (0.5 µg/mL) for 15 min and then were collected by centrifugation and washed with binding buffer. Propidium iodide (0.6 µg/mL) was added, and cells were immediately analyzed with a FACScan flow cytometer (Becton Dickinson, San Jose, CA) with simultaneous monitoring of green fluorescence (530 nm, 30 nm band-pass filter) for annexin V-FITC and red fluorescence (long-pass emission filter that transmits light >650 nm) associated with propidium iodide. Mitochondrial Transmembrane Potential Assay. The ∆Ψm was determined by the flow cytometric analysis of CMXRos-stained cells as previously described (31). CMXRos (C32H32Cl2N2O) with a thiol reactive chloromethyl moiety is a novel mitochondrion selective dye that is well-retained during cell fixation. Once the dye accumulates in the mitochondria, it can react with accessible thiol groups on peptides and proteins to form an aldehyde fixable conjugate. Cells (1 × 106 cells) were loaded with 250 nM CMXRos for the last 30 min of treatment with Mel, washed in PBS, fixed with 1% glutaraldehyde, and analyzed on an Epics cytofluorometer (Beckman Coulter, Inc.). The cells exposed to 50 µM CCCP for 1 h were used as a positive control of decrease in ∆Ψm. Measurement of Cytochrome c Release. HL-60 cells and HP100 cells were treated with 40 µg/mL Mel, harvested, and washed three times with ice-cold PBS. Cells were resuspended in isotonic buffer containing 10 mM HEPES, 0.3 M mannitol, and 0.1% bovine serum albumin, supplemented with 0.1 mM digitonin (1.5 × 106 cells/mL), left on ice for 5 min, and immediately centrifuged at 8500g for 5 min at 4 °C. The supernatant collected was used as the cytosolic fraction. The cytochrome c amount in the cytosolic fraction was measured by Western blotting using anti-human cytochrome c monoclonal antibody (BD Biosciences Pharmingen, San Diego, CA) (1:500) as described above. Statistical Evaluations. Data are expressed as means ( SE. Changes in variables for different assays were analyzed either by Student’s t-test (single comparisons) or by one way ANOVA for multiple comparisons. Differences were considered to be significant at p < 0.05.

Results Overexpression of CAT Is Responsible for Increased Resistance of HP100 Cells to Mel-Induced

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Apoptosis. 1. Comparison of CAT Activity, CAT Protein Expression, and CAT Gene Expression in HL-60 Cells and HP100 Cells and Their Resistance to H2O2-Induced Apoptosis. To characterize CAT expression in HP100 cells, we examined cellular CAT activity and CAT mRNA and protein content in HL-60 cells and HP100 cells. CAT activity was five times higher in HP100 cells than in parent HL-60 cells (Figure 1A). Western blot analysis displayed that CAT protein content was 2.3 times greater in HP100 cells than in HL-60 cells, whereas there were no differences between the two cell types in the contents of three other antioxidant enzymes, GPX, Cu/Zn-SOD, and Mn-SOD (Figure 1B). This is in line with previously reported data that there was no difference in GPX activity between HL-60 cells and HP100 cells (23). It has been reported that the level of Bcl-2 increased after H2O2 exposure leading to resistance to apoptosis (32). Therefore, we also measured an antiapoptotic protein Bcl-2 and a proapoptotic protein Bax in HL-60 cells and HP100 cells. However, two cell lines had the same amount of these Bcl-2 family proteins (Figure 1B). The CAT mRNA level in HL-60 cells and HP100 cells was determined by RT-PCR. Quantitative analysis of the blots on electrophoretic gels (CAT mRNA/ GAPDH mRNA) revealed that HP100 cells contained approximately two times more CAT mRNA than HL-60 cells (data not shown). We next examined whether HP100 cells actually exerted a higher resistance to apoptosis induced by exogenous H2O2 as evidenced by DNA ladder formation. We found that at concentrations g50 µM, H2O2 induced DNA laddering in HL-60 cells after 4 h of incubation, while HP100 cells remained resistant to DNA ladder formation even at H2O2 concentrations as high as 5 mM (Figure 1C). GPX and CAT are the two major enzymes responsible for control and maintenance of nontoxic levels of H2O2 in cells. Given that we did not find any differences in GPX levels as well as in the contents of Bcl-2 and Bax between HL-60 cells and HP100 cells, it is likely that a higher expression of CAT in the latter was responsible for their increased resistance to DNA laddering (apoptosis) induced by exogenously added H2O2. These results laid the ground for further comparative studies using these two cell lines to establish the role of endogenously formed H2O2 (ROS) in nonoxidant-induced apoptosis. 2. Resistance of HP100 Cells to Apoptosis Induced by Various Concentrations of Mel. We chose to utilize an alkylating agent, Mel, as a proapoptotic stimulus because it (i) triggers intrinsic (mitochondriadependent) apoptotic mechanisms that are likely to involve H2O2 production (33, 34) and (ii) does not contain chemical moieties that may be accountable for direct radical scavenging effects. To compare the sensitivity of HL-60 cells and HP100 cells to Mel-induced apoptosis, we treated both cell lines with Mel at concentrations ranging from 10 to 40 µg/mL for 4 h. A robust and dosedependent increase in the number of apoptotic cells (Figure 2A) and caspase-3 activity (Figure 2B) was detected in HL-60 cells after treatment with Mel at concentrations g20 µg/mL. In contrast, HP100 cells feebly responded to much higher concentrations of Mel (g30 µg/mL). At all doses of Mel tested, both the number of apoptotic cells and the caspase-3 activity were manyfold higher in HL-60 cells than in HP100 cells (Figure 2A,B).

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Figure 1. Comparison of CAT activities, antioxidant enzyme protein levels, Bcl-2 and Bax protein expression, and H2O2induced DNA fragmentation in HL-60 cells and HP100 cells. (A) Cellular CAT activities were measured spectrophotometrically as described in the Experimental Procedures. Data points represent the mean ( SE of 14-20 separate experiments. Note: some error bars are too small to visualize. (B) Western blot analysis of antioxidant enzymes, Bax, and Bcl-2 contents in cells. Protein contents of CAT, GPX, SOD, Bax, and Bcl-2 were determined by immunoblotting as described in the Experimental Procedures. The results of the densitometric analysis were shown as HP100/HL-60. Data points represent the mean ( SE of four separate experiments. **p < 0.01 vs all other protein expressions. (C) Internucleosomal DNA fragmentation in cells exposed to various concentrations of H2O2. Cells (5 × 106) were seeded into dishes and incubated with the indicated concentrations of H2O2 to determine the resistance to H2O2 for 4 h at 37 °C under 5% CO2 and 95% air. Cellular DNA was extracted, and 10 µg aliquots were electrophoresed on a 2% agarose gel. M is the 123 bp DNA ladder as a marker. Representative data from three separate experiments are shown.

H2O2 Mediates Melphalan-Induced Apoptosis

Figure 2. Changes in apoptotic biomarkers in HL-60 cells (closed circles) and HP100 cells (open circles) exposed to Mel. Dose response (A and B). Cells were incubated with the indicated concentration of Mel in 10% FCS containing RPMI 1640 for 4 h at 37 °C under a 5% CO2 and 95% air atmosphere. Time course (C and D). Cells were incubated with 40 µg/mL Mel in 10% FCS containing RPMI 1640 for the indicated time period at 37 °C under a 5% CO2 and 95% air atmosphere. (A and C) The number of apoptotic cells was determined by examining nuclear morphology after Hoechst 33342 staining. (B and D) Caspase-3 activities were measured using a fluorescent substrate as described in the Experimental Procedures. Data points represent the mean ( SE of 3-8 separate experiments. Note: some error bars are too small to visualize. ap < 0.01, bp < 0.05 vs HL-60 cells at the indicated concentration of Mel. cp < 0.01, dp < 0.05 vs nontreated control cells. ep < 0.01, fp < 0.05 vs HL-60 cells at the indicated time. gp < 0.01 vs value at 0 h in each cell line.

3. Comparison of the Time Course of Mel-Induced Apoptosis in HL-60 Cells and HP100 Cells. We next examined the time course of changes in apoptotic cell number and caspase-3 activity in HL-60 cells and HP100 cells following Mel treatment (40 µg/mL). Apoptotic cells (Figure 2C) and caspase-3 activation (Figure 2D) in HL60 cells became evident after 3 h of Mel treatment, while these levels of caspase-3 activity and number of apoptotic cells were achieved after 4 h of exposure of HP100 cells to Mel. After 4 h of treatment with Mel, the increase in the number of apoptotic cells was four times greater and activation of caspase-3 was three times greater in HL60 cells than in HP100 cells. 4. Effect of Mel on CAT Activity in HL-60 Cells and HP100 Cells. To test whether the difference in resistance of HP100 cells and HL-60 cells to Mel-induced apoptosis could be accounted for by the effects of Mel on CAT, we determined the effect of Mel on CAT activity in both cell lines. Cells were treated with 40 µg/mL Mel for 4 h and harvested, and CAT activity was measured spectrophotometrically. There was no difference in CAT activity in both cell lines before and after incubation with

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Mel (data not shown). 5. Effect of a CAT Inhibitor, 3-AT, on CAT Activity in HP100 Cells. If overexpression of CAT in HP100 cells was mainly responsible for their resistance to Melinduced apoptosis, then suppression of CAT activity to the level found in HL-60 cells should result in the increased Mel sensitivity comparable to that in HL-60 cells. To manipulate CAT activity in HP100 cells, we used different concentrations of a CAT inhibitor, 3-AT. CAT activity markedly declined at both 1 and 2 h incubations of HP100 cells in the presence of 25, 50, and 100 mM 3-AT without affecting cell viability (Figure 3A). Treatment with 50 or 100 mM 3-AT for 1 h yielded HP100 cells with approximately the same CAT activity as that in untreated HL-60 cells. Even a more pronounced decrease of CAT activity was observed after 2 h of incubation of HP100 cells with 3-AT. 6. Effect of 3-AT on Mel-Induced Apoptosis and Caspase-3 Activity in HP100 Cells. We further tested the effect of CAT inhibition on Mel-induced apoptosis in HP100 cells. HP100 cells were treated with 50 or 100 mM 3-AT for 1 h followed by 40 µg/mL Mel for 4 h. As shown in Figure 3B,C, 3-AT-pretreated HP100 cells exerted an increased sensitivity to Mel. In HP100 cells treated with 50 and 100 mM 3-AT, both the number of apoptotic cells (Figure 3B) and the caspase-3 activity (Figure 3C) increased and reached the same levels as those detected in HL-60 cells exposed to Mel (compare with Figure 2C,D, respectively). These results indicate that elevated CAT activity in HP100 cells is predominantly responsible for their resistance to Mel-induced apoptosis. 7. Effect of Exogenously Added CAT on MelInduced Apoptosis and Caspase-3 Activity in HL60 Cells. We have previously reported that extracellularly added CAT can efficiently inhibit apoptosis induced by a variety of stimuli (4, 18). Therefore, to further support our conclusion that different sensitivity of HL60 cells and HP100 cells to Mel-induced apoptosis is largely due to their different CAT activities, we examined whether exogenous CAT could protect HL-60 cells against Mel-induced apoptosis. When HL-60 cells were incubated with 40 µg/mL Mel in the presence of 50 U/mL CAT, the number of apoptotic cells decreased by 24% (Figure 4A) and caspase-3 activity was inhibited by 35% (Figure 4B) as compared with HL-60 cells in the absence of CAT. Thus, exogenous CAT was able to partially prevent Melinduced apoptosis in HL-60 cells. Characterization of Mel-Induced Oxidative Stress in HL-60 Cells and HP100 Cells. Because increased resistance of HP100 cells to Mel-induced apoptosis as compared to HL-60 cells was clearly associated with the CAT overexpression in the former, we next determined whether differences in ROS production and oxidative stress during execution of apoptotic program could be detected between these respective cell lines. 1. ROS Generation in HL-60 Cells and HP100 Cells following Exposure to Mel. Using DCFH2-DA as a probe, we observed ROS production (as assessed by an increase in DCF-derived fluorescence) in HL-60 cells but not in HP100 cells as early as 1 h after treatment with 40 µg/mL Mel (Figure 5). Treatment of HL-60 cells with Mel for 2 and 3 h exhibited a similar degree of ROS generation as compared with 1 h treatment with Mel (data not shown). No increase in ROS generation was

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Figure 3. (A) Inhibitory effect of 3-AT on CAT activities in HP100 cells. Cells were incubated with the indicated concentrations of 3-AT in 10% FCS containing RPMI 1640 for 1 (white bars) or 2 h (black bars) at 37 °C under a 5% CO2 and 95% air atmosphere. CAT activities were determined spectrophotometrically as described in the Experimental Procedures. Data points represent the mean ( SE of three separate experiments. Note: some error bars are too small to visualize. **p < 0.01 vs 1 h treatment at the indicated concentration of 3-AT. (B and C) 3-AT enforces HP100 cells to undertake Mel-induced apoptosis. Cells were treated with various concentrations of 3-AT in 10% FCS containing RPMI 1640 for 1 h at 37 °C under a 5% CO2 and 95% air atmosphere. Thereafter, cells were washed three times in 10% FCS containing RPMI 1640 and incubated in the presence (black bars) or absence (white bars) of 40 µg/mL Mel for 4 h. The number of apoptotic cells (B) and caspase-3 activity (C) were determined as described in the Experimental Procedures. Data points represent the mean ( SE of 3-5 separate experiments. **p < 0.01, *p < 0.05 vs Mel-treated HP100 cells in the absence of 3-AT.

Figure 4. Exogenous CAT protects HL-60 cells against Melinduced apoptosis. Cells were incubated with 40 µg/mL Mel in 10% FCS containing RPMI 1640 in the presence (black bars) or absence (white bars) of 50 U/mL CAT for 4 h at 37 °C under a 5% CO2 and 95% air atmosphere. The number of apoptotic cells (A) and caspase-3 activity (B) were determined as described in the Experimental Procedures. Data points represent the mean ( SE of 3-5 separate experiments. **p < 0.01, *p < 0.05 vs HL-60 cells in the absence of CAT.

detected following Mel treatment when HL-60 cells were cotreated with exogenous CAT (Figure 5). Notably, DCF

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Figure 5. Mel induces ROS production in HL-60 cells but not HP100 cells. Cells were treated with 40 µg/mL Mel in RPMI 1640 without phenol red for 1 h at 37 °C and incubated with 10 µM DCFH-DA for the last 30 min of the treatment in the dark. In some experiments, cells were treated with Mel in the presence of 50 U/mL CAT or treated with 100 mM 3-AT for 2 h before Mel exposure. The results are representative of three separate experiments.

fluorescence emission from naı¨ve HP100 cells was approximately five times lower than that from naı¨ve HL60 cells (Figure 5), suggesting that only minimal amounts of ROS were able to interact with the fluorescent probe in CAT overexpressing HP100 cells under normal culture conditions. When HP100 cells were pretreated with 3-AT, Mel caused ROS production in HP100 cells as well (Figure 5). DCFH2-DA reacts not only with H2O2 but also with other ROS including peroxynitrite (35, 36). However, given the fact that both exogenous CAT and endogenously overexpressed CAT completely protected Meltriggered ROS production in HL-60 cells and 3-AT suppressed resistance of HP100 cells to Mel-induced ROS generation, it is likely that most of the endogenously generated ROS in our experimental conditions were represented by H2O2. Thus, Mel-induced levels of H2O2 available for interactions with reductants (such as DCFH2) were markedly lower in HP100 cells than in HL-60 cells. 2. Effect of Mel Exposure on GSH Content in HL60 Cells and HP100 Cells. As GSH is the major intracellular reductant, we next measured its concentra-


Figure 6. Phospholipid peroxidation in HL-60 cells and HP100 cells exposed to Mel. PnA-loaded HL-60 and HP100 cells (2.5 µg PnA/106 cells, 2 h at 37 °C) were treated with 40 µg/mL Mel for 3 h. Phospholipids were extracted and separated by HPLC, as described in the Experimental Procedures. PnA-phospholipids (PE, PS, and PC) in HL-60 and HP100 cells were detected fluorometrically at 420 nm after excitation at 324 nm. In some experiments, HL-60 cells were cotreated with 50 U/mL CAT and HP100 cells were treated with 100 mM 3-AT 1 h before Mel exposure. Data points represent the mean ( SE of three separate experiments. **p < 0.01 vs Mel-treated HL-60 cells. #p < 0.05 vs Mel-treated HP100 cells without 3-AT treatment.

tion in HL-60 cells and HP100 cells over a 4 h period of time after treatment with Mel. Previous studies reported that elevated intracellular GSH may be associated with resistance to chemotherapeutic agents such as cisplatin and Mel in a number of tumors and tumor cell lines (37, 38). In the present study, however, there was no significant difference in cellular GSH levels between HL-60 cells and HP100 cells (GSH content under normal culture conditions: 45.59 ( 1.76 nmol/mg protein for HL-60 cells; 49.19 ( 0.84 nmol/mg protein for HP100 cells). GSH decreased slightly (by approximately 10% after 4 h) but not significantly following Mel treatment of HL-60 cells (data not shown). No GSH depletion was observed in HP100 cells (data not shown). There was no significant difference in GSH levels between the two cell lines after 4 h of exposure to Mel (data not shown). These results suggest that GSH was not likely the major target of Melinduced ROS production during apoptosis. 3. PS Oxidation during Mel-Induced Apoptosis in HL-60 Cells and HP100 Cells. We have previously demonstrated that PS is preferentially oxidized during oxidant- and nonoxidant-induced apoptosis (4, 15-18). Given that ROS production was detectable at early stages of Mel-induced apoptotsis in HL-60 cells but not in HP100 cells and that GSH was not the major target of oxidative stress, we were eager to compare PS peroxidation in these two cell lines. Therefore, we next examined phospholipid peroxidation in HL-60 cells and HP100 cells exposed to Mel. We used metabolic labeling of phospholipids with an oxidation sensitive fluorescent fatty acid, PnA, followed by fluorescence HPLC to analyze oxidized phospholipids (30, 39). At 3 h following treatment of HL-60 cells with Mel, only PS displayed marked peroxidation, detected as a loss of fluorescent PnA-PS, relative to the control HL-60 cells (Figure 6). This oxidation of PnA-PS was not detectable at earlier time points (2 h, data not shown). More abundant phospholipid classes, PC and PE, did not undergo oxidation (Figure 6). When HL-60 cells were incubated with Mel in the presence of exogenous CAT, PS oxidation decreased by 60% significantly (Figure 6). Mel did not induce any peroxidation of either PS or


Figure 7. PS externalization in HL-60 cells and HP100 cells following treatment with Mel. Cells (1 × 106 cells/mL) were incubated in the absence (control) or presence of 40 µg/mL Mel for 4 h at 37 °C. In some experiments, HP100 cells were treated with 50 mM 3-AT 1 h before Mel exposure. At the end of incubation, cells were collected and PS externalization was determined using Annexin V-FITC Apoptosis Detection Kit by flow cytometry as described in the Experimental Procedures. The results of the densitometric analysis are shown as fold increase to control. Data are means ( SE of three separate experiments. **p < 0.01 vs control HL-60 cells. #p < 0.01 vs Mel-treated HP100 cells without 3-AT treatment.

the other two classes of phospholipids (PC and PE) in HP100 cells (Figure 6). However, HP100 cells treated with Mel in the presence of 3-AT demonstrated similar PS oxidation as HL-60 cells treated with Mel (Figure 6). Combined, these results indicate that Mel-induced ROS were not involved in random oxidation of abundant intracellular reductants such as water soluble GSH or membrane phospholipids PC and PE. Rather, oxidizing equivalents of ROS produced during Mel-induced apoptosis in HL-60 cells were spent on selective oxidation of one particular class of phospholipids, PS. 4. Comparison of Mel-Induced PS Externalization on Plasma Membrane Surface in HL-60 Cells and HP100 Cells. Our previous work has linked PS oxidation with its transmembrane migration from the inner leaflet of the plasma membrane to its outer leaflet during apoptosis (40-44). Because Mel-induced apoptosis was accompanied by PS oxidation in HL-60 cells but not in HP100 cells, we further tested PS externalization in these two cell lines using annexin V binding (Figure 7). Exposure to Mel for 4 h elicited significantly increased externalization of PS in HL-60 cells and to a lesser extent in HP100 cells (Figure 7). Although HP100 cells were resistant to 5 mM H2O2 (Figure 1C), Mel caused a slight increase in apoptotic cell number (Figure 2C) and caspase-3 activity (Figure 2D) in HP100 cells at 4 h. This suggests that Mel-induced apoptosis may also involve some yet to be identified alternative mechanisms, whose contribution to apoptosis is relatively minor. However, when HP100 cells were pretreated with 3-AT, Mel caused similar PS externalization in HP100 cells as in HL-60 cells (Figure 7), indicating that H2O2 generated during Mel-induced apoptosis is indeed involved in mechanisms of PS externalization likely via PS oxidation. Assessment of Mitochondrial Involvement in ROS Generation and PS Oxidation/Externalization during Mel-Induced Apoptosis. In our previous studies, we have put forward a hypothesis that two important factors originating from mitochondria are involved in

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Figure 8. Time course of cytochrome c release into the cytosol in HL-60 cells and HP100 cells exposed to Mel. Cells (1 × 106 cells/mL) were incubated in the presence of 40 µg/mL Mel for 1-4 h at 37 °C under a 5% CO2 and 95% air atmosphere. At the indicated time points, a cytosolic fraction was prepared and 20 µg of protein was loaded into each lane for 12% SDS-PAGE, followed by Western blot analysis with anti-human cytochrome c monoclonal antibody as described in the Experimental Procedures. The results of the densitometric analysis are shown as fold increase to 0 h content. Data points represent the mean ( SE of three separate experiments. **p < 0.01 vs 0 h control in HL-60 cells. #p < 0.05 vs 4 h treatment of HL-60 cells with Mel.

selective redox catalysis of PS oxidation during apoptosiss cytochrome c and H2O2. Because our results demonstrated the critical role of H2O2 in PS oxidation, we were interested in comparative evaluation of mitochondrial events during Mel-induced apoptosis in HL-60 cells and HP100 cells. To this end, we performed measurements of cytochrome c release from mitochondria into the cytosol as well as changes in ∆Ψm in these cells. 1. Release of Mitochondrial Cytochrome c in HL60 Cells and HP100 Cells following Exposure to Mel. We measured cytochrome c in the cytosol of HL-60 cells and HP100 cells by Western blotting over 4 h after apoptosis induction by Mel. Cytosol fractions from HL60 cells exposed to Mel for 0-2 h did not contain any detectable cytochrome c protein (Figure 8). A significant accumulation of cytosolic cytochrome c was observed after 3 h of treatment with Mel, which further increased by 4 h. By contrast, only trace amounts of cytochrome c protein were detectable in the cytosol fractions from HP100 cells throughout 4 h of incubation with Mel (Figure 8). 2. Mel-Induced Mitochondrial Permeability Transition in HL-60 Cells and HP100 Cells. To assess the contribution of mitochondria to Mel-induced apoptosis (45), we examined changes in the ∆Ψm in HL-60 cells and HP100 cells. In HL-60 cells, ∆Ψm collapsed only 4 h after Mel treatment (Figure 9); we failed to detect any decrease in ∆Ψm up to 3 h (data not shown). No convincing changes in ∆Ψm over a 4 h incubation time was observed in HP100 cells (Figure 9 and data not shown for 1, 2, and 3 h). Treatment of HL-60 cells and HP100 cells with a mitochondrial uncoupling agent, CCCP (50 µM for 1 h), caused a remarkable drop in ∆Ψm (Figure 9).

Discussion Role of H2O2 in the Execution of the Apoptotic Program. Involvement of mitochondria-dependent mech-

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Figure 9. Changes in ∆Ψm in HL-60 cells and HP100 cells exposed to Mel. Cells (1 × 106 cells/mL) were incubated in the absence (control) or presence of 40 µg/mL Mel for 4 h at 37 °C under a 5% CO2 and 95% air atmosphere. Cells were labeled with 250 nM CMXRos for the last 30 min of treatment, and thereafter, ∆Ψm was analyzed by flow cytometry. As a positive control of ∆Ψm disruption, cells were exposed to 50 µM CCCP for 1 h. Representative data from three separate experiments are shown.

anisms (46, 47) is critical to execution of intrinsic apoptotic pathways and is also required for execution of extrinsic apoptotic program in so-called type II cells (48). One of the central and early events is release of cytochrome c from mitochondria into the cytosol, which is believed to cause disruption of electron transport and hence enhanced generation of one electron-reduced species of molecular oxygen such as O2- and its dismutation product, H2O2 (2, 49-57). H2O2 has been demonstrated to act as an important messenger in signal transduction pathways [e.g., via regulation of NF-kB (58, 59), mitogenactivated protein kinases (60) likely through oxidative modification of reactive cysteine residues (61)]. At least several components of apoptotic machineryscaspases (62, 63) and mitochondrial proteins forming the permeability transition pore (64-66)shave been shown to be sensitive to effects of H2O2. In line with this, overexpression of peroxiredoxins, antioxidant enzymes catalyzing H2O2 decomposition via a peroxidase reaction (67), is correlated with resistance to apoptosis (68). Thus, H2O2 is an attractive candidate as a secondary messenger for final common mediator of apoptosis; yet, a specific role for H2O2 in the execution or resolution of the apoptotic program has not been established. CAT is one of the most potent enzymes involved in elimination of H2O2, and cells overexpressing CAT have been used to investigate the potential role of H2O2 in a variety of biological experiments (14, 69, 70). Hence, we utilized CAT overexpressing HP100 cells, a subclone of human leukemia HL-60 cells, to determine the extent to which execution of apoptotic program, including its part related to PS signaling, is dependent on the availability of H2O2. With the exception of CAT, we found no convincing difference in the expression of either other primary antioxidant enzymes or apoptosis-related proteins (Bcl-2 and Bax) in HP100 cells as compared with the parental HL-60 cells (Figure 1). We also established that HP100 cells (but not HL-60 cells) were resistant to apoptosis induced by exogenously added H2O2. These results suggest that the HP100 cell line may be a valuable tool to examine involvement of endogenous H2O2 in apoptosis. To this end, we chose to utilize a model of intrinsic apoptosis induced by an alkylating agent, Mel (35, 71-75). Production of H2O2 (as measured by DCF fluorescence) was readily detectable as early as 1 h after Mel exposure


of HL-60 cells but not CAT-rich HP100 cells. Biomarkers of apoptosis, such as release of cytochrome c, disruption of ∆Ψm, caspase-3 activation, and chromatin condensation, became apparent much later, 3 h and onward after Mel treatment of HL-60 cells. The emergence of essentially all biomarkers of apoptosis was dramatically delayed in HP100 cells as compared with HL-60 cells. While the source of H2O2 produced at a very early phase of Mel-induced apoptosis in HL-60 cells remains unknown, it is unlikely that dismutation of O2- derived from disruption of mitochondrial electron transport is the major contributor as it happened significantly later. Alternatively, H2O2 can be produced directly by several oxidases such as 2-hydroxyacid oxidase and urate oxidase (76). Further studies employing SOD overexpressing cells may be instrumental in addressing the issue. Whatever the source of early H2O2, our results suggest that it is critical for triggering the Mel-induced apoptotic program as evidenced by (i) effective suppression of apoptosis in HP100 cells, (ii) reconstitution of sensitivity of HP100 cells to apoptosis after CAT inhibition by 3-AT, and (iii) partial inhibition of apoptosis in HL-60 cells by exogenously added CAT. Role of H2O2 in PS Signaling during Apoptosis. H2O2 produced during apoptosis may undergo enzymatic degradation by CAT and other enzymes (e.g., GPX, peroxiredoxins) and/or act as a secondary messenger interacting with sensitive intracellular targets and contributing to execution of apoptosis. Our results clearly demonstrate that CAT played the major role in elimination of Mel-induced endogenous production of H2O2. GPXdependent pathways were not likely involved in H2O2 scavenging as essentially no GSH consumption was detected over 4 h challenge of HL-60 cells or HP100 cells with Mel. Thus, GSH was not the major target of oxidative stress during Mel-induced apoptosis. Interestingly, our results on phospholipid peroxidation revealed that the two most abundant classes of membrane phospholipids, PC and PE, did not undergo any significant oxidation. Instead, a relatively minor phospholipid species, PS, was markedly oxidized after 3 h of treatment with Mel. This selective oxidation of PS was likely H2O2-mediated as it was observed only in HL-60 cells (but not in HP100 cells) where it was significantly inhibited by exogenously added CAT and became evident in 3-AT-treated HP100 cells. Measurements at 2 h time point showed no significant oxidation of PS or any other phospholipids in either HL-60 cells or HP100 cells. Two major questions can be asked. (i) What causes selectivity of PS oxidation? (ii) Given that H2O2 production was detectable as early as 1 h after Mel exposure of HL-60 cells and its steady state level was maintained during the subsequent 2-4 h of incubation, one may wonder why PS oxidation was delayed as compared to H2O2 generation. The answer is that there may be additional factor(s) required for effective and selective catalysis of PS oxidation. Our previous work has indicated that cytochrome c released from mitochondria into the cytosol played a critical role of a redox catalyst in the reaction of PS oxidation (15, 19, 20). Indeed, PS oxidation took place 3 h after exposure of HL-60 cells to Mel (Figure 6) and paralleled the departure of cytochrome c from mitochondria and its appearance in the cytosol (Figure 8). The fact that neither cytochrome c release nor PS oxidation occurred in Mel-treated HP100 cells is in line with our previous results (77) on the likely involvement


of cytochrome c in catalysis of PS oxidation (detected in HL-60 cells). However, further direct studies are necessary to establish whether cytochrome c release is, indeed, essential for PS oxidation. In mitochondria, cytochrome c specifically interacts with an anionic phospholipid, cardiolipin (78). Once released from mitochondria into the cytosol during apoptosis, cytochrome c interacts with apoptotic proteaseactivating factor-1 (Apaf-1), dATP, and cytosolic procaspase-9 to form “apoptosome” and activate downstream effector caspases including caspase-3 (79). This important role of cytochrome c does not entail its redox properties. We have previously hypothesized that in addition to the role in apoptosome formation, being a basic protein, cytochrome c electrostatically interacts with negatively charged PS on the inner leaflet of plasma membrane (15, 19, 20). We further assumed that in the presence of H2O2, cytochrome c can act as a peroxidase and catalyze oxidation of polyunsaturated phospholipids in its microenvironment, which happens to be PS. Moreover, interaction of cytochrome c with anionic phospholipids, such as PS, bolsters peroxidase activity of cytochrome c, thus enhancing its ability to oxidize PS. In cell-free model experiments, we used low temperature EPR spectroscopy and demonstrated that activation of cytochrome c by H2O2 to produce its oxoferryl species strongly depends on the presence of PS-containing liposomes (77). Interestingly, release of cytochrome c occurred earlier than dissipation of ∆Ψm but much later than production of H2O2 suggesting that elimination of mitochondrial membrane potential is not required for PS oxidation (see the time course of ∆Ψm in Mel-treated HL-60 cells mentioned in the Results). While in some apoptotic systems loss of ∆Ψm may be an early event in the apoptotic process, there is emerging data suggesting that depending on the model of apoptosis, the loss of ∆Ψm may not be an early requirement for apoptosis but on the contrary may be a consequence of the apoptotic signaling pathway (80). In fact, a number of recent reports state that loss of ∆Ψm is not required for cytochrome c release and that loss in inner membrane potential happens only after caspase-3 activation, which is downstream to release of cytochrome c (81, 82). Overall, depending on the cell system under investigation and the apoptotic stimuli used, dissipation of ∆Ψm may or may not be an early event in the apoptotic pathway. In particular, HL-60 cells can undergo apoptosis in response to a number of cytotoxic insults (actinomycin-D, etoposide, and staurosporine) without showing significant changes in ∆Ψm (83). Departure of cytochrome c from mitochondria is believed to be associated with peroxidation of cardiolipin (84, 85). It is tempting to speculate that very early Mel-induced generation of H2O2 may be involved in cytochrome c-catalyzed oxidation of cardiolipin. Hence, H2O2 may act as a messenger using its capacity to catalyze oxidation of two phospholipids with which it preferentially binds, cardiolipin inside mitochondria and PS in the cytosolic leaflet of plasma membrane. Further studies will reveal redox catalytic potency of cytochrome c toward cardiolipin in miotchondria during early stages of apoptosis. We have previously reported that oxidation of PS is closely related to its externalization and subsequent recognition and phagocytosis of apoptotic cells by macrophages (4, 15, 17, 18). Furthermore, we established that oxidized PS acts as an additional signal effectively

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recognized by macrophage receptor(s) (18). In the present study, we found that Mel-induced PS oxidation was also associated with pronounced externalization of PS on the surface of HL-60 cells. In addition, CAT inhibition by 3-AT restored the sensitivity of HP100 cells to ROS generation, PS oxidation, and PS externalization after Mel exposure. This suggests that the messenger role of Mel-induced H2O2 is important not only for PS oxidation but also for its externalization, hence for recognition of apoptotic cells by phagocytes. Furthermore, as interaction of externalized PS with its cognate macrophages receptor is indispensable for effective clearance of apoptotic cells without concomitant release of proinflammatory cytokines (86), it is likely that formation of endogenous H2O2 during apoptosis is important for successful resolution of inflammation.

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Acknowledgment. This work was supported by NIH HL70755.

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