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Originally published In Press as doi:10.1074/jbc.M400977200 on April 21, 2004

J. Biol. Chem., Vol. 279, Issue 27, 28614-28624, July 2, 2004
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Ceramide-induced Intracellular Oxidant Formation, Iron Signaling, and Apoptosis in Endothelial Cells

PROTECTIVE ROLE OF ENDOGENOUS NITRIC OXIDE*

Toshiyuki Matsunaga, Srigiridhar Kotamraju, Shasi V. Kalivendi, Anuradha Dhanasekaran, Joy Joseph, and B. Kalyanaraman{ddagger}

From the Department of Biophysics and Free Radical Research Center, Medical College of Wisconsin, Milwaukee, Wisconsin 53226

Received for publication, January 28, 2004 , and in revised form, March 31, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Sphingolipid ceramide (N-acetylsphingosine), a bioactive second messenger lipid, was shown to activate reactive oxygen species (ROS), mitochondrial oxidative damage, and apoptosis in neuronal and vascular cells. The proapoptotic effects of tumor necrosis factor-{alpha}, hypoxia, and chemotherapeutic drugs were attributed to increased ceramide formation. Here we investigated the protective role of nitric oxide (·NO) during hydrogen peroxide (H2O2)-mediated transferrin receptor (TfR)-dependent iron signaling and apoptosis in C2-ceramide (C2-cer)-treated bovine aortic endothelial cells (BAECs). Addition of C2-cer (5–20 µM) to BAECs enhanced ·NO generation. However, at higher concentrations of C2-cer (≥20 µM), ·NO generation did not increase proportionately. C2-cer (20–50 µM) also resulted in H2O2-mediated dichlorodihydrofluorescein oxidation, reduced glutathione depletion, aconitase inactivation, TfR overexpression, TfR-dependent uptake of 55Fe, release of cytochrome c from mitochondria into cytosol, caspase-3 activation, and DNA fragmentation. Nw-Nitro-L-arginine methyl ester (L-NAME), a nonspecific inhibitor of nitricoxide synthases, augmented these effects in BAECs at much lower (i.e. nonapoptotic) concentrations of C2-cer. The 26 S proteasomal activity in BAECs was slightly elevated at lower concentrations of C2-cer (≤10 µM) but was greatly suppressed at higher concentrations (>10 µM). Intracellular scavengers of H2O2, cell-permeable iron chelators, anti-TfR receptor antibody, or mitochondria-targeted antioxidant greatly abrogated C2-cer- and/or L-NAME-induced oxidative damage, iron signaling, and apoptosis. We conclude that C2-cer-induced H2O2 and TfR-dependent iron signaling are responsible for its prooxidant and proapoptotic effects and that ·NO exerts an antioxidative and cytoprotective role.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Ceramide belongs to a group of naturally occurring sphingolipid second messenger molecules that is formed by sphingomyelinase-catalyzed hydrolysis of sphingomyelin (1–3). There is growing interest on the potential role of ceramide-mediated proapoptotic cell signaling in response to treatment with reactive oxygen species (ROS)1 (e.g. superoxide and hydrogen peroxide) and other proapoptotic stress factors, including inflammatory cytokines such as tumor necrosis factor-{alpha} and lipopolysaccharide, hypoxia, and chemotherapeutic drugs (4–6). Exogenous treatment of endothelial cells and neuronal cells with ceramide also caused oxidative stress and activation of caspase-3 leading to apoptosis (7–12). Ceramide treatment has been shown to trigger both nitric oxide (·NO) and superoxide generation in endothelial cells (13–15). The relative ratio between superoxide and ·NO determine the ultimate cytotoxicity in ceramide-treated cells (14). Exposure of endothelial cells to lower concentrations of ceramide (~5 µM) causes an increase in ·NO formation due to Ca2+ activation and translocation of endothelial nitric-oxide synthase (eNOS) (14, 15). At higher concentrations (>20 µM) ceramide treatment induced ROS formation in cells (13, 14).

Previously, we have shown that treatment of endothelial cells with H2O2 induced intracellular oxidative stress, iron signaling, and apoptosis through stimulation of the transferrin receptor (TfR) (16, 17). More recently, we showed that ·NO mitigates peroxide-induced oxidative stress and apoptosis by inhibiting TfR-mediated iron signaling (18). Because ceramide induces both ·NO and ROS at different concentrations (14), we decided to explore in detail the effect of ceramide on oxidative cell signaling. In this study, we investigated the effects of C2-ceramide or N-acetylsphingosine, a cell-permeable ceramide analog, and C2-dihydroceramide, an inactive negative control for C2-ceramide (Fig. 1), on intracellular oxidant generation and iron signaling. The protective role of endogenously generated ·NO (19, 20) on ceramide-induced apoptosis was determined. Several fluorescence probes were used to monitor superoxide-, iron-, and peroxide-induced oxidant generation and ·NO/oxygen interaction. Results indicate that C2-ceramide induces ROS- and TfR-mediated iron signaling that are responsible for C2-cer-induced proapoptotic effects and that ·NO generation by ceramide affords cytoprotection against C2-ceramide-induced oxidative damage and apoptosis.



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  		FIG. 1.
Structures of C2-ceramide, naturally occurring ceramide, and the inactive C2-ceramide analog.

 

   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—C2-ceramide and C2-dihydroceramide were purchased from BIOMOL. Nw-nitro-L-arginine-methyl ester (L-NAME) was from Alexis. Glutathione monoethyl ester and desferal (or desferrioxamine) were obtained from Sigma. 2',7'-Dichlorodihydrofluorescein diacetate (DCFH-DA) and dihydroethidium (DHE) were from Molecular Probes. Fe(III)-tetrakis-(4-benzoic acid) porphyrin (FeTBAP) was synthesized according to a published method (21). Mito-Q was synthesized according to the published procedure (22), and monoclonal antibody 42/6, against human TfR (IgA class), was obtained from Dr. Ian Trowbridge (Salk Institute, San Diego, CA).

Endothelial Cell Culture—Bovine aortic endothelial cells (BAECs) were obtained from the Clonetics Corp. Cells were obtained at the third passage, transferred to 75-cm2 filter vent flasks (Costar, Cambridge, MA), and grown to confluence (5.2 x 106 cells/75 cm2) in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS), L-glutamine (4 mM), penicillin (100 units/ml), and streptomycin (100 µg/ml), incubated at 37 °C in a humidified atmosphere of 5% CO2 and 95% air. Cells were passaged as described by Balla et al. (23) and used between passages 4 and 12. On the day of the treatment, the medium was replaced with DMEM containing 2% FBS, which contains ~25–30 µg of transferrin/ml. The above experimental conditions were used in all the experiments performed in this study.

Measurement of Oxidative Stress—The level of intracellular oxidant production was estimated by oxidations of DHE and DCFH. Following treatment of BAECs with C2-ceramide, the medium was aspirated, and cells were washed with DPBS and incubated in 2 ml of fresh culture medium without FBS. DHE and DCFH-DA were added at a final concentration (10 µM) and incubated for 20 min, respectively. The cells were then washed twice with DPBS and maintained in 1 ml of culture medium. Fluorescence was monitored using a Nikon fluorescence microscope equipped with rhodamine and FITC filters. The intensity values were calculated using Metamorph software.

Measurement of 55Fe Uptake—55Fe uptake into the cells was measured as described previously (16, 24). Briefly, 0.2 µCi/ml 55Fe (ferric chloride) was added to the medium for 0–8 h, and its levels were measured as a function of time. Cells were washed twice with DPBS and lysed with PBS containing 0.1% Triton X-100, and the cell lysate was counted in a beta counter.

Western Blotting of TfR, PARP, Hsp-70, and Bcl-2—After the treatment with C2-ceramide, the cells were washed with ice-cold DPBS and resuspended in 150 µl of radioimmune precipitation assay buffer (20 mM Tris-HCl, pH 7.4, 2.5 mM EDTA, 1% Triton X-100, 1% sodium deoxycholate, 1% SDS, 100 mM NaCl, 100 mM sodium fluoride) containing 1 mM sodium vanadate, 10 µg/ml aprotinin, 10 µg/ml leupeptin, and 10 µg/ml pepstatin inhibitors. Cells were homogenized by passing the suspension through a 25-gauge needle (10 strokes). The lysate was centrifuged at 750 x g for 10 min at 4 °C to pellet out the nuclei. The remaining supernatant was centrifuged for 30 min at 12,000 x g. Protein was determined by Lowry method, and 20 µg of the lysate was used for the Western blot analysis. Proteins were resolved on SDS-polyacrylamide gels and blotted onto nitrocellulose membranes. Membranes were washed with TBS (140 mM NaCl, 50 mM Tris-HCl, pH 7.2) containing 0.1% Tween 20 (TBST) and 5% skim milk to block the nonspecific protein binding. Membranes were incubated with 1 µg/ml mouse anti-human transferrin receptor monoclonal antibody (Zymed Laboratories Inc., San Francisco, CA), mouse anti-bovine poly(ADP-ribose) polymerase (PARP) monoclonal antibody (Zymed Laboratories Inc.), mouse anti-human Hsp-70 antibody (Zymed Laboratories Inc.) or hamster anti-human Bcl-2 monoclonal antibody (BD Pharmingen) in TBST for 2 h at room temperature, washed 5 times, and then incubated with goat anti-mouse IgG-horseradish peroxidase-conjugated secondary antibody for TfR, PARP, Hsp-70, and mouse anti-hamster (1:5,000) for Bcl-2 for 1.5 h at room temperature. The band was detected using the ECL method (Amersham Biosciences).

Mitochondrial Cytochrome c Release—The release of mitochondrial cytochrome c into the cytosol in C2-ceramide-treated BAECs was measured according to the methods as described previously (17, 25). Briefly, BAECs were washed with DPBS and homogenized in PBS supplemented with 40 µg/ml saponin. Lysate was centrifuged at 750 x g for 10 min and followed by 12,000 x g for 20 min. The supernatant was used as the cytosolic fraction to measure the released cytochrome c into the cytosol by Western blot analysis using a mouse anti-cytochrome c antibody (BD Pharmingen). Detection was by horseradish peroxidase-conjugated goat anti-mouse antibody using the ECL method.

Measurement of Caspase Activities—Cytosolic enzymatic activities of caspase-3, caspase-6, caspase-8, and caspase-9 were measured as described previously (26). Briefly, cells were washed twice with DPBS following treatment with C2-ceramide and then lysed with 50 mM HEPES buffer (pH 7.4) containing 5 mM CHAPS and 5 mM dithiothreitol. After cytosolic fraction was taken by centrifugation at 12,000 x g for 30 min, the activities of caspase-3, caspase-6, caspase-8, and caspase-9 were measured using the substrates of ac-DEVD-pNA (acetyl-Asp-Glu-Val-Asp-p-nitroanilide), ac-VEID-pNA (acetyl-Val-Glu-Ile-Asp-pNA), ac-IETD-pNA (acetyl-Ile-Glu-Thr-Asp-pNA), and ac-LEHD-pNA (acetyl-Leu-Glu-His-Asp-pNA) respectively. The absorbance at 405 nm of the released pNA was monitored in a spectrophotometer and quantitated using a pNA standard.

Measurement of Apoptosis by TUNEL Assay—The terminal deoxynucleotidyl transferase-mediated nick-end labeling (TUNEL) assay was used for microscopic detection of apoptosis (26). This assay is based on labeling of 3' free hydroxyl ends of the fragmented DNA with fluorescein-dUTP catalyzed by terminal deoxynucleotidyl transferase. Procedures were followed according to a commercially available kit (ApoAlert) from Clontech. Apoptotic cells exhibit a strong nuclear green fluorescence that can be detected using a standard fluorescein filter (520 nm). All cells stained with propidium iodide exhibit a strong red cytoplasmic fluorescence at 620 nm. The areas of apoptotic cells were detected by fluorescence microscopy equipped with rhodamine and FITC filters. The quantification of apoptosis was performed using the Metamorph image analysis package.

Measurement of Intracellular ·NO—Intracellular ·NO levels were monitored using a DAF-2 fluorescence probe (27, 28). The treated cells were washed with DPBS and incubated in 2 ml of fresh culture medium without FBS. DAF-2 was added at a final concentration of 10 µM, and the cells were incubated for 20 min. The cells were washed twice with DPBS and maintained in 1 ml of culture medium for monitoring the fluorescence using a Nikon fluorescence microscope (excitation, 488 nm; emission, 610 nm) equipped with an FITC filter. The values of fluorescent intensity were calculated using the Metamorph software.

Measurement of Aconitase Activity—BAECs were washed twice with cold DPBS and lysed with buffer containing 0.2% Triton X-100, 100 µM diethylenetriaminepentaacetic acid, and 5 mM citrate in PBS. The activity of aconitase was measured in 100 mM Tris-HCl (pH 8.0) containing 20 mM DL-trisodium isocitrate. An extinction coefficient for cis-aconitate of 3.6 mM-1 at 240 nm was used (29).

Measurement of Glutathione—The level of glutathione (GSH) was measured by HPLC as the o-phthalaldehyde (OPA) adduct at pH 8.0 (30). BAECs were washed twice with DPBS, suspended in 250 µl of PBS, and lysed by sonication. After centrifugation at 10,000 x g for 2 min, 200 µl of the clear supernatant was derivatized by incubation for 30 min at room temperature with OPA. An aliquot of sample was injected onto a Kromasil C-18 column and eluted isocratically with a mobile phase consisting of 150 mM sodium acetate:methanol (91.5:8.5, v/v). The OPA-GSH adduct was monitored using a fluorescence detector operating at excitation and emission wavelengths at 250 and 410 nm, respectively. The levels of intracellular GSH were quantified using a GSH solution as a standard.

Proteasome Function Assays—For 26 S proteasome, proteasomal activity was measured as previously reported (31, 32). Briefly, cells were washed with buffer I (50 mM Tris, pH 7.4/2 mM dithiothreitol/5 mM MgCl2/2 mM ATP) and homogenized with buffer I containing 250 mM sucrose. Twenty micrograms of 10,000 x g supernatant was diluted with buffer I to a final volume of 1 ml. The fluorogenic proteasome substrates SucLLVY-AMC (chymotrypsin-like) and Z-Leu-Leu-Lys-7-amido-4-methylcoumarin (trypsin-like) were added in a final concentration of 100 and 80 µM, respectively. Proteolytic activity was measured by monitoring the release of the fluorescent group 7-amido-4-methylcoumarin (excitation, 380 nm; emission, 460 nm).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Ceramide Induces Intracellular Superoxide and Hydrogen Peroxide—Intracellular ROS levels were measured in BAECs treated with different concentrations of C2-cer for different time periods. The oxidation of DCFH, a nonfluorescent probe, to a fluorescent dichlorofluorescein (DCF) was used to measure intracellular H2O2-derived oxidants. Although H2O2 itself does not react with DCFH to form DCF, it was proposed that intracellular peroxidases or redox-active metal ions could catalyze the oxidation of DCFH to DCF in the presence of H2O2 (17, 33). Results show that C2-ceramide induced a dose- and time-dependent increase in DCF staining (Fig. 2, A–C). DCF fluorescence was noticeable in cells treated with 20 µM ceramide and reached a maximum in cells exposed to 50 µM C2-ceramide for 8 h (Fig. 2, B and C). The oxidation of the probe gradually increased over a 2- to 8-h time period in cells treated with 50 µM C2-ceramide. Incubation with an inactive form of ceramide, C2-dihydroceramide, lacking a double bond did not induce any appreciable DCF green fluorescence (Fig. 2, A and B). These results suggest that the active C2-ceramide induces intracellular oxidant generation as detected by DCF fluorescence.



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  		FIG. 2.
Ceramide-induced superoxide and H2O2 generation in endothelial cells using hydroethidine and dichlorodihydrofluorescein probes. A, BAECs were treated with various concentrations of C2-ceramide (a–d) or 50 µM of C2-dihydroceramide (e) for 8 h. Also cells were treated with 50 µM C2-ceramide for different time periods (f–j). After the treatments the medium was aspirated, and the cells were washed twice with DPBS and subsequently incubated with 10 µM DCFH-DA (A) or 10 µM dihydroethidium (D) for 20 min. The cells were then washed with DPBS and maintained in 1 ml of DMEM. The green fluorescence characteristic of DCF (A–C) and red fluorescence caused by ethidium (D–F) were measured using fluorescein isothiocyanate and rhodamine filters, respectively, in a Nikon fluorescence microscope. The data shown are representative of three independent experiments.

 
Next, we determined the effect of C2-cer in cells treated with dihydroethidium (DHE), a fluorescent probe that reacts with superoxide to form a characteristic red fluorescence. Recent reports indicate that superoxide anion reacts with dihydroethidium to form a product that is distinctly different from ethidium (34). As shown in Fig. 2 (D–F), there was a dose- and time-dependent increase in the intensity of red fluorescence with ceramide treatment, indicative of enhanced superoxide generation. Again the inactive form of C2-cer, C2-dihydroceramide, did not induce red fluorescence in cells treated with HE (Fig. 2, D and E).

Ceramide-induced 55Fe Uptake and TfR Expression—We determined the role of transferrin iron in ceramide-induced apoptosis and oxidant formation in endothelial cells by measuring TfR levels and 55Fe uptake. BAECs were treated with different concentrations of C2-cer for various time periods as described in Fig. 3C, and TfR levels were measured under these conditions. Results show that ceramide treatment increased TfR levels in a dose- and time-dependent manner (Fig. 3, A and B). TfR levels were found to be significantly increased by 2 h with 50 µM C2-ceramide (Fig. 3B). The inactive analog of C2-ceramide did not appreciably affect TfR levels in BAECs. Consistent with the increase in TfR levels, the uptake of 55Fe was considerably increased in C2-ceramide-treated cells (Fig. 3C). Pretreatment with anti-TfR antibody inhibited ceramide-induced 55Fe uptake (not shown). These results clearly indicate a definite role for Tf-iron in ceramide-induced oxidative stress in endothelial cells.



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  		FIG. 3.
The effect of C2-ceramide on transferrin receptor levels and iron uptake in endothelial cells. BAECs were treated with different concentrations of C2-ceramide (A) and for different time periods either with 50 µM C2-ceramide or 50 µM C2-dihydroceramide (B) and transferrin receptor levels were measured by Western analysis using a monoclonal anti-TfR antibody. C, BAECs were treated with 50 µM C2-ceramide for various time periods along with 0.2 µCi/ml 55Fe, and the uptake of the labeled iron was measured as described under "Experimental Procedures." Data shown are representative of three separate experiments. *, significantly different (p < 0.05) compared with untreated conditions.

 
Ceramide-induced Apoptosis in Endothelial Cells—To investigate the mitochondrial involvement in ceramide-induced apoptosis, we measured the release of cytochrome c from the mitochondria into the cytosol. Incubation of BAECs with C2-ceramide for 8 h caused a dose-dependent increase in the release of cytochrome c into the cytosol (Fig. 4A). BAECs treated with 50 µM of C2-ceramide induced a decrease in the antiapoptotic protein Bcl-2 located on the outer membrane of the mitochondria (Fig. 4B) as well as in the Hsp-70 protein levels (Fig. 4B). We then identified the actual caspase family (caspases 3, 6, 8, and 9) that was activated during ceramide-induced apoptosis. As shown in Fig. 4, only the activity of the effector caspase-3 began to increase as early as 4 h in ceramide-treated cells, reaching a peak value by 8 h. In addition, the increase in caspase-3 activity was detected in cells only when the concentration of C2-ceramide exceeded 20 µM (Fig. 4C). In contrast, the caspase-3 activation was not evident in BAECs treated with an inactive analog of C2-ceramide, namely C2-dihydroceramide (Fig. 1). Although there was a marginal increase in the activation of caspase-6 and caspase-8 (Fig. 4, E and F), their significance in C2-ceramide-induced apoptosis was not apparent, because their activities became noticeable only at 16 h; at this time point, however, the execution of DNA fragmentation had already occurred (Fig. 4H). The marginal increase observed in the caspase-9 activity (Fig. 4G) by 4-h treatment with C2-ceramide would likely activate the caspase-3 activity, initiating a possible feedback loop that further increased their individual activities.



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  		FIG. 4.
Ceramide-induced mitochondrial release of cytochrome c and alterations in antiapoptotic protein levels and caspase activation in endothelial cells. A, BAECs were treated with different concentrations of C2-ceramide for 8 h and release of cytochrome c from mitochondria was measured by Western analysis using an anti-cytochrome c antibody. B, the alterations in the levels of Hsp-70 and Bcl-2 (antiapoptotic proteins) in response to 50 µM C2-ceramide, treated for various time periods. C, dose response of C2-ceramide in the activation of caspase-3 enzyme, and the data shows that caspase-3 activation occurs at 50 µM or higher concentrations of C2-ceramide. D, the activation of caspase-3 with 50 µM C2 ceramide from 8 h of incubation. E–G, effect of C2-ceramide in the activation of other caspases like 6, 8, and 9, respectively. H, effect of 50 µM C2-ceramide on PARP cleavage as a function of time, and, as shown above, the cleavage of inactive PARP (~116 kDa) to its active fragment (~85 kDa) starts at 4 h of incubation with ceramide. I, cells were treated with different concentrations of C2-ceramide for 16 h and stained for TUNEL-positive cells as an index extent of DNA fragmentation examined by fluorescence microscopy (original magnification, x100). Photographs are overlaid images of propidium iodide- and FITC-stained cells (TUNEL-positive cells as indicated by arrows). Yellow and red denote apoptotic and nonapoptotic cells, respectively.

 
The time-dependent increase in caspase-3 activity was further correlated with its downstream target, namely, poly(ADP-ribose) polymerase (PARP) cleavage (Fig. 4H), which is responsible for DNA fragmentation. The PARP cleavage started to appear at 4 h of treatment with C2-ceramide and significantly increased at 8 and 16 h of incubation with C2-ceramide. The inactive PARP precursor protein (~116 kDa) was cleaved to form a ~85-kDa active fragment (Fig. 4H). This active fragment of PARP translocates into the nucleus and cleaves the DNA. Finally, a dose-dependent increase in DNA fragmentation was shown by TUNEL staining (Fig. 4I) wherein cells were treated with different concentrations of C2-ceramide (0–50 µM) for a period of 8 h. As shown, C2-ceramide treatment increased the TUNEL-positive staining in cells, from 1.5% (0 µM) to 48.4% (50 µM). Treatment with C2-dihydroceramide, the inactive ceramide, did not yield DNA fragmentation as detected by TUNEL staining (Fig. 4C).

Antioxidants and Iron Chelators Mitigate Ceramide-induced Oxidative Stress and Apoptosis—The effects of different antioxidants and iron chelators in C2-ceramide-treated cells were investigated. BAECs were pretreated for 2 h with GSH ester (5 mM), FeTBAP (25 µM), desferal (20 µM), or anti-TfR antibody (12 µg/ml, 42/6, IgA class, which specifically binds to the extracellular domain of the transferrin receptor and inhibits receptor endocytosis) prior to treating cells with 50 µM C2-ceramide for 8 h. These agents inhibited ceramide-induced DCFH oxidation (Fig. 5, A and B) and caspase-3 activation (Fig. 5D). Pretreatment with a mitochondria-targeted antioxidant (e.g. Mito-Q) showed similar inhibitory effects, suggesting that mitochondrial generation of ROS is responsible for ceramide-mediated oxidative stress and apoptosis (Fig. 5C). Under these conditions, these antioxidants and iron chelators also inhibited ceramide-induced DNA fragmentation as shown by TUNEL staining (Fig. 5D).



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  		FIG. 5.
Effect of antioxidants and iron chelators on ceramide-induced H2O2 generation and DNA fragmentation. A, BAECs were pre-treated with either GSH ester (5 mM), FeTBAP (25 µM), desferal (20 µM), or anti-TfR antibody (IgA class, 12 µg/ml) for 2 h before the addition of 50 µM C2-ceramide for 8 h, and DCF staining was carried out as described in Fig. 1 and under "Experimental Procedures." B, fluorescence intensity of data shown in A and C, same as A, except that after the treatments caspase-3 activity was measured spectrophotometrically at 405 nm by following the release p-nitroanilide. D, same as A and C except that cells were treated for 16 h with C2-ceramide and other antioxidants as indicated and DNA fragmentation was measured by TUNEL staining. The data represent mean ± S.D. of three independent experiments.

 
Effect of C2-ceramide on Intracellular ·NO Generation in Endothelial Cells—At low concentrations (5–20 µM), ceramide enhanced ·NO release in BAECs (15). Thus, intracellular ·NO levels were monitored using DAF-2 fluorescence. It has been shown that DAF-2 forms a fluorescent triazolo-type product in the presence of an oxidant derived from ·NO and molecular oxygen (28). We observed a dose- and time-dependent increase in DAF-2 fluorescence in endothelial cells treated with C2-ceramide, but during longer exposure time there was a disproportionate increase in DAF-2 fluorescence that started to plateau in cells treated with 50 µM C2-cer (Fig. 6, A and B). Ceramide-induced intracellular generation of ·NO was further confirmed using a nitric-oxide synthase inhibitor, L-NAME. Pretreatment of L-NAME for 2 h before the addition of 50 µM C2-ceramide for 8 h inhibited ceramide-induced DAF-2 fluorescence in a dose- and time-dependent manner (Fig. 6, C–F).



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  		FIG. 6.
Effect of ceramide and NOS inhibitor on nitric oxide generation in endothelial cells. A, BAECs were treated with different concentrations of C2-ceramide for either 1 or 8 h and at the end of the experiment, cells were washed twice with DPBS and replaced with 2 ml of culture medium, and 5 µM DAF-2-DA was added and further incubated for 15 min. Cells were washed free of extracellular DAF-2-DA and immediately viewed under the fluorescence microscope equipped with an FITC filter to measure the green fluorescence as an index of nitric oxide levels. B, fluorescence intensity of data shown in A, and C, same as A except that cells were pretreated with different concentrations of L-NAME for 2 h before the addition of 50 µM C2-ceramide for 8 h. D, fluorescence intensity of data shown in C; E, same as A except that cells were treated with 50 µM C2-ceramide for different time points in the absence or presence of 2 mM L-NAME for 8 h. F, fluorescence intensity of data shown in E. Data represent the mean ± S.D. of three separate experiments.

 
NOS Inhibitor Exacerbates Ceramide-induced Intracellular ROS—Ceramide, at lower concentrations (5–20 µM), induced little or no ROS (Fig. 6A) (15). However, in the presence of L-NAME, ROS generation increased in cells treated with lower concentrations of ceramide, as measured by DCF and hydroethidium fluorescence (Fig. 7, A and B). This suggests that ceramide-induced ·NO counteracts the effects of ROS or ROS generation induced by ceramide; at higher concentrations of C2-ceramide, ROS generation overwhelms the protective effects of ·NO. NOS inhibition sensitized these cells to ceramide-induced oxidative stress. This is in agreement with the data showing a disproportionate increase in ·NO in cells treated with C2-ceramide during prolonged incubation (Fig. 6, A and B), as compared with shorter period of incubation. These results show that L-NAME exacerbates C2-ceramide-induced ROS.



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  		FIG. 7.
Effect of NOS inhibitor on ceramide-induced H2O2 and superoxide generation in endothelial cells. BAECs were treated with low concentrations of C2-ceramide (5 and 20 µM) for 8 h either in the presence or absence of 2 mM L-NAME. After the treatments H2O2 (A) and superoxide (B) staining were carried out by DCF and ethidium staining, respectively, as described under "Experimental Procedures." Data shown represent mean ± S.D. of three independent experiments.

 
The next step was to investigate whether Tf-iron could play a potential role in mediating ceramide toxicity. To this end, GSH levels were measured in the presence or absence of L-NAME in C2-ceramide treated cells. Results show that L-NAME significantly enhanced the depletion of GSH levels caused by C2-ceramide (≤20 µM) (Fig. 8A). Under these conditions, the total aconitase activity was significantly lower in cells treated with both L-NAME and ceramide (Fig. 8B). The TfR levels were increased at much lower levels of ceramide in cells pretreated with L-NAME (Fig. 8C). This result prompted us to measure the iron uptake under similar conditions. We found that L-NAME treatment synergistically enhanced ceramide-mediated uptake of radiolabeled iron (Fig. 8D). At lower concentrations of C2-ceramide (5 µM), L-NAME treatment increased iron uptake compared with C2-ceramide alone (Fig. 8D). Thus, intracellular depletion of ·NO augments ceramide-induced iron signaling.



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  		FIG. 8.
Effect of NOS inhibitor on ceramide-induced GSH depletion, aconitase activity, TfR expression, and iron uptake in endothelial cells. A, BAECs were treated with different concentrations of C2-ceramide in the presence or absence of 2 mM L-NAME (pretreatment for 2 h), and the GSH levels were determined by measuring the GSH-o-phthalaldehyde adduct using HPLC. B, same as A except that the total aconitase activity was measured spectrophotometrically at 240 nm. C and D, same as A except that the transferrin receptor levels and 55Fe uptake were measured with and without L-NAME. Cells were treated with different concentrations of C2-ceramide in the presence or absence of 2 mM L-NAME for 8 h, and the uptake of labeled iron was measured as described under "Experimental Procedures." Data represent mean ± S.D. of at least three separate experiments. *, significantly different (p < 0.05) compared with untreated conditions.

 
In addition to enhancing oxidant generation, L-NAME treatment exacerbated C2-ceramide-mediated apoptotic effects (Fig. 9A). The release of cytochrome c from the mitochondria was increased in the presence of L-NAME and C2-ceramide, as compared with C2-ceramide or L-NAME alone. Similar results were observed with caspase-3 activity, i.e. lower concentrations of C2-ceramide (10–20 µM) in the presence of L-NAME increased the caspase-3 activity (Fig. 9B). The DNA fragmentation (Fig. 9C)as measured by the TUNEL staining of cells increased in cells treated with 5 µM and 20 µM C2-ceramide for 16 h in the presence of L-NAME as compared with C2-ceramide alone.



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  		FIG. 9.
NOS inhibitor exacerbates ceramide-induced apoptosis in endothelial cells. A, BAECs were pretreated with 2 mM L-NAME for 2 h and subsequently treated with 20 µM C2-ceramide for 8 h, and the release of cytochrome c from the mitochondria was measured by Western analysis using an anti-cytochrome c antibody. B, cells were treated with low concentrations of C2-ceramide (5–20 µM) for 8 h in the presence or absence of 2 mM L-NAME (2-h pretreatment), and caspase-3 activity was measured spectrophotometrically. C, same as in B except that cells were treated for 16 h with 5 and 20 µM of C2-ceramide either in the presence or absence of L-NAME, and DNA fragmentation was measured by TUNEL-positive staining as described under "Experimental Procedures."

 
In the presence of antioxidants and iron chelator, C2-ceramide/L-NAME-induced ROS generation was inhibited. As shown in Fig. 10, C2-ceramide/L-NAME-mediated DCF fluorescence (Fig. 10, A and B) and caspase-3 activity (Fig. 10C) in cells treated with either GSH ester (5 mM), FeTBAP (25 µM), Mito-Q (1 µM), desferal (20 µM), or anti-TfR antibody (12 µg/ml, IgA class) was considerably decreased. These results indicate that C2-ceramide induces the formation of ROS to a greater extent in L-NAME-treated cells and that Tf-iron plays a predominant role in ceramide-mediated toxicity.



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  		FIG. 10.
Effect of co-incubation of NOS inhibitor and antioxidants and iron chelators on ceramide-induced H2O2 generation and caspase-3 activation in endothelial cells. A, BAECs were treated with 20 µM C2-ceramide alone or ceramide plus L-NAME or ceramide plus L-NAME plus antioxidants or iron chelators for 8 h, and H2O2 generation was measured as an index of DCF fluorescence. Note that L-NAME or other compounds were pretreated for 2 h before the addition of C2-ceramide. In the case of antioxidant or iron chelator plus L-NAME plus ceramide groups, antioxidant or iron chelator was added 2 h prior to the addition of L-NAME, which was added 2 h prior to the addition of ceramide. B, fluorescence intensity of data shown in A; C, same as A except that caspase-3 activity was measured spectrophotometrically at 405 nm by following the release of p-nitroanilide. ++, cells were pretreated with both L-NAME (2 mM) and different antioxidants before they were treated with C2-ceramide (20 µM). Data represent the mean ± S.D. of three separate experiments.

 
The Biphasic Effect of Ceramide on Proteasomal Activities—Next, we investigated whether C2-cer treatment modulates the proteasomal activity in BAECs. Fig. 11 shows the trypsin-like and chymotrypsin-like activities of the 26 S proteasome in BAECs treated with different concentrations of C2-cer. In the presence of C2-cer (10 µM), there was initially a slight increase in the proteasomal activity that drastically decreased at higher C2-cer concentrations (Fig. 11, A and B). This biphasic effect is attributed to the fact that lower ceramide concentration stimulates ·NO generation, whereas ROS formation is favored at higher ceramide concentrations (14, 15). This is in agreement with the previous report indicating that ·NO, both exogenous and endogenous, enhanced the proteasomal activity, whereas ROS suppressed the proteasomal function in BAECs (18). Pretreatment of cells with antioxidants (Mito-Q and desferal) considerably increase the proteasomal activity in ceramide-treated cells (Fig. 11, C and D). This is attributed to a decrease in ROS and increase in ·NO in Mito-Q- and desferal-treated cells (data not shown).



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  		FIG. 11.
Effect of ceramide on the proteasomal activity in BAECs. BAECs were treated with different concentrations of C2-ceramide in the presence and absence of L-NAME (2 mM), Mito-Q (1 µM), or desferal (20 µM) for 6 h. The chymotrypsin-like (A) and trypsin-like (B) activities of 26 S proteasome were measured in cell lysates as described under "Experimental Procedures." Data represent the mean ± S.D. of three separate experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we demonstrated that a cell-permeable bioactive ceramide analog induces ·NO at lower concentrations (≤20 µM) and ROS at higher concentrations (>20 µM) in bovine aortic endothelial cells. Results showed that ceramide-induced TfR-dependent iron uptake was responsible for its prooxidant and proapoptotic effects, because pretreatment with TfR antibody or cell-permeable iron chelators greatly mitigated these effects. Depletion of intracellular ·NO augmented ceramide-induced iron signaling, oxidative stress, and apoptosis. ·NO suppressed the prooxidant and proapoptotic effects of ceramide by maintaining intracellular iron homeostasis.

DCF Fluorescence as an Indicator of Oxidant-induced Transferrin Iron Signaling—Ceramide-induced oxidative stress was assessed by monitoring the increase in DCF fluorescence (Fig. 2A). Because ceramide-induced DCF fluorescence was inhibited by anti-TfR antibody, we concluded that TfR-transported Tf-iron was responsible for catalyzing intracellular oxidation of DCFH to DCF. We have recently shown that oxidative inactivation of iron-sensing iron-sulfur proteins (e.g. aconitase) was responsible for cellular iron uptake through increased iron-regulatory protein (IRP) activity (16, 17). Inactivation of aconitase by oxidative disassembly of the [4Fe-4S] cluster is accompanied by enhanced IRP activation through increased mRNA-binding activity associated with the iron-responsive element (35, 36). Increased IRP1/iron-responsive element binding stabilizes the TfR mRNA, leading to enhanced mRNA translation, TfR synthesis, and Tf-iron uptake (37). The exogenous addition of bolus or continuously generated H2O2 to endothelial cells caused enhanced oxidation of DCFH to DCF that was regulated by TfR-mediated uptake of Tf iron (17). Pretreatment of cells with anti-TfR antibody that specifically binds to the extracellular domain of TfR inhibited H2O2-induced iron signaling and ROS-mediated DCF fluorescence. The present data indicate that intracellular oxidation of DCFH is catalyzed by ceramide-induced H2O2 and Tf-iron transported through TfR.

Superoxide/Hydroethidine-induced Intracellular Red Fluorescence—Hydroethidine or dihydroethidium has been widely used to detect intracellular superoxide (16, 34). This assay is based on the fact that the product formed from the reaction between superoxide and dihydroethidium exhibits a red fluorescence. This product has long been thought to be ethidium (38). We recently reported that the fluorescence characteristics of the superoxide/hydroethidine product are distinctly different from those of ethidium (34). In addition, the HPLC retention time of ethidium is different from that of the product formed from the reaction between superoxide and hydroethidine (34). Although the exact structure of this product is not known, HE is still a viable fluorescent probe for detecting intracellular superoxide by monitoring the "red fluorescence" formed from HE. Preliminary experiments show that cells treated with hydroethidine and C2-cer exhibit a product whose fluorescence yield was enhanced in the presence of DNA and whose HPLC profile was distinctly different from that of ethidium (not shown). Clearly, elucidating the reaction mechanism between hydroethidine and superoxide is pivotal to our understanding of the oxidative reactions induced by ceramides and other bioactive lipid mediators.

Mitochondria as a Source of Superoxide Generation in Ceramide-treated Cells—Based on the published data (4, 39–41), we propose that ceramide-induced ROS generation inhibits mitochondrial enzymes (aconitase and complex activity) associated with the electron transport chain, which in turn leads to more superoxide and H2O2. Inhibition of complex-1 activity stimulated superoxide formation through increased auto-oxidation of ubisemiquinone (43, 44). Ceramide-induced oxidative stress was thought to result from inhibition of mitochondrial complex III activity (45). The molecular signaling events induced by the lipid second messenger, C2-ceramide, and their role in inhibiting the respiratory enzymes are not fully known. The activation of rac 1, the protooncogene family member and a regulatory component of NADPH oxidase, in ceramide-treated endothelial cells was reported to induce mitochondrial oxidative stress (39). The role of mitochondria in ceramide-induced ROS generation is implicated, because pretreatment with mitochondria-targeted antioxidants (Mito-quinone or Mito-vitamin E) (22) greatly inhibited C2-cer-induced DCF and HE fluorescence (data not shown).

Nitric Oxide, Iron Homeostasis, and Proteasomal Activation—It has been previously shown that addition of low concentrations of C2-ceramide (5 µM) to BAECs significantly increases the eNOS activity due to translocation of eNOS from the endothelial cell membrane to intracellular sites (15). However, at higher concentrations of ceramide, even though there was an increased eNOS mRNA and protein expression, there was a decrease in the levels of bioactive ·NO (13). These findings are in accordance with the present data in that at low concentrations of C2-ceramide (< 20 µM), there was an increase in the generation of ·NO and a decrease in ROS generation, whereas, at higher concentrations of ceramide (> 20 µM), ROS generation becomes dominant. Although the reasons for the shift in mechanism are not completely understood, it is likely that ROS can rapidly react with ·NO, thereby reducing its bioactivity. The antioxidative and cytoprotective effects of ·NO were recently attributed to ·NO-induced proteasomal activation (18). Proteasomal inhibitors abrogate ·NO-mediated cytoprotection and antioxidative effects (18). ·NO as a stimulator of proteasomal function is still a nascent concept (18). It was suggested that ·NO mitigated peroxide-induced transferrin iron uptake, DCFH oxidation, and apoptosis by enhancing the proteolytic activity in endothelial cells (17, 18). Depletion of endogenous ·NO with L-NAME decreased the trypsin-like activity of 26 S proteasome in endothelial cells (18). Previous results also indicated that ·NO controls peroxide-induced iron signaling, intracellular iron homeostasis, and oxidative stress through increased proteasomal activation (18). In the present study, C2-cer-induced Tf-iron uptake, DCFH oxidation, and caspase-3 activation were greatly enhanced in the presence of L-NAME. Treatment with C2-cer activated the proteasomal function at lower concentrations (which induced ·NO), and higher concentrations of C2-cer (which induced more ROS and less ·NO) inhibited the intracellular proteasomal function (Fig. 11). C2-ceramide-induced proteolytic activation was inhibited by L-NAME, suggesting a role for ·NO. In contrast, L-NAME did not inhibit the proteasomal inactivation observed at higher concentrations of C2-cer (Fig. 11). If any, L-NAME enhanced the proteasomal inactivation induced by C2-cer at higher concentrations (Fig. 11). This is consistent with the enhanced iron uptake and oxidant generation observed in BAECs treated with C2-cer (5–20 µM) and L-NAME (Fig. 3C).

Ceramides: Prooxidant Lipid Mediators in Cardiovascular and Neurodegenerative Diseases?—Ceramide-induced oxidative stress was proposed to play a key role in the pathogenesis of age-related diseases with lipid abnormalities, leading to the accumulation of sphingomyelin, ceramide, and cholesterol esters (46–48). Altered lipid metabolism resulting in elevated ceramide levels and increased oxidative stress had been implicated in atherosclerosis, amyotrophic lateral sclerosis, and Alzheimer's disease (46, 49, 50). The ceramide-induced ROS-mediated apoptotic signaling pathway has been suggested to play a key role in the degeneration of dopaminergic neurons in patients with Parkinson's disease (7, 12). In these pathologies, perturbed iron metabolism or elevated iron levels were shown to be prominent. Thus, ceramide-induced oxidative stress and iron signaling reported in the present work explain in part the mechanism of initiation and propagation of lipid peroxidative processes in the pathogeneses of these diseases. ·NO levels are impaired in age-related cardiovascular and neurodegenerative diseases (42, 51). The present data suggest that there exists an inverse relationship between endogenous ·NO and cellular iron levels. Clearly, future studies should be directed at investigating more thoroughly the intriguing connection between ceramide, ·NO, and iron in neurovascular and cardiovascular pathologies.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants HL073056-01 and 1PO1HL68769-01. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed: Dept. of Biophysics, Medical College of Wisconsin, 8701 Watertown Plank Rd., Milwaukee, WI 53226. Tel.: 414-456-4035; Fax: 414-456-6512; E-mail: balarama{at}mcw.edu.

1 The abbreviations used are: ROS, reactive oxygen species; BAEC, bovine aortic endothelial cell; C2-cer, C2-ceramide; DCF, 2',7'-dichlorofluorescein; DCFH-DA, 2',7'-dichlorodihydrofluorescein diacetate; DAF-2-DA, diaminofluorescein diacetate; FeTBAP, Fe(III)-tetrakis-(4-benzoic acid) porphyrin; GSH, reduced glutathione; Tf, transferrin; TfR, transferrin receptor; DHE, dihydroethidium; Mito-Q, mixture of mitoquinone and mitoquinol conjugated to triphenyl phosphonium ion; L-NAME, Nw-nitro-L-arginine methyl ester; TUNEL, terminal deoxynucleotidyltransferase-mediated nick-end labeling; TBS, Tris-buffered saline; PBS, phosphate-buffered saline; DPBS, Dulbecco's PBS; eNOS, endothelial nitric-oxide synthase; FITC, fluorescein isothiocyanate; HE, hydroethidium; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; PARP, poly(ADP-ribose) polymerase; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; ac-, acetyl-; pNA, p-nitroanilide; HPLC, high-performance liquid chromatography; OPA, o-phthalaldehyde. Back



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Chatterjee, S. (1998) Arterioscler. Thromb. Vasc. Biol. 18, 1523-1533[Abstract/Free Full Text]
  2. Levade, T., Auge, N., Veldman, R. J., Cuvillier, O., Negre-Salvayre, A., and Salvayre, R. (2001) Circ. Res. 89, 957-968[Abstract/Free Full Text]
  3. Hannun, Y. A. (1996) Science 274, 1855-1859[Abstract/Free Full Text]
  4. Corda, S., Laplace, C., Vicaut, E., and Duranteau, J. (2001) Am. J. Respir. 24, 762-768
  5. Xu, J., Yeh, C.-H., Chen, S., He, L., Sensi, S. L., Canzoniero, L. M. T., Choi, D. W., and Hsu, C. Y. (1998) J. Biol. Chem. 273, 16521-16526[Abstract/Free Full Text]
  6. Barsacchi, R., Perrotta, C., Sestili, P., Cantoni, O., Moncada, S., and Clementi, E. (2002) Cell Death Differ. 9, 1248-1255[CrossRef][Medline] [Order article via Infotrieve]
  7. France-Lanord, V., Brugg, B., Michel, P. P., Agid, Y., and Ruberg, M. (1997) J. Neurochem. 69, 1612-1621[Medline] [Order article via Infotrieve]
  8. Escargueil-Blanc, I., Andrieu-Abadie, N., Caspar-Bauguil, S., Brossmer, R., Levade, T., Negre-Salvayre, A., and Salvayre, R. (1998) J. Biol. Chem. 273, 27389-27395[Abstract/Free Full Text]
  9. Erdreich-Epstein, A., Tran, L. B., Bowman, N. N., Wang, H., Cabot, M. C., Durden, D. L., Vlckova, J., Reynolds, C. P., Stins, M. F., Groshen, S., and Millard, M. (2002) J. Biol. Chem. 277, 49531-49537[Abstract/Free Full Text]
  10. Pettus, B. J., Chalfant, C. E., and Hannun, Y. A. (2002) Biochim. Biophys. Acta 1585, 114-125[Medline] [Order article via Infotrieve]
  11. Ito, A., Uehara, T., Tokumitsu, A., Okuma, Y., and Nomura, Y. (1999) Biochim. Biophys. Acta 1452, 263-274[Medline] [Order article via Infotrieve]
  12. Hunot, S., Brugg, B., Ricard, D., Michel, P. P., Muriel, M.-P., Ruberg, M., Faucheux, B. A., Agid, Y., and Hirsch, E. C. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 7531-7536[Abstract/Free Full Text]
  13. Li, H., Junk, P., Huwiler, A., Burkhardt, C., Wallerath, T., Pfeilschifter, J., and Forstermann, U. (2002) Circulation 106, 2250-2256[Abstract/Free Full Text]
  14. Sawai, H., Okazaki, T., Takeda, Y., Tashima, M., Sawada, H., Okuma, M., Kishi, S., Umehara, H., and Domae, N. (1997) J. Biol. Chem. 272, 2452-2458[Abstract/Free Full Text]
  15. Igarashi, J., Thatte, H. S., Prabhakar, P., Golan, D. E., and Michel, T. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 12583-12588[Abstract/Free Full Text]
  16. Kotamraju, S., Chitambar, C. R., Kalivendi, S. V., Joseph, J., and Kalyanaraman, B. (2002) J. Biol. Chem. 277, 17179-17187[Abstract/Free Full Text]
  17. Tampo, Y., Kotamraju, S., Chitambar, C. R., Kalivendi, S. V., Keszler, A., Joseph, J., and Kalyanaraman, B. (2003) Circ. Res. 92, 56-63[Abstract/Free Full Text]
  18. Kotamraju, S., Tampo, Y., Keszler, A., Chitambar, C. R., Joseph, J., Haas, A. L., and Kalyanaraman, B. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 10653-10658[Abstract/Free Full Text]
  19. Kotamraju, S., Tampo, Y., Kalivendi, S. V., Joseph, J., Chitambar, C. R., and Kalyanaraman, B. (2004) Arch. Biochem. Biophys. 423, 74-80[CrossRef][Medline] [Order article via Infotrieve]
  20. Tang, C. H., and Grimm, E. A. (2004) J. Biol. Chem. 279, 288-298[Abstract/Free Full Text]
  21. Day, B. J., Shawen, S., Liochev, S. I., and Crapo, J. D. (1995) J. Pharmacol. Exp. Ther. 275, 1227-1232[Abstract/Free Full Text]
  22. Kelso, G. F., Porteous, C. M., Coulter, C. V., Hughes, G., Porteous, W. K., Ledgerwood, E. C., Smith, R. A., and Murphy, M. P. (2001) J. Biol. Chem. 276, 4588-4596[Abstract/Free Full Text]
  23. Balla, G., Jacob, H. S., Balla, J., Rosenberg, M., Nath, K., Apple, F., Eaton, J. W., and Vercellotti, G. M. (1992) J. Biol. Chem. 267, 18148-18153[Abstract/Free Full Text]
  24. Chitambar, C. R., and Seligman, P. A. (1986) J. Clin. Invest. 78, 1538-1546[Medline] [Order article via Infotrieve]
  25. Heinloth, A., Brüne, B., Fischer, B., and Galle, J. (2002) Atherosclerosis 162, 93-101[CrossRef][Medline] [Order article via Infotrieve]
  26. Kotamraju, S., Konorev, E. A., Joseph, J., and Kalyanaraman, B. (2000) J. Biol. Chem. 275, 33585-33592[Abstract/Free Full Text]
  27. Kojima, H., Sakurai, K., Kikuchi, K., Kawahara, S., Kirino, Y., Nagoshi, H., Hirata, Y., and Nagano, T. (1998) Chem. Pharm. Bull. 46, 373-375
  28. Nakatsubo, N., Kojima, H., Kikuchi, K., Nagoshi, H., Hirata, Y., Maeda, D., Imai, Y., Irimura, T., and Nagano, T. (1998) FEBS Lett. 427, 263-266[CrossRef][Medline] [Order article via Infotrieve]
  29. Kennedy, M. C., Emptage, M. H., Dreyer, J.-L., and Beinert, H. (1983) J. Biol. Chem. 258, 11098-11105[Abstract/Free Full Text]
  30. Tietze, F. (1969) Anal. Biochem. 27, 502-522[CrossRef][Medline] [Order article via Infotrieve]
  31. Coux, O., Tanaka, K., and Goldberg, A. L. (1996) Annu. Rev. Biochem. 65, 801-847[CrossRef][Medline] [Order article via Infotrieve]
  32. Pajonk, F., Riess, K., Sommer, A., and McBride, W. H. (2002) Free Radic. Biol. Med. 32, 536-543[CrossRef][Medline] [Order article via Infotrieve]
  33. Rota, C., Fann, Y. C., and Mason, R. P. (1999) J. Biol. Chem. 274, 28161-28168[Abstract/Free Full Text]
  34. Zhao, H., Kalivendi, S., Zhang, H., Joseph, J., Nithipatikom, K., Vasquez-Vivar, J., and Kalyanaraman, B. (2003) Free Radic. Biol. Med. 34, 1359-1368[CrossRef][Medline] [Order article via Infotrieve]
  35. Pantopoulos, K., Mueller, S., Atzberger, A., Ansorge, W., Stremmel, W., and Hentze, M. W. (1997) J. Biol. Chem. 272, 9802-9808[Abstract/Free Full Text]
  36. Hentze, M. W., Rouault, T. A., Harford, J. B., and Klausner, R. D. (1989) Science 244, 357-359[Abstract/Free Full Text]
  37. Rouault, T. A., and Klausner, R. D. (1996) Trends Biochem. Sci. 21, 174-177[CrossRef][Medline] [Order article via Infotrieve]
  38. Sorescu, D., Weiss, D., Lassegue, B., Clempus, R. E., Szocs, K., Sorescu, G. P., Valppu, L., Quinn, M. T., Lambeth, J. D., Vega, J. D., Taylor, W. R., and Griendling, K. K. (2002) Circulation 105, 1429-1435[Abstract/Free Full Text]
  39. Deshpande, S. S., Qi, B., Park, Y. C., and Irani, K. (2003) Arterioscler. Thromb. Vasc. Biol. 23, e1-e6[Abstract/Free Full Text]
  40. Garcia-Ruiz, C., Colell, A., Mari, M., Morales, A., and Fernandez-Checa, J. C. (1997) J. Biol. Chem. 272, 11369-11377[Abstract/Free Full Text]
  41. Phillips, D. C., Allen, K., and Griffiths, H. R. (2002) Arch. Biochem. Biophys. 407, 15-24[CrossRef][Medline] [Order article via Infotrieve]
  42. McCarty, M. F. (1998) Med. Hypotheses. 51, 465-476[CrossRef][Medline] [Order article via Infotrieve]
  43. Lenaz, G., Bovina, C., D'Aurelio, M., Fato, R., Formiggini, G., Genova, M. L., Giuliano, G., Pich, M. M., Paolucci, U., Castelli, G. P., and Ventura, B. (2002) Ann. N. Y. Acad. Sci. 959, 199-213[Medline] [Order article via Infotrieve]
  44. Schuler, F., and Casida, J. E. (2001) Biochim. Biophys. Acta 1506, 79-87[Medline] [Order article via Infotrieve]
  45. Sparagna, G. C., Hickson-Bick, D. L., Buja, L. M., and McMillin, J. B. (2000) Am. J. Physiol. 279, H2124-H2132
  46. Cutler, R. G., Pedersen, W. A., Camandola, S., Rothstein, J. D., and Mattson, M. P. (2002) Ann. Neurol. 52, 448-457[CrossRef][Medline] [Order article via Infotrieve]
  47. Puglielli, L., Ellis, B. C., Saunders, A. J., and Kovacs, D. M. (2003) J. Biol. Chem. 278, 19777-19783[Abstract/Free Full Text]
  48. Ariga, T., Jarvis, W. D., and Yu, R. K. (1998) J. Lipid Res. 39, 1-16[Abstract/Free Full Text]
  49. Hui, D. Y., and Howles, P. N. (2002) J. Lipid Res. 43, 2017-2030[Abstract/Free Full Text]
  50. Irizarry, M. C. (2003) Ann. Neurol. 54, 7-8[CrossRef][Medline] [Order article via Infotrieve]
  51. Mendes Ribeiro, A. C., Brunini, T. M., Ellory, J. C., and Mann, G. E. (2001) Cardiovasc. Res. 49, 697-712[Abstract/Free Full Text]

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