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J. Biol. Chem., Vol. 279, Issue 7, 6056-6064, February 13, 2004
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**
From the
Departments of Environmental and Occupational Health and Pharmacology, **Cancer Research Institute, University of Pittsburgh, Pittsburgh, Pennsylvania 15260, ¶Department of Cancer Biology, University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030, and ||Division of Toxicology, Institute of Environmental Medicine, Karolinska Institutet, 17177 Stockholm, Sweden
Received for publication, September 8, 2003 , and in revised form, November 13, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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We have recently shown that PS externalization during apoptosis is preceded by its selective oxidation, likely catalyzed by cytochrome c released from mitochondria (5, 6). We have also demonstrated that oxidized PS (PS-OX) may serve as an "eat me" signal for macrophage receptor(s), thus facilitating recognition and PS-dependent engulfment of apoptotic cells. (7, 8) Based on these observations, we hypothesized that PS oxidation acts as an essential component of the signaling pathway that is required for PS externalization and the safe clearance of apoptotic cells by macrophages (5, 8, 9, 10). Our hypothesis predicts that lipid antioxidants capable of blocking PS oxidation will inhibit PS externalization and/or recognition of apoptotic cells by phagocytes. An important feature of such a lipid antioxidant, however, is that it should block phospholipid oxidation without affecting other redox-sensitive mechanisms.
Our previous work has established that a phenolic antitumor drug, etoposide (a topoisomerase II inhibitor), acts as a powerful lipid antioxidant but does not protect thiols against oxidation because of a relatively high reactivity of etoposide phenoxyl radicals toward SH-groups (11). Etoposide has been reported to cause DNA damage and induce apoptosis accompanied by ROS generation (12, 13). It is conceivable that the fastidious lipid antioxidant traits of etoposide may dissociate PS-dependent signaling pathways of apoptosis from the final common pathway for apoptosis by inhibiting PS oxidation. However, etoposide effects on phospholipid peroxidation, as they relate to PS-dependent signaling pathways of apoptosis, have not been documented to date.
In the present study, we used etoposide as a tool to dissect the mechanism of PS signaling during apoptosis. We found that etoposide fails to trigger PS oxidation and externalization in HL-60 cells. Moreover, etoposide was able to block oxidation of all major phospholipids, including PS, during oxidant (H2O2)-induced apoptosis in these cells. The blockade of phospholipid oxidation correlated with the etoposide-dependent inhibition of PS externalization and phagocytosis of apoptotic HL-60 cells by J774A.1 macrophages. The etoposide-mediated dissociation of PS signaling during apoptosis could be overcome by the integration into HL-60 cells of exogenous PS, and even more so upon integration of PS-OX, thus emphasizing the essential role of PS oxidation in the clearance of apoptotic cells. These results demonstrate that etoposide can modulate PS-dependent externalization and recognition of apoptotic cells by macrophages by means of regulation of PS oxidation.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Cell Culture—HL-60 human promyelocytic leukemia cells (American Type Culture Collection) were grown in RPMI 1640 medium supplemented with 12.5% heat-inactivated fetal bovine serum at 37 °C in a humidified atmosphere (5% CO2 plus 95% air) at 37 °C. Cells from passages 25-40 were used for the experiments. The density of cells at collection time was 0.5 x 106 cells/ml. HL-60 cells were incubated in the presence of etoposide (10 µM or 50 µM) and/or H2O2 in fetal bovine serum-free RPMI 1640 medium without phenol red for 2 h at 37 °C. H2O2 (25 µM) was added four times (every 30 min of incubation). In the case of combination of etoposide with H2O2, etoposide was added 15 min before the addition of H2O2. Macrophages J774A.1 (from the American Type Culture Collection) were grown in Dulbecco's modified Eagle's medium supplemented with 10% heat-inactivated fetal bovine serum, 100 units/ml penicillin, 100 µg/ml streptomycin, and 50 µg/ml gentamycin sulfate in a humidified atmosphere (5% CO2 plus 95% air) at 37 °C.
Apoptotic Nuclear Morphology—HL-60 cells were incubated in the presence of etoposide and/or H2O2 for 2 h at 37 °C. At the end of the incubation, cells were washed and re-suspended in PBS. Hoechst 33342 (2 µg/ml) was added, and cells were examined under fluorescent microscopy. Results are 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 (>200 cells).
Caspase-3 Activity—The activity of caspase-3 was determined by using a commercially available kit (Molecular Probes). At specified time intervals, aliquots of etoposide- and/or H2O2-stimulated HL-60 cell suspensions were taken, cells were washed twice with PBS and lysed for 30 min in lysis buffer (Molecular Probes). The suspensions were centrifuged at 4 °C, and supernatants were collected as lysates. For measurement of caspase activity, 50 µl of lysates were combined with 50 µl of reaction buffer, containing 20 mM PIPES, 4 mM EDTA, 10 mM dithiothreitol, 0.2% CHAPS, 0.2 mM Asp-Glu-Val-Asp 7-amino-4-methylcoumarin (DEVD-AMC, a fluorogenic peptide substrate), pH 7.4, and incubated for 30 min at 25 °C. After incubation, fluorescence was measured in a Packard FusionTM Multifunctional plate reader (PerkinElmer Life Sciences) using excitation at 365 ± 50 nm and emission at 465 ± 35 nm. The protein concentration in cell lysates was measured using the Bio-Rad assay.
MCLA-enhanced Chemiluminescence—HL-60 cells (2 x 106 cells/ml) were pre-warmed in PBS containing 0.5 mM CaCl2, 1 mM MgCl2, and 30 mM glucose for 5 min at 37 °C. After that, etoposide (50 µM) was added, and cells were monitored for 30 min at 37 °C using Luminescence Analyzer 633 (Coral Biomedical Inc., San Diego, CA) set at continuous mixing in the presence of 4 µM MCLA. Assays were performed in the absence and in the presence of superoxide dismutase (50 units/ml) added 5 min prior to the addition of etoposide. The total amount of O2 produced was estimated as the area under the curve (mV x s) upon etoposide stimulation and after subtracting values obtained in the presence of superoxide dismutase. Data were collected and analyzed with the Multiuse PC software version 2.0.2 for Luminoskan 1251 Carousel (Labsystems).
Assay of GSH—GSH content in the cells was determined fluorometrically using ThioGloTM 1 as previously described (14). Briefly, cells treated with etoposide and/or H2O2 for 2 h at 37 °C were collected by centrifugation, washed, and re-suspended in PBS. GSH was measured in cell lysates prepared by freezing and thawing cells. Immediately after the addition of ThioGloTM 1 to the cell lysates, fluorescence was measured in a Packard FusionTM Multifunctional plate reader (PerkinElmer Life Sciences) using excitation of 390 ± 15 nm and emission of 515 ± 30 nm.
Annexin V Staining of Externalized PS—PS exposure was determined by flow cytometric detection of annexin V staining using a protocol outlined in the annexin V-FITC apoptosis detection kit (BioVision Research Products, Mountain View, CA). Briefly, HL-60 cells (0.5 x 106) exposed to etoposide and/or H2O2, washed once with PBS and re-suspended in binding buffer were stained with annexin V (0.5 µg/ml) and propidium iodide (0.6 µg/ml) for 5 min at room temperature. After staining, cells were immediately analyzed using a FACScan flow cytometer (BD Biosciences) 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. 10,000 events were collected and analyzed using the LYSISTM II software (BD Biosciences).
Derivatization of Externalized PS with Fluorescamine—Labeling of externalized PS with fluorescamine (a probe that reacts with lipids containing primary amino groups) was performed as described previously (8). Briefly, HL-60 cells (3 x 107) exposed to etoposide and/or H2O2 were suspended in labeling buffer (150 mM NaCl, 5 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 5 mM NaHCO3, 5 mM glucose, and 20 mM HEPES, pH 8.0). Cells were gently agitated in the presence of fluorescamine (200 µM) for 30 s. The reaction was stopped by the addition of 40 mM Tris-HCl, pH 7.4. Cells were collected by centrifugation and lipids were extracted by the Folch procedure (15). Lipids were analyzed by two-dimensional HPTLC using a solvent system of chloroform:methanol:28% ammonium hydroxide (65:25:5, v/v/v) in the first direction and chloroform:acetone:methanol:glacial acetic acid:water (50:20:10:10:5, v/v/v/v/v) in the second. Fluorescamine-modified PS was localized by exposure of HPTLC plates to UV light by using a Fluor-STM MultiImager (Bio-Rad). Unmodified phospholipids were visualized under visible light in a Fluor-STM MultiImager (Bio-Rad) after exposure of HPTLC plates to iodine vapor. The phosphorus content of phospholipids was determined according to Bottcher et al. (16) after scraping representative spots from the plate. The amount of modified PS is expressed as percentages of total PS (unmodified plus modified) recovered from the plate on the basis of phosphorus content assay.
Assay of Phospholipid Peroxidation—cis-PnA was incorporated into HL-60 cells as described previously (17). Briefly, HL-60 cells were incubated in serum-free RPMI medium 1640 without phenol red in the presence of cis-PnA (1 µg cis-PnA/106 cells) for 2 h at 37 °C. At the end of incubation, cells were washed with PBS containing human serum albumin (fatty acid-free, 0.5 mg/ml) to remove an excess of unbound cis-PnA. cis-PnA-labeled cells were treated with etoposide (10 or 50 µM) in the presence or in the absence of H2O2 for 2-6 h at 37 °C in 25 mM HEPES buffer containing 137 mM NaCl, 2.7 mM KCl, 1.5 mM KH2PO4, 8 mM Na2HPO4, 10 mM glucose, pH 7.4. H2O2 (25 µM) was added every 30 min during the initial 2 h of incubation. At the end of the incubation, lipids were extracted by the Folch procedure (15). The lipid extracts were separated by normal-phase HPLC using a 5-µm microsorb MV column (4.6 x 250 mm; Rainin Instrument Co. Inc.) as described previously (17). The separations were performed by using a Shimadzu LC-600 high-performance liquid chromatography system with an in-line configuration of RF 10 AXL fluorescence detector. Fluorescence of cis-PnA was measured at 420 nm emission after excitation at 324 nm. Data were processed and stored in digital form with Shimadzu EZChrom software. Lipid phosphorus was determined by a micro-method (18).
Assay of Aminophospholipid Translocase Activity—Aminophospholipid translocase (APT) activity was measured using modifications of the methods of McIntyre and Sleight (19) and Williamson and et al. (20). HL-60 cells (4 x 106) treated with etoposide (50 µM, 2 h at 37 °C) in the presence or absence of H2O2 were centrifuged (400 x g, 10 min) and washed once in incubation buffer (136 mM NaCl, 2.7 mM KCl, 2 mM MgCl2, 5 mM glucose, 10 mM HEPES, pH 7.5). Cell pellets were re-suspended in incubation buffer (5 x 106 cells/ml) containing 500 µM phenylmethylsulfonyl fluoride, transferred to a microfuge tube, and placed in ice-water for 10 min. NBD-labeled PS (ethanol solution) was added to cells (final concentration, 10 µM) and incubated for 10 min at 4 °C. Labeled cells were centrifuged and re-suspended at the same density in incubation buffer with phenylmethylsulfonyl fluoride. Cell suspensions were placed in a 28 °C water bath to initiate internalization, and 75-µl aliquots of cell suspension were removed at various time intervals (1-20 min) and placed into 2.5 ml of incubation buffer including the reducing agent and 10 mM sodium dithionite. Fluorescence (excitation = 470 nm, emission = 540 nM) was then recorded (within 30-60 s). Samples from the last time point were also placed in incubation buffer without dithionite to obtain total fluorescence intensity (FLtotal). Internalized fluorescence at various times (FLt) was normalized as a percent of the total fluorescence by the following equation: % internalized = (FLt - FL0)/(FLtotal - FL0) x 100. To inhibit APT activity, HL-60 cells were incubated in the presence of NEM (10 µM) for 10 min at 37 °C. At the end of incubation, cells were washed twice with PBS, and APT activity was determined as described above.
Phagocytosis of HL-60 Cells by J774A.1 Macrophages—J774A.1 macrophages were seeded into an eight-well chamber slide (5 x 104 cells/well) and cultured overnight in DMEM medium. Before adding target (control or etoposide and/or H2O2-treated HL-60) cells, stimulated cells were washed with serum-free RPMI medium 1640 without phenol red and fluorescently labeled with Cell TrackerTM Orange (5 µg/ml, 10 min at 37 °C) and subsequently washed again (three times) with PBS. Fluorescently labeled cells (5 x 105 cells/well) were co-cultured with macrophages for 1 h at 37 °C. After incubation, unbound target cells were washed three times with RPMI medium 1640 and three times with PBS; well contents were fixed with a solution of 2% formaldehyde in PBS containing Hoechst 33342 (1 µg/ml) for 30 min at room temperature. The cells were examined under a Nikon ECLIPSE TE 200 fluorescence microscope (Tokyo, Japan) equipped with a digital Hamamatsu charge-coupled device camera (C4742-95-12NBR) and analyzed using the MetaImaging SeriesTM software version 4.6 (Universal Imaging Corp., Downingtown, PA). Macrophages that had a side-by-side connection with target cells (binding) and/or internalized target cells (engulfment) were considered phagocytosis-positive. A minimum of 300 macrophages were analyzed per experimental condition. Results are expressed as the percentage of the phagocytosis-positive macrophages.
Preparation of PS- and PS-OX-containing Liposomes—PS was oxidized by incubation with a water-soluble azo-initiator of peroxy radicals, 2,2'-azo-bis-(2-amidinopropane) hydrochloride. PS in chloroform was dried under nitrogen and PBS was added to achieve the final concentration of 5 mM. The lipid was incubated with 2,2'-azo-bis-(2-amidinopropane) hydrochloride (50 mM) at 37 °C for 4 h and then extracted with chloroform:methanol (2:1, v/v). Oxidation was assessed by measuring the absorbance of PS hydroperoxides with conjugated dienes at 234 nm. Approximately 30% of PS molecules were estimated to be oxidized after a 4-h incubation, whereas 70% remained non-oxidized; this mixture of PS species is hereafter referred to as oxidized PS (PS-OX). Small unilamellar liposomes containing 50% phosphatidylcholine (PC) and 50% PS (nonoxidized PS or PS-OX) were produced as described by Fadok et al. (21). Individual phospholipids, stored in chloroform, were dried under nitrogen. PBS was added to obtain a phospholipid concentration of 1 mM, and the lipid mixture was vortexed and sonicated for 3 min on ice. All liposomes were used immediately after preparation.
Treatment of HL-60 Cells with Liposomes—Naïve or etoposide-treated (50 µM for 2 h at 37 °C) apoptotic HL-60 cells were incubated with NEM (10 µM, 10 min at 37 °C) to inhibit APT. At the end of the incubation, cells were centrifuged and NEM was washed out. NEM-treated cells were incubated in the presence of PC+PS or PC+PS+PS-OX liposomes (2.5-150 µM) for 30 min at 37 °C. After that, cells were washed with PBS labeled with Cell TrackerTM Orange, and phagocytosis assays were performed as described above.
Statistics—The results are presented as mean ± S.E. values from at least three experiments, and statistical analyses were performed by either paired or unpaired Student's t test or one-way ANOVA. The statistical significance of differences was set at p < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Etoposide Treatment of HL-60 Cells Prevents H2O2-induced Oxidation of PS—Execution of the apoptotic program triggered by both oxidant and non-oxidant stimuli may be associated with the production of reactive oxygen species (ROS) and ensuing oxidative stress (22). Therefore, we determined ROS production and oxidative stress in our model. To determine whether apoptosis was accompanied by the production of superoxide, we used a superoxide-specific enhancer of chemiluminescence, MCLA. The results in Fig. 2A show that superoxide production was significantly increased in HL-60 cells exposed to etoposide (50 µM for 2 h at 37 °C) as compared with untreated cells. To assess the degree of oxidative stress, we measured the level of intracellular GSH in HL-60 cells. We found that after etoposide (50 µM) stimulation for 2 h at 37 °C, the GSH content significantly decreased to 63.6 ± 2.0% of its initial level (Fig. 2B). Similarly, H2O2 (25 µM x 4, 2 h) caused a pronounced depletion of GSH. Finally, when HL-60 cells were exposed to a combination of etoposide and H2O2, an even stronger depletion of GSH was evident. This suggests that etoposide is not likely to interfere with thiol-dependent redox signaling pathways of apoptosis.
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7% cells remained annexin V-positive. At 10 µM, etoposide did not significantly reduce the number of H2O2-induced annexin V-positive cells (data not shown). Increasing the treatment time of HL-60 cells with etoposide (50 µM) to 4 and 6 h, however, resulted in a significantly increased number of cells with externalized PS (up to 8.1 ± 1.3 and 11.3 ± 0.4%, respectively) (Fig. 5A). After 4 and 6 h of exposure of HL-60 cells to H2O2, 22.4 ± 2.5 and 30.8 ± 2.4% cells were annexin V-positive. At these later time points, etoposide did not decrease H2O2-induced externalization of PS, although it remained lower than the expected additive sum (30.5 and 42.5% at 4 and 6 h, respectively). Most importantly, the dependence of PS externalization on its oxidation held true for these later time-points as well. In particular, elevated levels of PS externalization were associated with PS oxidation revealed by decreased contents of cis-PnA-PS in HL-60 cells treated with etoposide alone, H2O2 alone, or a combination of H2O2 plus etoposide at both 4 and 6 h. Although etoposide did not protect PS against H2O2-induced oxidation at either 4 or 6h, in both cases the oxidation levels were lower than the expected additive effects of etoposide alone and H2O2 alone. Our incubation conditions (50 µM etoposide and/or H2O2 25 µM x 4) were not associated with any significant accumulation of necrotic (PI-positive/annexin V-positive) cells during 2, 4, or 6 h exposures as compared with their content in control samples (1.5-2.0%).
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15 min of incubation. In contrast, almost all NBD-PC remained on the external surface over the 20-min incubation period, indicating that internalization was specific for aminophospholipids (data not shown). Pretreatment of cells with NEM (10 µM, 10 min at 37 °C) resulted in an essentially complete inhibition of APT (Fig. 7A). H2O2 (25 µM x 4, 2 h) reduced the ability of cells to internalize NBD-PS by 2-fold (Fig. 7A), and etoposide did not affect H2O2-induced APT inhibition (Fig. 7B). Accordingly, the rate of NBD-PS internalization was 1.6-fold higher in control cells than in H2O2-exposed cells (both in the absence and in the presence of etoposide; Fig. 7B). No significant changes in the ability of HL-60 cells to internalize NBD-PS were observed after exposure to etoposide alone (2 h at 37 °C). Thus, etoposide did not protect APT against H2O2-induced inactivation.
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2.0-2.5%) were detected when control (non-treated) HL-60 cells were incubated (for 1 h) with J774A.1 macrophages. Etoposide-treated HL-60 cells were phagocytosed at a slightly (albeit significantly, 1.3-fold) higher level than control cells. In contrast, H2O2-treated HL-60 cells were readily ingested by J774A.1 macrophages, yielding a 2.1-fold increase in the number of phagocytosis-positive cells. Remarkably, etoposide (at 50 µM) caused inhibition of phagocytosis of HL-60 cells treated with H2O2 (Fig. 8A) insofar as the level of phagocytosis-positive macrophages was not different from that detected in the presence of etoposide alone.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Our data clearly demonstrate that etoposide can trigger oxidative stress and induce apoptosis in HL-60 cells. Despite these effects, etoposide effectively prevented phospholipid oxidation during the execution of the apoptotic program. Moreover, in line with previous work (33), etoposide-induced apoptosis in HL-60 cells was not accompanied by PS externalization at early stages of apoptosis. In H2O2-treated cells, etoposide conferred complete protection of phospholipids against oxidation and inhibited PS externalization. Importantly, etoposide did not block the execution of other features of the H2O2-induced apoptotic program (i.e. caspase activation and nuclear fragmentation) and did not prevent H2O2-induced inhibition of APT. Etoposide is thus able to dissociate PS signaling from the common pathway of apoptosis by inhibiting oxidation of phospholipids, particularly PS. It is worth noting that PS externalization does, in fact, occur during late stages of the etoposide-induced apoptotic program (4 and 6 h) (34-36). These findings would seem to contradict our conclusion concerning antioxidant blockade of PS externalization during etoposide-triggered apoptosis. However, one needs to keep in mind that scavenging of reactive radicals by phenolic antioxidants is associated with their oxidation, i.e. depletion of antioxidant capacity (37, 38). In fact, we observed that low doses of etoposide (10 µM) were much less effective in protecting against phospholipid peroxidation as well as in inhibiting PS-dependent signaling compared with etoposide used at 50 µM. It is likely that oxidative metabolism of the phenolic moiety of etoposide (known to form quinone derivatives; Ref. 39) is responsible for its ineffectiveness as an inhibitor of PS externalization during late stages of apoptosis. In line with this, we found that exposure of HL-60 cells to etoposide and a combination of H2O2 plus etoposide for 4 and 6 h resulted in a significant PS oxidation. Notably, even at these later time points, PS externalization correlated with PS oxidation, suggesting the existence of a causative link between the two processes.
APT, a P-type ATPase, maintains the asymmetrical distribution of PS in cells (2). Inhibition of APT has been suggested as playing an important role in PS externalization during apoptosis (40, 41). However, the link between APT inactivation and the execution of the apoptotic program remains unclear. It has been suggested that APT is not a substrate for caspases (42), indicating that alternative mechanisms may be involved in the inhibition or inactivation of this enzyme. In particular, depletion of ATP may contribute to APT inhibition during apoptosis (34, 43). The current data indicate that etoposide inhibits PS externalization through its effect on PS oxidation, rather than by means of a direct inhibition of APT. This is supported by our results demonstrating that etoposide did not prevent H2O2-induced inhibition of APT. Moreover, because etoposide alone did not affect APT activity in HL-60 cells but triggered production of ROS and caspase activation, one may assume that its prevention of phospholipid peroxidation is likely responsible, at least in part, for the lack of APT inhibition. It is tempting to speculate that oxidized PS may contribute to APT inactivation and this effect is minimized in the presence of etoposide. It is possible that oxidized PS acts as an essential regulator of APT activity and PS externalization in intrinsic apoptotic pathways, but it is relatively less important in extrinsic mitochondria-independent (type I) apoptosis.
Etoposide-mediated Inhibition of PS Oxidation Blocks Macrophage Engulfment of HL-60 Cells—Because the externalization of PS on the surface of apoptotic cells acts as an important "eat me" signal for phagocytes, we assumed that etoposide-treated apoptotic HL-60 cells that fail to externalize PS would be less readily engulfed by macrophages. Indeed, we found that etoposide-triggered, early apoptotic cells were poorly engulfed by J774A.1 macrophages. In contrast, apoptosis induced by H2O2 was accompanied by PS externalization, and these cells were successfully eliminated by macrophages. Moreover, the combination of etoposide and H2O2 resulted in a significant inhibition of both H2O2-induced PS externalization and phagocytosis of these cells. Thus, etoposide not only dissociates the common final pathway of apoptosis from the PS signaling pathway essential for phagocytosis, but also blocks this signaling during H2O2-induced apoptosis. Similarly, overexpression of the antiapoptotic protein Bcl-2 prevents PS exposure and macrophage engulfment but does not affect DNA fragmentation or the activation of downstream caspases in Fas-triggered SKW6.4 cells (44). Taken together, these data suggest that the execution of cell death and the clearance of cell corpses are governed, in part, by distinct mechanisms or subprograms.
Our data suggest that etoposide-dependent inhibition of PS oxidation contributes to the failure of macrophages to recognize apoptotic HL-60 cells. In line with these findings, we found that enforced PS externalization in apoptotic HL-60 cells could overcome the etoposide-induced concealment from macrophage recognition. PS-OX integrated into the plasma membrane of etoposide-treated HL-60 cells was an even more potent recognition signal and yielded more effective engulfment by macrophages. These findings concur with our previous studies in which viable Jurkat, Raji, and HL-60 cells were ingested after enrichment with PS and/or PS-OX (8), and further underscore the importance of PS oxidation in PS-dependent signaling pathways of phagocytic clearance of apoptotic cells. Of note, etoposide alone, or in combination with other drugs, was recently shown to induce apoptosis and subsequent phagocytosis of Burkitt lymphoma cell lines; this effect was inhibited upon the addition of H2O2 (45, 46). The apparent contradiction between our findings and those of Shacter et al. (45, 46) is likely explained by the relatively high concentration of H2O2 (200 µM) utilized by these investigators resulting in potent inhibition of apoptosis and accumulation of relatively large amounts of necrotic and "late" apoptotic cells recognizable by macrophages. Indeed, Hampton and Orrenius (47) have shown that H2O2 exerts a dual regulation of caspases, such that low doses (50 µM) are able to trigger apoptosis, whereas high doses (above 200 µM) yield oxidative inactivation of these enzymes and subsequent switching of the mode of cell death to necrosis. Notably, our experimental conditions (50 µM etoposide and/or H2O2 25 µM x 4) did not yield any significant increase in the number of necrotic cells over an incubation of 2-6 h with etoposide and/or H2O2. When high concentrations of H2O2 were used (250 µM), a significant accumulation of necrotic HL-60 cells (up to 30%) was observed already after a 2-h exposure, in agreement with the data by Shacter et al. (45, 46)
Since its introduction in 1971, etoposide has become one of the most widely used and effective antitumor drugs in the treatment of different malignancies in both adults and children (48). As an anticancer drug, etoposide is commonly used in combination therapy in both conventional and high-dose regimens (49-55). Although most common conventional regimens result in etoposide plasma concentrations around 2-10 µM (49-52), it is not unusual in clinical practice to utilize high dose etoposide (500 mg/m2) that may yield peak plasma concentrations of 100 µM or higher (53). Utilization of higher doses of etoposide (1500-2400 mg/m2; Refs. 54, 55) can be expected to achieve even higher plasma concentrations. Therefore, the concentration of etoposide used in the current study is well within the range of therapeutic levels achievable during etoposide treatment. One of the unfortunate side effects of etoposide, particularly in children, is therapy-related acute myelogenous leukemias (AML; Refs. 56, 57). Although specific mechanisms responsible for the induction of AML are not fully understood, they include inhibition of topoisomerase II, likely enhanced by myeloperoxidase-catalyzed one-electron metabolism of etoposide in bone marrow progenitor cells (58). The latter is associated with the production of different reactive free radical intermediates potentially leading to direct genotoxic effects as well as non-genotoxic carcinogenesis (31, 39, 59, 60). Apoptosis and safe elimination of transformed cells by phagocytes are of obvious importance for the prevention of pro-carcinogenic effects of etoposide. Our results suggest that impaired PS signaling and elimination of etoposide-triggered apoptotic cells may contribute to the deleterious side effects of this drug, particularly those leading to AML.
To conclude, we have shown that etoposide, an antitumor agent with unique antioxidant properties, can suppress phospholipid oxidation and PS externalization in HL-60 cells. Moreover, we have shown that PS signaling can be dissociated from other features of the apoptotic program insofar as H2O2-induced apoptosis was potentiated, whereas H2O2-induced PS oxidation and externalization as well as phagocytosis by J774A.1 macrophages were blocked upon incubation of cells with etoposide. Both externalization of PS and oxidation of phospholipids (including PS and PC) have been identified as important determinants of macrophage clearance of apoptotic cells (1, 8, 61, 62). We speculate that oxidation of PS may combine these features of the apoptotic process and facilitate externalization of PS as well as PS-OX, with subsequent recognition of these phospholipid species by neighboring macrophages. Future studies should aim to quantify amounts of PS and its oxidized counterpart on the surface of apoptotic cells as well as to identify the macrophage receptor(s) that are involved in the recognition of PS/PS-OX. A more detailed understanding of the mechanisms that govern the safe removal of apoptotic cells will aid in the design of novel therapeutic strategies in conditions of unscheduled cell death and/or impairment of cell clearance.
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FOOTNOTES |
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To whom correspondence should be addressed: Dept. of Environmental and Occupational Health, University of Pittsburgh, Pittsburgh, PA 15260. Tel.: 412-383-2136; Fax: 412-383-2123; E-mail: kagan{at}pitt.edu.
1 The abbreviations used are: PS, phosphatidylserine; PI, phosphatidylinositol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; cis-PnA, cis-parinaric acid; MCLA, 2-methyl-6-(4-methoxyphenyl)-3,7-dihydroimidazol(1,2-a)pyrazin-3-one hydrochloride; NBD-PS, 1-palmitoyl-2-(6-((7-nitro-2-1,3-benzoxadiazol-4-yl)amino)hexanoyl)-sn-glycero-3-phosphoserine; NBD-PC, 1-palmitoyl-2-(6-((7-nitro-2-1,3-benzoxadiazol-4-yl)amino)hexanoyl)-sn-glycero-3-phosphochilone; PS-OX, oxidized phosphatidylserine; APT, aminophospholipid translocase; PBS, phosphate-buffered saline; NEM, N-ethylmaleimide; ROS, reactive oxygen species; z-VAD-fmk, benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl ketone; HPLC, high performance liquid chromatography; HPTLC, high performance thin layer chromatography; PIPES, 1,4-piperazinediethanesulfonic acid; CHAPS, 3-((3-cholamidopropyl)dimethylammonio)-1-propanesulfonic acid; ANOVA, analysis of variance; AML, acute myelogenous leukemia.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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, control; 
, H2O2 (25 µM x 4); 
, etoposide (50 µM); •, etoposide (50 µM) plus H2O2 (25 µM x 4). Data are mean ± S.E. (n = 3-6); *, p < 0.05 versus control.
