Oxidized Low Density Lipoprotein Induces Apoptosis in Cultured Human Umbilical Vein Endothelial Cells by Common and Unique Mechanisms*

Mariko Harada-ShibaDagger §, Mikio KinoshitaDagger §, Hiroshi Kamido, and Kentaro ShimokadoDagger par

From the Dagger  National Cardiovascular Center Research Institute, 7-1 Fujishirodai 5-chome, Suita, Osaka 565-8565 and the  Department of Medicine, Kurume University School of Medicine, Asahi-Machi, Kurume, Fukuoka 830-0011, Japan

    ABSTRACT
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Oxidized low density lipoprotein (oxLDL) induces apoptosis in vascular cells. To elucidate the mechanisms involved in this apoptosis, we studied the apoptosis-inducing activity in lipid fractions of oxLDL and the roles of two common mechanisms, ceramide generation and the activation of caspases, in apoptosis in human umbilical vein endothelial cells treated with oxLDL. We also studied the effects of antioxidants and cholesterol. oxLDL induced endothelial apoptosis in a time- and dose-dependent fashion. Apoptosis-inducing activity was recovered in the neutral lipid fraction of oxLDL. Various oxysterols in this fraction induced endothelial apoptosis. Neither the phospholipid fraction nor its component lysophosphatidylcholine induced apoptosis. oxLDL induced ceramide accumulation temporarily at 15 min in a dose-dependent fashion. Two inhibitors of acid sphinogomyelinase inhibited both the increase in ceramide and the apoptosis induced by oxLDL. Furthermore, a membrane-permeable ceramide (C2-ceramide) induced endothelial apoptosis. These findings demonstrated that ceramide generation by acid sphingomyelinase is indispensable for the endothelial apoptosis induced by oxLDL. Inhibitors of both caspase-1 and caspase-3 inhibited the apoptosis, suggesting that oxLDL induced apoptosis by activating these cysteine proteases. The antioxidants butylated hydroxytoluene and superoxide dismutase but not catalase inhibited the apoptosis induced by oxLDL or 25-hydroxycholesterol. This suggests not only that superoxide plays an important role but also that a critical interaction between oxLDL and the cell takes place on the outer surface of the membrane, because superoxide dismutase is not membrane-permeable. Exogenous cholesterol also inhibited the apoptosis. Our study demonstrated that neutral lipids in oxLDL induce endothelial apoptosis by activating membrane sphingomyelinase in a superoxide-dependent manner, as well as by activating caspases.

    INTRODUCTION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Oxidized low density lipoprotein (oxLDL)1 is a key substance in atherogenesis (1, 2). oxLDL is generated by auto-oxidation in the presence of transition metals (3, 4), by cell-mediated mechanisms (5-7), and by enzyme-mediated mechanisms (8-11). It induces the early changes of atherosclerosis: the expression of adhesion molecules on endothelial cells (12), a decrease in production of endothelial cell-derived relaxing factor (13) and prostacyclin (14), the transformation of macrophages and smooth muscle cells to foam cells (15), the production of various proinflammatory cytokines and growth factors by almost all vascular cells (16, 17), the proliferation and migration of vascular cells (18-20), the retardation of endothelial regeneration (21), and changes in the balance between procoagulant and anticoagulant activity on the vascular cell surface (22). These changes consequently trigger a series of cellular responses in the arterial wall that result in the formation of atheromatous lesions. oxLDL also affects the later stage of atherosclerosis by its toxicity. oxLDL and its lipid components cause the release of lactic dehydrogenase from cultured vascular smooth muscle cells, endothelial cells, and fibroblasts (6) and decrease the number of these cells (6). This cytotoxicity of oxLDL is one of the factors that make atheromatous plaques unstable and prone to rupture (23).

Recently, the cytotoxicity of oxLDL has been partly attributed to induction of apoptosis. oxLDL induces both the morphological changes and DNA fragmentation characteristic of apoptosis in cultured smooth muscle cells (24), macrophages (24, 25), endothelial cells (26, 27), and lymphoid cells (28). The apoptosis of vascular cells plays a role in both the progression and the regression of atherosclerotic lesions (23, 29-31). However, it is not clear how oxLDL induces apoptosis in endothelial cells and other vascular cells. Different agents, such as tumor necrosis factor-alpha , ionizing radiation, UV radiation, hydrogen peroxide (32), high glucose (33), and growth factor deprivation (34), induce apoptosis in many cell types by both unique and common pathways (35-37). For example, Fas and tumor necrosis factor receptor family members transduce the signal of apoptosis through death domain-containing molecules, such as FADD/MORT1 (37), whereas many other apoptosis-inducing agents do not use molecules with a death domain. On the other hand, almost all known apoptosis-inducing agents share the activation of caspases (formerly called ICE family proteases) (35-37). Typically, caspase-1 (ICE) activates caspase-3 (CPP32/apopain/YAMA), which then cleaves death substrates such as PARP and lamins (37). Ceramide accumulation has been proposed to be a common pathway of apoptosis; most apoptosis-inducing agents induce an accumulation of intrinsic ceramide and a concomitant decrease in sphingomyelin, a precursor of ceramide; C2-C8 ceramides, which are membrane-permeable synthetic analogues of ceramide, activate caspases and induce apoptosis in many cells (35, 38-40). However, some of the increase in ceramide has been suggested to be a result rather than a cause of cell death (40, 41).

In this study, we investigated the mechanisms involved in the endothelial apoptosis induced by oxLDL. We found that ceramide generation, as well as caspase activation, is indispensable for the endothelial apoptosis induced by oxLDL and that some unique mechanisms are also involved in this process.

    Experimental Procedures
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Materials-- The inhibitor for caspase-3 (CPP32/apopain), ac-DEVD-CHO, and the inhibitor for caspase-1 (interleukin-1beta -converting enzyme), ac-YVAD-CHO, were purchased from Peptide Institute, Inc. (Osaka, Japan). Cholesterol (5-cholesten-3beta -ol), 7beta -hydroxycholesterol (5-cholesten-3beta ,7beta -diol), 25-hydroxycholesterol (5-cholesten-3beta ,25-diol), cholesterol-5alpha ,6alpha -epoxide (5alpha ,6alpha -epoxycholestan-3beta -ol), 3beta ,5alpha ,6beta -trihydroxycholestane (cholestane-3beta ,5alpha ,6beta -triol), L-lysophosphatidyl-choline palmitoyl (lysoPC), butylated hydroxytoluene (BHT), superoxide dismutase (SOD) (EC 1.15.1.1; bovine erythrocytes), and catalase (EC 1.11.1.6; bovine liver) were purchased from Sigma, and 7-keto-cholesterol (5-cholesten-3beta -ol-7-one) was from Makor Chemical Ltd. (Jerusalem, Israel). C2-ceramide (N-acetylsphingosine) was from BIOMOL Research Laboratory (Plymouth Meeting, PA).

Cells-- Human umbilical vein endothelial cells (HUVECs) (42) were seeded at a density of 5 × 104/cm2 and cultured in Dulbecco's modified Eagle's medium (DMEM) (Nissui Pharmaceutical Co., Tokyo, Japan) supplemented with penicillin (100 units/ml), streptomycin (100 µg/ml), 20% (v/v) fetal calf serum (FCS) (Life Technologies, Inc.), and 10 ng/ml recombinant human basic fibroblast growth factor (bFGF) (Pepro Tech Inc., Rocky Hill, NJ) for 24 h before the medium was changed to DMEM supplemented with 0.1% bovine serum albumin (BSA), 10 ng/ml bFGF, and various concentrations of oxLDL. The number of cells in this confluent culture did not change significantly in the serum-free medium for the next 48 h unless oxLDL was added. The cells were used for the experiments between passages 2 and 7.

Preparation of Modified LDL-- LDL (1.019 < d < 1.063) was prepared from normal human serum by sequential ultracentrifugation as described previously (43). Fifteen ml of DMEM was mixed with LDL (500 µg/ml) and Cu(II) sulfate (20 µM), and incubated with a confluent culture of HUVECs in a 15-cm dish at 37 °C, 5% CO2, and 100% humidity for 48 h. The medium was centrifuged at 3500 rpm for 10 min, sterilized with a 0.45 µm Millipore filter, and used as oxLDL. The incubation of LDL in DMEM without either HUVECs or Cu(II) sulfate did not cause significant oxidation of LDL. Acetylated LDL was prepared as reported previously (44). All procedures were carried out under sterile conditions. The endotoxin was measured by a radioimmunoassay (Endospec SP test, Seikagaku-kogyo, Tokyo, Japan) and was under the detection limit in all native and oxLDL preparations. The oxidation of LDL was evaluated by agarose gel electrophoresis (Universal gel, Ciba Corning Diagnostic Corp., Alameda, CA) and by the measurement of thiobarbituric acid reacting substance (6). oxLDL had a thiobarbituric acid reacting substance value of 23.2 nmol of malondialdehyde/mg of protein (the mean ± S.E. of four separate preparations), whereas native and acetylated LDL each had no detectable thiobarbituric acid reacting substance. The oxLDL and acetylated LDL had 2-3-fold higher Rf values on agarose gel electrophoresis compared with the native LDL. Gas chromatography (45) revealed that a typical preparation of ox LDL contained 4.0 µg of 7alpha -hydroxycholesterol/mg of LDL protein, 13.1 µg of epoxide/mg of LDL protein, 14.9 µg of 7-keto-cholesterol/mg of LDL protein, and 1.1 mg of unidentified oxycholesterol/mg of LDL protein.

Quantitative Analysis of Apoptotic Cells-- HUVECs were seeded at 5 × 104 cells/well in gelatin-coated 8-well chamber slides (Nunc, Inc., Naperville, IL) and cultured for 24 h before the medium was changed to DMEM containing 0.1% BSA, 10 ng/ml bFGF, and the indicated concentration of oxLDL. The cells were then cultured in the presence of oxLDL for the indicated periods, followed by fixation in 3% paraformaldehyde in phosphate-buffered saline for 20 min and staining with a solution of 4',6-diamidino-2-phenylindole (10 mM Tris-HCl pH 7.4, 10 mM EDTA, 100 mM NaCl, 500 ng/ml 4',6-diamidino-2-phenylindole) for 10 min at room temperature (46). The number of apoptotic cells was counted in nine high power fields under a fluorescent microscope (approximately 1000-2000 cells/well). The percentage of apoptotic cells was calculated as the number of apoptotic cells/number of total cells × 100%. Each experiment was conducted in triplicate and repeated at least twice.

Fractionation of Lipids-- Total lipid was extracted from oxLDL with chloroform/methanol and resolved by TLC (Silica gel 60, Merk, Darmstadt, Germany) using heptane/isopropyl ether/acetic acid (60:40:4) as the developing solvent. Neutral lipids and phospholipids were extracted from the TLC with chloroform/methanol (1:1).

Measurement of Ceramide-- The cellular ceramide content was measured by an Escherichia coli diacylgylcerol kinase assay (47, 48). Briefly, total lipids were extracted from HUVECs (2 × 106 cells) by chloroform/methanol (1:1) and dried under N2 gas. The lipid was solubilized by sonication for 2 min into 20 µl of aqueous 7.5% n-octyl-beta -D-glucopyranoside (Dojindo), 5 mM cardiolipin (Sigma), and 1 mM diethylenetriaminepentaacetic acid (Dojindo, Kumamoto, Japan). The solubilized lipid was mixed with 20 µl of reaction buffer (250 mM Tris-HCl, 500 mM NaCl, 10 mM EGTA, 25 mM MgCl2, pH 7.0), 10 µl of 1 mg/ml E. coli diacylglycerol kinase (Calbiochem), 20 µl of 10 mM ATP (Sigma), and 5 µl of [-32P]ATP (111 TBq/mmol, DuPont). The reaction was allowed to proceed at room temperature for 40 min and then was stopped by the addition of 1 ml chloroform/methanol/1 N HCl (100:100:1, v/v), 340 µl of buffered saline solution, and 60 µl of 100 mM EDTA. The lower organic phase was dried under N2. Labeled ceramide was resolved by TLC using chloroform/acetone/methanol/acetic acid/water (10:4:2:2:1, v/v) as the developing solvent. The autoradiogram was analyzed by a Fuji Bas 2000 Bioimaging analyzer, and the ceramide content was determined with a standard curve of 0-1.7 nmol of ceramides (type III, Sigma). The ceramide content was standardized with total cellular phospholipid determined by a commercial kit (Wako Phospholipid Test, Wako Pure Chemicals, Osaka, Japan).

Statistical Analysis-- Statistical analysis was conducted with Student's t test. Differences were considered significant when probability values less than 5% were obtained.

    Results
Top
Abstract
Introduction
Procedures
Results
Discussion
References

oxLDL Induces Apoptosis in HUVECs-- oxLDL induced the apoptosis in a time- and dose-dependent manner (Fig. 1). oxLDL induced significant apoptosis in HUVECs as early as at 12 h of incubation, and the number of apoptotic cells increased up until 48 h of incubation (Figs. 1 and 2). Twelve to 18% of the HUVECs became apoptotic after 48 h incubation with 25 µg of protein/ml of oxLDL. This value is approximately the same as that obtained with the deprivation of both serum and bFGF (Fig. 2A). At higher concentrations of oxLDL, the percentage of apoptotic cells decreased (Fig. 2B), as did the total cell number (data not shown). After 72 h, the HUVECs became apoptotic regardless of the presence or absence of oxLDL, probably due to serum starvation, and the effect of oxLDL became less prominent (data not shown). Incubation with native LDL or acetylated LDL did not induce apoptosis beyond the control level (less than 5% of total cells) (data not shown). These findings indicated that oxLDL induces apoptosis in cultured human endothelial cells.


View larger version (85K):
[in this window]
[in a new window]
 
Fig. 1.   Endothelial apoptosis induced by oxLDL. HUVECs were cultured in 8-well Lab-Tek chamber slides at 5 × 104 cells/cm2 in DMEM containing 20% FCS and 10 ng/ml bFGF. After 24 h, the medium was changed to DMEM containing 0.1% BSA and 10 ng/ml bFGF, cultured further for 48 h in the presence or absence of oxLDL (25 µg of protein/ml), and then stained with 4',6-diamidino-2-phenylindole as described under "Experimental Procedures." A, cells cultured with oxLDL; B, the same cells with a higher magnification; C, cells without oxLDL. Bar indicates 10 µm.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 2.   Time- and dose-dependent induction of apoptosis in HUVECs by oxLDL. A, time dependence. HUVECs were seeded as described in the legend to Fig. 1, and cultured in the presence (black-square) or absence (black-triangle) of oxLDL (25 µg of protein/ml) for the indicated periods. As a positive control (open circle ), cells were cultured in the absence of bFGF. The number of apoptotic cells was counted in nine high power fields under a fluorescent microscope. Each point represents the mean ± S.E. of quintuple samples for the negative control and oxLDL groups and of quadruplicate samples for the positive control. B, dose dependence. HUVECs were seeded and incubated with indicated concentrations of oxLDL for 48 h. As a positive control, cells were cultured in the absence of bFGF. The values shown are the mean ± S.E. of quadruplicate samples.

Lipid Components in oxLDL Induce Apoptosis-- The cytotoxicity of oxLDL has been attributed to its lipid components (49, 50). To examine whether the apoptotic activity of oxLDL is also due to the lipid fractions, oxLDL was extracted with organic solvent, and neutral lipids and phospholipids were separated by TLC. All of the apoptosis-inducing activity of oxLDL was recovered in the total lipid fraction (Table I). More than 90% of the activity was recovered in the neutral lipid fraction, and no significant activity was recovered in the phospholipid fraction. Gas chromatography revealed that the major components of the neutral lipid fraction generated in oxLDL were a series of oxysterols, such as 7-ketocholesterol and 7-hydroxycholesterol (see preparation of modified LDL under "Experimental Procedures"). We therefore investigated the apoptosis-inducing activity of various oxysterols found in oxLDL. All oxysterols examined induced apoptosis, to various degrees; significant apoptosis was induced by 5 µg/ml 7-keto-cholesterol, 25-hydroxycholesterol, or triol and by 20 µg/ml 7-hydroxycholesterol or epoxide (Table II). Oxysterol tended to induce apoptosis to a greater degree compared with oxLDL. Cholesterol did not induce apoptosis at 20 µg/ml. LysoPC did not induce significant endothelial apoptosis (Table II). These data suggest that neutral lipids, such as oxysterols, are responsible for the apoptosis-inducing activity of oxLDL.

                              
View this table:
[in this window]
[in a new window]
 
Table I
Apoptosis-inducing activity of various lipid fractions of oxLDL
oxLDL was fractionated as described under "Experimental Procedures." Each fraction was tested at a concentration of lipid equivalent to 25 µg of protein/ml of oxLDL. Values are the mean ± S.E. of six samples.

                              
View this table:
[in this window]
[in a new window]
 
Table II
Oxysterols induce apoptosis in HUVECs
The cells were seeded as described in the legend to Fig. 1. On day 2, the medium was changed to DMEM containing the indicated concentration of oxysterol. After 48 h, apoptotic cells were counted. Each value represents the mean ± S.E. of quadruplicate samples. 7beta -OH-cholesterol, 7beta -hydroxycholesterol; 7-keto-cholesterol, 5-cholesten-3beta -ol-7-one; epoxide, cholesterol 5alpha , 6alpha -epoxide; 25-OH-cholesterol, 25-hydroxycholesterol; Triol, cholestane-3beta ,5alpha ,6alpha -triol. NS, not significant.

oxLDL Induces Apoptosis by Generating Ceramide-- Although ceramide has been implicated in the apoptosis induced by a variety of agents, its precise role is still controversial (35, 40). Therefore, we studied whether ceramide was involved in oxLDL-induced apoptosis. First, we examined whether oxLDL affected the cellular ceramide content. Ceramides were detected as two major spots on TLC, as reported previously (Fig. 3A) (48). oxLDL (0-50 µg/ml) increased the cellular ceramide transiently at 15 min in a dose-dependent fashion (Fig. 3). After 24 h of oxLDL treatment, the cellular ceramide increased again to a similar extent (data not shown). An oxysterol, 7-keto-cholesterol (5 µg/ml), also stimulated ceramide generation at 15 min (3.2-fold increase; average of two independent experiments). Second, we studied whether the inhibition of ceramide generation prevented endothelial apoptosis. Two different inhibitors of sphingomyelinase, desipramine and chlorpromazine (51), inhibited the oxLDL-induced apoptosis completely (Fig. 4A). At the same concentration, these inhibitors inhibited the ceramide generation induced by oxLDL (Fig. 4B). Third, we studied whether exogenous ceramide induced apoptosis in our experimental conditions. In accordance with a previous report with exogenous C6-ceramide (52), C2-ceramide, a membrane-permeable form of ceramide, induced endothelial apoptosis (Table III). These findings demonstrated that the increase in cellular ceramide is indispensable for the induction of endothelial apoptosis by oxLDL.


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 3.   oxLDL increases the cellular ceramide content. A and B, time course. HUVECs were seeded at 5 × 104 cm2 in DMEM supplemented with 20% FCS and 10 ng/ml bFGF and cultured for 24 h. The medium was changed to DMEM supplemented with 0.1% BSA, 10 ng/ml bFGF, and oxLDL (25 µg of protein/ml). At the indicated times, the total cellular lipid was extracted, and the ceramide content was measured as described under "Experimental Procedures" (A). The standard (std) was 1.7 nmol of ceramide. Quantitative analysis was conducted with BAS200 image analyzer (B). Each data point indicates the mean ± S.E. of values obtained from three independent experiments conducted in duplicate. Ceramide content of time 0 was 2.9 ± 0.7 mol/104 mol of phospholipid (the mean ± S.E. of six samples). C, dose dependence. HUVECs were cultured in the presence of various concentrations of oxLDL for 15 min. Ceramide content of the sample containing no oxLDL was 2.1 ± 0.6 mol/104 mol of phospholipid (the mean ± S.E. of six samples).


View larger version (30K):
[in this window]
[in a new window]
 
Fig. 4.   Effect of sphingomyelinase inhibitors on ceramide generation and apoptosis. A sphingomyelinase inhibitor was added to the culture 22 h after seeding. After 2 h, the medium was changed to DMEM supplemented with 0.1% BSA, 10 ng/ml bFGF, oxLDL (25 µg of protein/ml), and the inhibitor. The percentage of apoptotic cells (A) was determined after 48 h, and ceramide content (B) was determined at 15 min. Each column represents the mean ± S.E. of three independent experiments conducted in duplicate.

                              
View this table:
[in this window]
[in a new window]
 
Table III
Effects of C2-ceramide on endothelial apoptosis
HUVECs were seeded as described in the legend to Fig. 1 and cultured in the presence of C2-ceramide for 48 h. Values are the mean ± S.E. of three independent experiments conducted in duplicate.

Other Mechanisms Involved in oxLDL-induced Apoptosis-- To elucidate other mechanisms involved in this endothelial apoptosis, we studied the effects of inhibitors of the enzymes involved in apoptosis, antioxidants, and cholesterol. Although most apoptosis-inducing agents activate caspases (35-37), there are some exceptions (53). Therefore, we tried to confirm whether oxLDL induces apoptosis by activating caspases. The inhibitor of caspase-1 (ac-YVAD-CHO) and the inhibitor of caspase-3 (ac-DEVD-CHO) each inhibited apoptosis (Fig. 5), confirming that oxLDL induces apoptosis by activating caspases, as do most other apoptosis-inducing factors. Because oxLDL could be further oxidized in the presence of HUVECs and the radicals generated during this process could induce apoptosis, we studied the effects of antioxidants on endothelial apoptosis. Among the antioxidants, SOD and BHT inhibited the apoptosis but catalase had no effect on oxLDL-induced apoptosis (Fig. 6), suggesting that superoxide, but not hydrogen peroxide, is responsible for the ability of oxLDL to induce of apoptosis. Similarly, both SOD and BHT inhibited the apoptosis-inducing activity of 25-hydroxycholesterol (data not shown). A recent report that caspase-3 cleaves sterol regulatory element binding proteins and may up-regulate cholesterol synthesis (54) indicates a close relationship between apoptosis and cholesterol metabolism, prompting us to study the effect of cholesterol on endothelial apoptosis. Exogenous cholesterol potently inhibited the apoptosis induced by both oxLDL and oxysterols (Fig. 7).


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 5.   Effects of caspase inhibitors on oxLDL-induced apoptosis. HUVECs were seeded as described in the legend to Fig. 1. The cells were cultured in DMEM supplemented with 0.1% BSA, 10 ng/ml bFGF, and oxLDL (25 µg of protein/ml) in the presence or absence of an inhibitor for 48 h. The values shown are the mean ± S.E. of quadruplicate samples.


View larger version (15K):
[in this window]
[in a new window]
 
Fig. 6.   Effects of antioxidants on oxLDL-induced apoptosis. HUVECs were seeded as described in the legend to Fig. 1. The cells were cultured in DMEM supplemented with 0.1% BSA, 10 ng/ml bFGF, and oxLDL (25 µg of protein/ml) in the presence or absence of antioxidants for 48 h. For antioxidants, 50 µM of BHT, 100 µg/ml SOD, and 100 µg/ml catalase were used. The vehicle of these antioxidants (ethanol and phosphate-buffered saline) alone did not affect the apoptosis. BHT was dissolved in ethanol, and SOD and catalase were dissolved in phosphate-buffered saline. The values shown are the mean ± S.E. of quadruplicate samples.


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 7.   Effect of cholesterol on apoptosis induced by oxLDL or oxysterols. HUVECs were seeded as described in the legend to Fig. 1. After 24 h, the medium was changed to DMEM supplemented with 0.1% BSA, 10 ng/ml bFGF, and oxLDL (25 µg of protein/ml) or oxysterols. 25-OH Chol, 25-hydroxycholesterol (5 µg/ml), Triol, cholestane-3,5,6-triol (5 µg/ml). Cells were cultured in the presence or absence of 50 µg/ml cholesterol for 48 h. Each column represents the mean ± S.E. of quadruplicate values.

    DISCUSSION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

In this study, we observed that oxLDL induces apoptosis in cultured human umbilical vein endothelial cells by both common and unique mechanisms. The common mechanisms include the accumulation of intrinsic ceramide and the activation of caspases. A unique mechanism underlying oxLDL-induced apoptosis is the involvement of superoxide. We also found that cholesterol inhibits oxLDL-induced apoptosis, although its precise mode of action is unknown.

Previous reports indicated that oxysterols and oxLDL could induce apoptosis in endothelial cells (26, 27, 55). We confirmed these observations and showed that oxysterols (and potentially some other lipids) in the neutral lipid fraction of oxLDL account for the apoptosis-inducing activity of oxLDL. Most apoptosis-inducing activity was recovered in the neutral lipid fraction that contains a series of oxysterols, as detected by gas chromatography. Oxysterols potently induced the apoptosis at concentrations comparable to those in oxLDL (Table II). Although lysoPC in oxLDL has some important biological effects (12), neither the phospholipid fraction nor isolated lysoPC induced endothelial apoptosis in the present study.

One of the mechanisms involved in this apoptosis is the accumulation of intrinsic ceramide. This conclusion was drawn from three sets of observations. First, oxLDL increased the cellular ceramide content, as reported with other apoptosis-inducing agents such as tumor necrosis factor-alpha , anti-FAS antibody, and ionizing radiation. This ceramide generation did not need sphingomyelin in oxLDL as a substrate of sphingomyelinase (56) because an oxysterol also increased the cellular ceramide content (see "Results"). Second, two different nonspecific inhibitors of acid sphingomyelinase completely inhibited both ceramide generation and the apoptosis. These inhibitors do not inhibit other lysosomal enzymes or neutral sphingomyelinase (51). Third, a membrane-permeable homologue of ceramide, C2-ceramide, induced apoptosis in HUVECs at concentrations comparable to those of intrinsic ceramide. A similar membrane-permeable ceramide, C6-ceramide, was reported to induce apoptosis in HUVECs (53). Although accumulating data suggest that the increase in ceramide is a common mechanism among various apoptosis-inducing agents (35-37), there are some concerns about the role of ceramide in apoptosis. For example, although Fas-induced apoptosis is independent of transcription or translation, C2 ceramide-induced apoptosis is inhibited by blocking the transcription of AT-1, suggesting that FAS, although it increases ceramide generation, induces apoptosis by a mechanism unrelated to this increase in ceramide. The inhibition of caspase-1 blocked both the ceramide generation and apoptosis induced by REAPER, suggesting that ceramide generation is a result rather than a cause of the activation of caspase-1, a key enzyme of apoptosis (41). Our finding that cellular ceramide increased temporarily at 15 min and again at 24 h after stimulation with oxLDL suggests that cellular ceramide increases as both a cause and a result of apoptosis. The increase of ceramide at 15 min is clearly a cause of apoptosis, because the inhibitor of sphingomyelinase blocked both ceramide generation at 15 min and apoptosis at 6 h later. The increase of ceramide at 15 min could not be a result of apoptosis, because no apoptosis was detected at that point. The significance of the increase in ceramide at early time points agrees well with a recent report concerning the temporal profile of apoptosis in a cell-free system, in which ceramide caused fragmentation of nuclei only when it was added in the first 90 min of the time course (57). Interleukin-1beta induces E-selectin in HUVECs through a transient ceramide generation similar to that observed in the present study (48). Contrary to the ceramide increase at 15 min, the ceramide increase after 24 h could be a result of apoptosis because significant apoptosis had occurred by this time.

Another controversy regarding the ceramide pathway of apoptosis centers on the enzymes involved in the ceramide generation. Tumor necrosis factor-alpha and Fas ligand activate acid sphingomyelinase in many cell types (58-61), whereas ionizing radiation induces apoptosis in bovine aortic endothelial cells by activating neutral sphingomyelinase (62). Ceramide synthase accounts for ceramide generation in daunorubicin-induced apoptosis (63). We employed two sphingomyelinase inhibitors that inhibit acid sphingomyelinase (64, 65) and demonstrated that acid sphingomyelinase is responsible for the ceramide generation in oxLDL-induced apoptosis. Acid sphingomyelinase was recently found to be localized not only in the lysosome but also in association with the caveola, a membrane domain that can undergo an internalization cycle (66). Most membrane sphingomyelin and ceramide are also localized in the caveola, and interleukin-1 can activate acid sphingomyelinase and increase ceramide in the caveola (66). As discussed later, our data suggest that a critical interaction between oxLDL and endothelial cells takes place on the outer surface of the cell. Taken together, the present and above-mentioned findings suggest that acid sphingomyelinase in the caveola plays a role in the ceramide generation and apoptosis induced by oxLDL.

Another common mechanism involved in this apoptosis is the activation of caspases (35-37). Typically, apoptotic agents activate caspase-1 and caspase-3 sequentially. However, there are many alternative caspases to cleave death substrates (37), or even a pathway independent of classical interleukin 1beta -converting enzyme (53). oxLDL has been found to activate caspase-3 (CPP32) and induce apoptosis in HUVECs (27). Our findings with specific inhibitors confirm that both caspase-1 and caspase-3 are involved in the endothelial apoptosis induced by oxLDL.

We identified a unique mechanism underlying the oxLDL-induced endothelial apoptosis: the involvement of superoxide. Our finding that SOD but not catalase inhibits oxLDL-induced apoptosis indicates that superoxide play an important role in the apoptosis. Oxygen radical has been proposed as a target of bcl-2, but now its role is questioned based on observations that apoptosis is induced even under an anaerobic condition (67, 68). An important difference between previous reports and ours is that superoxide plays a role outside the cell in our experimental system, whereas intracellular radical-associated mechanisms have been studied as intracellular mechanisms of apoptosis. Because SOD is not membrane-permeable, the site of action of this enzyme would expected to be outside the plasma membrane. Therefore, although oxLDL and oxysterols can get into the cell, critical interactions between these lipids and the cell are thought to take place on the outer surface of the cell. It is also noteworthy that LDL modified with reactive oxygen species is not sufficient and needs the further involvement of reactive oxygen species to induce endothelial apoptosis. This is in contrast with a previous report that toxicity of oxLDL is not blocked by antioxidants once LDL is oxidized (3). Our findings suggest that oxysterols (and potentially other lipid components) in oxLDL play a role in the propagation of the chain reaction initiated by superoxide to induce apoptosis.

An interesting finding of our study is that exogenous cholesterol inhibited the apoptosis induced by oxLDL or oxysterols. Although previous investigators have pointed out a potential relationship between cholesterol metabolism and apoptosis (54), direct evidence of an antiapoptotic effect of cholesterol has not been reported. Based on an observation that caspase-3 was found to cleave sterol regulatory element binding proteins and to potentially up-regulate cholesterol metabolism (54), it was suggested that cholesterol is needed to maintain membrane integrity during apoptosis. If so, cholesterol could inhibit apoptosis by reducing the intrinsic cholesterol supply that is needed for apoptosis. However, this is not likely, because hydroxycholesterols are more potent negative regulators for HMG-CoA reductase, whereas hydroxycholesterols induce rather than inhibit apoptosis. Our findings that oxLDL activates caspase-3 and that cholesterol inhibits apoptosis suggest that caspase-3 up-regulates cholesterol synthesis as a negative feedback mechanism to prevent apoptosis, whereas it cleaves the death substrate to induce apoptosis. Although the precise mode of action of cholesterol remains to be elucidated, cholesterol can inhibit sphingomyelin degradation and therefore can potentially inhibit the ceramide pathway of apoptosis.

In summary, we reported mechanisms involved in the oxLDL-induced apoptosis in vascular endothelial cells. Our data suggest that neutral lipids, such as oxysterols, in oxLDL activate acid sphingomyelinase on the outer membrane of the cell in a superoxide-dependent manner to generate ceramide, which eventually activates caspases.

    ACKNOWLEDGEMENTS

We thank Dr. Atsushi Masamune for helpful advice on ceramide measurement, Dr. Shigeko Takaichi for electron microscopy, and Dr. Hisayuki Matsuo for encouragement during this project.

    FOOTNOTES

* This study was supported by grants from the Science and Technology Agency, from the Ministry of Education, Science and Culture, and from the Ministry of Health and Welfare.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§ The first two authors contributed equally to this work.

par To whom correspondence should be addressed. Tel.: 81-6-833-5012, ext. 2590; Fax: 81-6-872-7485; E-mail: kshimoka{at}res.ncvc.go.jp.

1 The abbreviations used are: oxLDL, oxidized low density lipoprotein; BHT, butylated hydroxytoluene; SOD, superoxide dismutase; HUVEC, human umbilical vein endothelial cell; DMEM, Dulbecco's modified Eagle's medium; lysoPC, lysophosphatidylcholine; bFGF, basic fibroblast growth factor; BSA, bovine serum albumin.

    REFERENCES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

  1. Steinberg, D., Parthasarathy, S., Carew, T. E., Khoo, J. C., and Witztum, J. L. (1989) New Engl. J. Med. 320, 915-924[Medline] [Order article via Infotrieve]
  2. Ross, R. (1993) Nature 362, 801-809[CrossRef][Medline] [Order article via Infotrieve]
  3. Morel, D. W., Hessler, J. R., and Chisolm, G. M. (1983) J. Lipid Res. 24, 1070-1076[Abstract]
  4. Lamb, D. J., Michinson, M. J., and Leake, D. S. (1995) FEBS Lett. 374, 12-16[CrossRef][Medline] [Order article via Infotrieve]
  5. Steinbrecher, U. P., Parthasarathy, S., Leake, D. S., Witztum, J. L., and Steinberg, D (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 3883-3887[Abstract]
  6. Morel, D. W., DiCorleto, P. E., and Chisolm, G. M. (1984) Arteriosclerosis 4, 357-364[Abstract]
  7. Hiramatsu, K., Rosen, H., Heinecke, J. W., Wolfbauer, G., and Chait, A. (1987) Arteriosclerosis 7, 55-60[Abstract]
  8. Ehrenwald, E., Chisolm, G. M., and Fox, P. L. (1994) J. Clin. Invest. 93, 1493-1501[Medline] [Order article via Infotrieve]
  9. Parthasarathy, S., Steinbrecher, U. P., Barnett, J., Witztum, J. L., and Steinberg, D. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 3000-3004[Abstract]
  10. Daugherty, A., Dunn, J. L., Rateri, D. L., and Heinecke, J. W. (1994) J. Clin. Invest. 94, 437-444[Medline] [Order article via Infotrieve]
  11. Yla-Herttuala, S., Luoma, J., Viita, H., Hiltunen, T., Sisto, T., and Nikkari, T. (1995) J. Clin. Invest. 95, 2692-2698[Medline] [Order article via Infotrieve]
  12. Kume, N., Cybulsky, M. I., and Gimbrone, M. A., Jr. (1992) J. Clin. Invest. 90, 1138-1144[Medline] [Order article via Infotrieve]
  13. Mangin, E. L., Kugiyama, K., Nguy, J. H., Kerns, S. A., and Henry, P. D. (1993) Circ. Res. 72, 161-166[Abstract]
  14. Thorin, E., Hamilton, C. A., Dominiczak, M. H., and Reid, J. L. (1994) Arterioscler. Thromb. 14, 453-459[Abstract]
  15. Henriksen, T., Mahoney, E. M., and Steinberg, D. (1981) Proc. Natl. Acad. Sci. U. S. A. 78, 6499-6503[Abstract]
  16. Kume, N., and Gimbrone, M. A., Jr. (1994) J. Clin. Invest. 93, 907-911[Medline] [Order article via Infotrieve]
  17. Nakano, T., Raines, E. W., Abraham, J. A., Klagsbrun, M., and Ross, R. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 1069-1073[Abstract]
  18. Quinn, M. T., Parthasarathy, S., and Steinberg, D. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 2805-2809[Abstract]
  19. Auge, N., Andrieu, N., Negre-Salvayre, A., Thiers, J.-C., Levade, T., and Salvayre, R. (1996) J. Biol. Chem. 271, 19251-19255[Abstract/Free Full Text]
  20. Yui, S., Sasaki, T., Miyazaki, A., Horiuchi, S., and Yamazaki, M. (1993) Arterioscler. Thromb. 13, 331-337[Abstract]
  21. Murugesan, G., and Fox, P. L. (1996) J. Clin. Invest. 97, 2736-2744[Abstract/Free Full Text]
  22. Ishii, H., Kizaki, K., Horie, S., and Kazama, M. (1996) J. Biol. Chem. 271, 8458-8465[Abstract/Free Full Text]
  23. Bjorkerud, S., and Bjorkerud, B. (1996) Am. J. Pathol. 149, 367-380[Abstract]
  24. Bjorkerud, B., and Bjorkerud, S. (1996) Arterioscler. Thromb. Vasc. Biol. 16, 416-424[Abstract/Free Full Text]
  25. Reid, V. C., Mitchinson, M. J., and Skepper, J. N. (1993) J. Pathol. 171, 321-328[Medline] [Order article via Infotrieve]
  26. Escargueil-Blanc, I., Meilhac, O., Pieraggi, M.-T., Arnal, J.-F., Salvayre, R., and Negre-Salvayre, A. (1997) Arterioscler. Thromb. Vasc. Biol. 17, 331-339[Abstract/Free Full Text]
  27. Dimmeler, S., Haendeler, J., Galle, J., and Zeiheer, A. M. (1997) Circulation 95, 1760-1763[Abstract/Free Full Text]
  28. Escargueil-Blanc, I., Salvayre, R., and Negre-Salvayre, A. (1994) FASEB J. 8, 1075-1080[Abstract/Free Full Text]
  29. Isner, J. M., Kearney, M., Bortman, S., and Passeri, J. (1995) Circulation 91, 2703-2711[Abstract/Free Full Text]
  30. Han, D. K. M., Haudenschild, C. C., Hong, M. K., Tinkle, B. T., Leon, M. B., and Liau, G. (1995) Am. J. Pathol. 147, 267-277[Abstract]
  31. Bochaton-Piallat, M.-L., Gabbiani, F., Redard, M., Desmouliere, A., and Gabbiani, G. (1995) Am. J. Pathol. 146, 1059-1064[Abstract]
  32. de Bono, D. P., and Yang, W. D. (1995) Atherosclerosis 114, 235-245[CrossRef][Medline] [Order article via Infotrieve]
  33. Baumgartner-Parzer, S. M., Wagner, L., Pettermann, M., Grillari, J., Gessl, A., and Waldhausl, W. (1995) Diabetes 44, 1323-1327[Abstract]
  34. Araki, S., Shimada, Y., Kaji, K., and Hayashi, H. (1990) Biochem. Biophys. Res. Commun. 168, 1194-1200[Medline] [Order article via Infotrieve]
  35. Hale, A. J., Smith, C. A., Sutherland, L. C., Stoneman, V. E. A., Longthorne, V. L., Culhane, A. C., and Williams, G. T. (1996) Eur. J. Biochem. 236, 1-25[Abstract]
  36. Fraser, A., and Evan, G. (1996) Cell 85, 781-784[Medline] [Order article via Infotrieve]
  37. Nagata, S. (1997) Cell 88, 355-365[Medline] [Order article via Infotrieve]
  38. Obeid, L. M., Linardic, C. M., Karolak, L. A., and Hannun, Y. A. (1993) Science 259, 1769-1771[Medline] [Order article via Infotrieve]
  39. Jarvis, W. D., Kolesnick, R. N., Fornari, F. A., Traylor, R. S., Gewirtz, D. A., and Grant, S. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 73-77[Abstract]
  40. Hannun, Y. A. (1996) Science 274, 1855-1859[Abstract/Free Full Text]
  41. Pronk, G. J., Ramer, K., Amiri, P., and Williams, L. T. (1996) Science 271, 808-810[Abstract]
  42. Jaffe, E. A., Nachman, R. L., Becker, C. G., and Minick, C. R. (1973) J. Clin. Invest. 52, 2745-2756[Medline] [Order article via Infotrieve]
  43. Havel, R., Eder, H., and Braigon, J. (1955) J. Clin. Invest. 39, 1345-1363
  44. Brown, M., Goldstein, J., Krieger, M., Ho, Y., and Anderson, R. (1979) J. Cell Biol. 82, 597-613[Abstract]
  45. Kamido, H., Kuksis, A., Marai, L., and Myher, J. J. (1993) Lipids 28, 331-336[Medline] [Order article via Infotrieve]
  46. Shimokado, K., Umezawa, K., and Ogata, J. (1995) Exp. Cell Res. 220, 266-273[CrossRef][Medline] [Order article via Infotrieve]
  47. Van Veldhoben, P. P., Bishop, W., R., Yurivich, D., A., and Bell, R., M. (1995) Biochem. Mol. Biol. Int. 36, 21-30[Medline] [Order article via Infotrieve]
  48. Masamune, A., Igarashi, Y., and Hakomori, S. (1996) J. Biol. Chem. 271, 9368-9375[Abstract/Free Full Text]
  49. Chisolm, G. M., Ma, G., Irwin, K. C., Martin, L. L., Gunderson, K. G., Linberg, L. F., Morel, D. W., and DiCorleto, P. E. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 11452-11456[Abstract/Free Full Text]
  50. Sevanian, A., Hodis, H. N., Hwang, J., McLeod, L. L., and Peterson, H. (1995) J. Lipid Res. 36, 1971-1986[Abstract]
  51. Masson, M., Spezzatti, B., Chapman, J., Battisti, C., and Baumann, N. (1992) J. Neurosci. Res. 31, 84-88[Medline] [Order article via Infotrieve]
  52. Slowik, M. R., De Luca, L. G., Min, W., and Pober, J. S. (1996) Circ. Res. 79, 736-747[Abstract/Free Full Text]
  53. Woodle, E., Smith, D., Bluestone, J., Kirkman, W., Green, D., and Skowronski, E. (1997) J. Immunol. 158, 2156-2164[Abstract]
  54. Wang, X., Zelenski, N. G., Yang, J., Sakai, J., Brown, M. S., and Goldstein, J. L. (1996) EMBO J. 15, 1012-1020[Abstract]
  55. Lizard, G., Deckert, V., Dubrez, L., Moisant, M., Gambert, P., and Lagrost, L. (1996) Am. J. Pathol. 148, 1625-1638[Abstract]
  56. Kinscherf, R., Claus, R., Deigner, H., P., Nauen, O., Gehrke, C., Hermetter, A., Russwurm, S., Daniel, V., Hack, V., and Metz, J. (1997) FEBS Lett. 405, 55-59[CrossRef][Medline] [Order article via Infotrieve]
  57. Farschon, D. M., Couture, C., Mustelin, T., and Newmeyer, D. D. (1997) J. Cell Biol. 137, 1117-1125[Abstract/Free Full Text]
  58. Cifone, M. G., DeMaria, R., Roncaioli, P., Rippo, M. R., Azuma, M., Lanier, L. L., Santoni, A., and Testi, R. (1993) J. Exp. Med. 177, 1547-1552
  59. Cifone, M. G., Roncaioli, P., De Maria, R., Camarda, G., Santoni, A., Ruberti, G., and Testi, R. (1995) EMBO J. 14, 5859-5868[Abstract]
  60. Higuchi, M., Singh, S., Jaffrezou, J.-P., and Aggarwal, B. B. (1996) J. Immunol. 156, 297-304[Abstract]
  61. Santana, P., Pena, L. A., Haimovitz-Friedman, A., Martin, S., Green, D., McLoughlin, M., Cordon-Cardo, C., Schuchman, E. H., Fuks, Z., and Kolesnick, R. (1996) Cell 86, 189-199[Medline] [Order article via Infotrieve]
  62. Haimovitz-Friedman, A., Kan, C.-C., Ehleiter, D., Persaud, R. S., McLoughlin, M., Fuks, Z., and Kolesnick, R. N. (1994) J. Exp. Med. 180, 525-535[Abstract]
  63. Bose, R., Verheij, M., Haimovitz-Friedman, A., Scotto, K., Fuks, Z., and Kolesnick, R. (1995) Cell 82, 405-414[Medline] [Order article via Infotrieve]
  64. Albouz, S., Le Saux, F., Wenger, D., Hauw, J. J., and Baumann, N. (1986) Life Sci. 38, 357-363[CrossRef][Medline] [Order article via Infotrieve]
  65. Lister, M. D., Crawford-Redick, C. L., and Loomis, C. R. (1993) Biochim. Biophys. Acta 1165, 314-320[Medline] [Order article via Infotrieve]
  66. Liu, P., and Anderson, R. G. W. (1995) J. Biol. Chem. 270, 27179-27185[Abstract/Free Full Text]
  67. Jacobson, M. D., and Raff, M. C. (1995) Nature 374, 814-816[CrossRef][Medline] [Order article via Infotrieve]
  68. Muschel, R., Bernhard, E., Garza, L., McKenna, W., and Koch, C. (1995) Cancer Res. 55, 995-998[Abstract]


Copyright © 1998 by The American Society for Biochemistry and Molecular Biology, Inc.