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Am J Physiol Heart Circ Physiol 289: H852-H861, 2005. First published April 1, 2005; doi:10.1152/ajpheart.00015.2005
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Oxidized LDL induces mitochondrially associated reactive oxygen/nitrogen species formation in endothelial cells

Jaroslaw W. Zmijewski,1,2,* Douglas R. Moellering,2,* Claire Le Goffe,1,2 Aimee Landar,1,2 Anup Ramachandran,2 and Victor M. Darley-Usmar1,2

1Center for Free Radical Biology and 2Department of Pathology, University of Alabama, Birmingham, Alabama

Submitted 7 January 2005 ; accepted in final form 21 March 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Exposure of cells to complex mixtures of oxidized lipids such as those found in oxidized low-density lipoprotein (oxLDL) induce reactive oxygen and nitrogen species (ROS/RNS) formation. The source of the ROS/RNS within cells is unknown; it is thought they may be involved in redox cell signaling. Although this possibility was initially overlooked, it is becoming clear that mitochondria, which are a source of superoxide and hydrogen peroxide, may play a critical role in the response of cells on exposure to oxidized lipids. In this study, we tested the possibility that mitochondria are a potential source of oxLDL-dependent formation of ROS/RNS in endothelial cells. Using confocal microscopy, we demonstrated that a significant proportion of oxLDL-dependent dichlorodihydrofluorescein (DCF) fluorescence is colocalized to mitochondria. In support of this concept, rho0 endothelial cells showed a substantial decrease in ROS/RNS formation stimulated by oxLDL. In contrast, mostly nonmitochondrial DCF fluorescence was detected in cells exposed to an extracellular source of hydrogen peroxide. The exposure of cells to a nitric oxide synthase inhibitor and urate resulted in a decrease in oxLDL-induced DCF fluorescence that was restored by addition of nitric oxide donors to the medium. Taken together, these results suggest that oxLDL-dependent DCF fluorescence is mitochondrially associated and may be due to the formation of peroxynitrite.

reactive oxygen species; reactive nitrogen species; mitochondria; low-density lipoprotein; atherosclerosis

OXIDIZED LIPIDS FORMED BY either nonspecific or enzymatic pathways have the potential to activate a diverse series of cell signaling events in a broad variety of cells (8, 31, 56). This is particularly important in the field of atherosclerosis, where a strong body of evidence supports the concept that production of oxidants, most likely hydrogen peroxide, superoxide, and peroxynitrite, are enhanced during the atherosclerotic process (21, 46, 49). Furthermore, it has been shown that both complex mixtures of oxidized lipids and specific individual molecules, particularly reactive electrophiles, are capable of inducing cell signaling events (18, 20, 26, 27, 33, 37). Several groups of enzymes can generate reactive oxygen and nitrogen species (ROS/RNS) in cells including the NADPH oxidases, nitric oxide (NO) synthase (NOS), and mitochondria (46, 49). A role for mitochondria in this regard has not been examined in the vasculature. It is now becoming evident that this organelle can transduce a number of oxidative and nitrosative signals through mechanisms that remain to be defined (6, 7, 10, 11, 48).

Although it has been shown that oxidized low-density lipoprotein (oxLDL) increases the intracellular formation of ROS/RNS (13, 24), the source of these active species remains uncertain. This is important, because oxLDL can initiate signaling cascades including activation of NF-{kappa}B and the MAP kinases, which are known to involve redox changes in cells (14, 20, 35, 39–41). Several studies of cultured cells have established that oxLDL is an important factor that enhances arterial apoptosis with involvement of both the mitochondrial and death receptor pathways (Fas/Fas ligand, tumor necrosis factor receptors I and II) and oxidative stress (30, 41). For example, oxLDL induces cellular apoptosis in a mechanism dependent upon ROS generation in vascular smooth muscle cells (24). In addition, adenovirus-mediated overexpression of catalase attenuates oxLDL-induced apoptosis in human aortic endothelial cells (35). In contrast, oxLDL can also contribute to the adaptive antiatherogenic responses of endothelium through the transcriptional regulation of antioxidant enzymes (2, 23, 36). Mitochondrial involvement is implied from the finding that nontoxic concentrations of oxLDL lead to induction of manganese superoxide dismutase and complex I (9, 50). Many of these responses can be recapitulated using defined lipid oxidation products such as the electrophilic cyclopentenones (25, 33). Thus it becomes important to establish the source of ROS/RNS formation on exposure to oxidized lipids, because it is likely that this is a major determinant in both the adaptive and cytotoxic responses of cells under conditions of chronic inflammation (32, 36). Early studies of the role of NO in atherosclerosis indicated that in endothelial cells, its reaction with superoxide contributed to vascular dysfunction in the disease (22, 46). The product of the reaction of NO with superoxide or peroxynitrite at high concentrations can lead to damage to the vasculature; but lower levels, such as those formed by endothelial NOS (eNOS), may lead to the activation of cell signaling pathways (46). An interesting example is the activation of the c-Jun NH2-terminal kinase by oxLDL, which requires activation of eNOS and formation of peroxynitrite (20). Recent data provide additional support for a link between cellular ROS formation, NO, and mitochondria with the findings that a proportion of the NOS in endothelial cells may be mitochondrially associated, and NO can modulate mitochondrial function including ROS formation (45, 52).

In the present study, we hypothesized that oxLDL promotes the formation of mitochondrial ROS/RNS via the electron transport chain. To address these questions, we applied a 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA) fluorogenic probe to measure oxLDL-induced ROS/RNS generation in endothelial cells. Several studies with this probe have shown that it is capable of detecting both hydrogen peroxide and peroxynitrite (15, 38). Interestingly, detection of hydrogen peroxide within cells occurs through a mechanism dependent upon intracellular ROS and availability of iron (54). This probe was also used to examine intracellular localization of ROS/RNS generation induced by oxLDL via confocal scanning microscopy. We show that the oxLDL-dependent DCF fluorescence is mitochondrially associated and requires NO derived from eNOS. Our data suggest that mitochondria are capable of transducing the signal from oxLDL to ROS/RNS through the action of the respiratory chain.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. We obtained 4,4,4-trifluoro-1-(2-thienyl)-1,3-butanedione (TTFA), myxothiazol, rotenone, glucose oxidase (GO), and N{omega}-nitro-L-arginine (L-NNA) from Sigma (St. Louis, MO). MitoTracker deep red 633, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazol-carbocyanine iodide (JC-1 probe), DCFH-DA, coelenterazine, Hoechst-33258, and anti-oxPhos complex IV subunit I MAb were purchased from Molecular Probes (Eugene, OR). N,N'-bis(2-hydroxybenzyl)ethylenediamine-N,N'-diacetic acid dihydrochloride dihydrate (HBED) was purchased from BioMol (Plymouth Meeting, PA); (Z)-1-[2-(2-aminoethyl)-N-(2-ammonioethyl)amino]diazen-1-ium-1,2-diolate (DETA-NONOate) was from Alexis (San Diego, CA); and diethylamine NONOate-AM (DEA-AM) was from Calbiochem (San Diego, CA). All other reagents used were of analytical grade.

Cell culture. Bovine aortic endothelial cells (BAECs) previously harvested from descending thoracic aortas were obtained and maintained (at 37°C in 5% CO2) in DMEM growth medium (GIBCO) that contained glutamine (4 mM), pyruvate (1 mM), sodium bicarbonate (3.7 g/l), glucose (1 g/l), and fetal calf serum (10%; Atlanta Biologicals; Norcross, GA) with penicillin (100 U/ml) and streptomycin (100 ng/ml). Cells used in this study were between passages 4 and 12 (20). GO was used as a source of extracellular hydrogen peroxide formation as described by Tampo et al. (54) without additional supplementation of the medium with glucose in a volume of 0.5 ml. To create mitochondrial DNA-depleted (rho0) cells, BAECs were incubated in DMEM growth medium supplemented with ethidium bromide (250 ng/ml), uridine (50 µg/ml), and sodium pyruvate (110 µg/ml) for 7–14 days (11). Control and rho0 cells treated with oxLDL were between passages 8 and 12. Additionally, membrane potential ({Delta}{psi}) and protein levels of complex IV subunit I of rho0 cells were determined.

Isolation and oxidation of LDL. Human LDL was isolated from plasma of healthy donors by differential centrifugation using a method previously described (12). The concentration of LDL protein was determined using Bradford protein assay reagent (Bio-Rad). All native (nLDL) or oxLDL concentrations used in the experiments were normalized to this protein concentration. For oxidation, LDL (1–2 mg/ml) was dialyzed against sterile PBS and diethylenetriamine pentaacetic acid (DTPA, 10 µM) and incubated with oxidant CuSO4 (25 µM) for 14–16 h at 37°C. The oxidation reaction was stopped by the addition of DTPA (100 µM). The relative electrophoretic mobility of LDL was determined on agarose gels using the lipoprotein electrophoresis system supplied by Beckman following the manufacturer's instructions and was typically between 1.2 and 1.8. A control sample was prepared (PBS, 10 µM DTPA, and 25 µM CuSO4) and added to the cells under identical conditions to oxLDL treatment.

Imaging of DCF fluorescence. Cells were grown in a four-well chambered coverglass (Nalge; Naperville, IL) until confluent, incubated with DCFH-DA (20 µM) for 60 min, and subsequently treated with various concentrations of oxLDL for 30 min at 37°C. Cells were also incubated with MitoTracker deep red 633 (0.5 µM) for 15 min before being imaged. To examine the effects of an iron chelator, the peroxynitrite scavenger urate, or mitochondrial complex inhibitors, cells were preincubated with HBED for 60 min or urate, rotenone, or TTFA for 30 min before application of oxLDL. Cells were washed twice with culture medium, and images were acquired from three or more randomly chosen fields using inverted epifluorescence microscopy (model IX70; Olympus). The levels of DCF fluorescence were quantitated and displayed as three-dimensional surface scans using SimplePCI software (Compix; Cranberry Township, PA). The subcellular source of ROS generation was assessed by single bidirectional scans of live cells using a Leica DMIRBE inverted epifluorescence/Nomarski microscope outfitted with Leica TCS NT laser confocal optics. Images were acquired using a x100 oil-immersion objective and a pinhole setting of 0.2 Airy units. Laser excitation was set (5–10%) to produce as little photooxidation of the dye as possible. Images were merged and processed using IPLab Spectrum and Adobe Photoshop (Adobe Systems) software.

Measurement of mitochondrial {Delta}{psi}. The mitochondrial inner membrane electrochemical {Delta}{psi} was assessed using epifluorescence microscopy with 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazol-carbocyanine iodide (JC-1). Briefly, BAECs or rho0 cells were grown in a four-well chambered coverglass until 90–100% confluence. Live cells were then incubated in culture medium supplemented with JC-1 for 20 min and were further incubated with Hoechst-33258 (2.5 µg/ml) for 10 min at 37°C to identify nuclei. Next, cells were washed with serum complete medium, and images were acquired from three or more randomly chosen fields using an inverted epifluorescence microscope with suitable filter setup (model IX70; Olympus). The levels of JC-1 red and green fluorescence were measured, and nuclei were counted in each captured image using SimplePCI software. Quantitative data were expressed as total red channel pixel intensity per number of nuclei. Images were merged and processed as described (see Imaging of DCF fluorescence).

Measurement of ROS using coelenterazine. Cells were detached from confluent monolayers with trypsin, treated for 30 min with various specific pharmacological inhibitors, and then treated and gently mixed for an additional 30 min with or without oxLDL (75 µg/ml) (55). For measurement of ROS, coelenterazine (10 µM) was added to cells (2–5 x 106 cells/ml) resuspended in PBS supplemented with Ca2+ (1 mM) and glucose (1 g/l). Chemiluminescence was monitored for 2 min using an AutoLumat LB-953 luminometer.

Treatment with L-NNA and NO donors. BAECs were grown to confluence in a four-well chambered coverglass and incubated with DCFH-DA (20 µM) for 60 min and in the presence of L-NNA (250 µM) for 30 min before treatment with various concentrations of oxLDL (75–150 µg/ml) for the next 30 min. In the case of NO donors, media were supplemented with DETA-NONOate (500 µM) or DEA-AM (2 µM) during incubation of untreated cells and cells treated with L-NNA or L-NNA with oxLDL. Cells were then washed twice with serum-containing medium, and images from three or more randomly chosen fields were acquired.

Statistical analysis. All experiments were performed independently three or more times. Data are expressed as means ± SE and were analyzed for statistical significance using unpaired Student's t-test. A P value of <0.05 was considered significant.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
OxLDL stimulates ROS production in BAECs. Several studies (13, 24, 44) have demonstrated that treatment of vascular cells with oxLDL stimulates ROS/RNS formation as measured by an increase in DCF fluorescence. However, the molecular species within the oxLDL particle that causes these effects is unknown as is the source of the intracellular oxidants. In the first series of experiments, the ability of oxLDL to induce ROS generation in endothelial cells was confirmed. BAECs were incubated with DCFH-DA (20 µM) for 60 min and then with oxLDL (0–200 µg/ml) or nLDL (200 µg/ml) for 30 min. After this period, the medium was changed, and images were acquired (Fig. 1A). DCF fluorescence intensity was measured and is expressed as the average pixel intensity per cell (Fig. 1, B and C). The DCF fluorescence intensity was markedly increased in cells as a function of oxLDL concentration, whereas nLDL or the copper-DTPA control had no effect.



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  		Fig. 1. Effects of native and oxidized low-density lipoproteins (nLDL and oxLDL, respectively) on reactive oxygen species (ROS)-induced oxidation of dichlorodihydrofluorescein (DCF). A: bovine aortic endothelial cells (BAECs) in four-well chambers were loaded with 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA, 20 µM) for 60 min before exposure to nLDL or oxLDL (200 µg/ml) for 30 min. Live cells were imaged on an inverted fluorescence microscope (top). Levels of DCF fluorescence in captured images were then processed to three-dimensional (3-D) surface scans (bottom). B: average DCF fluorescence intensity values for cells treated as in A. Values are means ± SE; n = 3. *P < 0.001 compared with control (CTL). C: oxLDL concentration-dependent increase in DCF fluorescence; n = 3. *P < 0.005 compared with CTL.

 
It was recently shown (54) that DCF fluorescence stimulated by exogenous hydrogen peroxide can be inhibited by the addition of a cell-permeable iron chelator such as HBED. HBED (100 µM) caused a substantial decrease in the level of oxLDL-dependent DCF fluorescence as was also found with external generation of hydrogen peroxide by GO (Fig. 2).



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  		Fig. 2. N,N'-bis(2-hydroxybenzyl)ethylenediamine-N,N'-diacetic acid dihydrochloride dihydrate (HBED)-dependent inhibition of oxLDL- or glucose oxidase (GO)-induced ROS generation. Cells were incubated with DCFH-DA in the presence or absence of HBED (100 µM) for 30 min before treatment with oxLDL (150 µg/ml) or GO (5 mU) for 30 min. A: live cells were washed with serum-containing medium and imaged on an inverted epifluorescence microscope. B: quantitative data for oxLDL or GO-induced DCF fluorescence in the presence or absence of HBED; n = 3 or 4. *P < 0.001 compared with oxLDL or GO alone.

 
Impact of inhibitors of electron transport chain complexes and oxLDL-dependent DCF fluorescence. A potential source of both hydrogen peroxide and iron in cells is the respiratory complexes in the mitochondria. To demonstrate the capability of DCF fluorescence to detect mitochondrially derived ROS, we determined the effects of rotenone, which is an inhibitor of complex I, on levels of endogenous ROS generation. Incubation with rotenone (10 µM) alone caused an increase in DCF fluorescence compared with control cells and was ~60% of that found with oxLDL (Fig. 3A). Because the experiments of rotenone in combination with oxLDL proved difficult to interpret, we used other inhibitors to determine the potential contribution of respiratory chain complexes to oxLDL-dependent DCF fluorescence. In contrast with rotenone, TTFA (10 µM) alone did not increase DCF fluorescence (Fig. 3, A and B). Preincubation of BAECs with TTFA (10 µM) and subsequent exposure of cells to oxLDL (75 µg/ml) showed an 86% decrease in DCF fluorescence compared with treatment with oxLDL alone (Fig. 3, A and B). It was reported (59) that TTFA can inhibit esterases. This mechanism could prevent DCF from accumulating in cells or mitochondria and so account for inhibition of the oxLDL-dependent increase in DCF fluorescence. As a control for this property, cells were preincubated with TTFA in the presence of GO (5 mU) as a nonmitochondrial source of ROS. TTFA had no effect on GO-dependent DCF fluorescence (Fig. 3C), thereby ruling out an artifactual contribution of esterases to the data shown in Fig. 3B.



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  		Fig. 3. Effects of 4,4,4-trifluoro-1-(2-thienyl)-1,3-butanedione (TTFA) on oxLDL- or GO-induced ROS generation. BAECs were cultured with DCFH-DA for 60 min in the presence or absence of mitochondrial complex II inhibitor TTFA (10 µM). Cells were then exposed to oxLDL (75 µg/ml) or GO (5 mU) for 30 min. A: images were acquired. Compared with untreated cells, DCF fluorescence of cells exposed to rotenone (10 µM) for 60 min increased. B and C: levels of DCF fluorescence were averaged as pixel intensity per cell; n = 3. *P < 0.01 compared with oxLDL alone (B) or with CTL (C).

 
OxLDL-induced ROS/RNS generation measured with coelenterazine. As a mechanistically and structurally distinct alternative to DCF, we used the chemiluminescent probe coelenterazine. This molecule does not require peroxidase or iron for its mode of action and reacts with both superoxide and peroxynitrite (55). It is likely that mitochondria initially generate superoxide that is then converted to hydrogen peroxide through the enzymatic action of superoxide dismutase or, in the presence of NO, can be converted to peroxynitrite. As found with the use of DCF as a probe, a dramatic increase in ROS production was evident with coelenterazine after 30 min of treatment with oxLDL (Fig. 4A). The complex II inhibitors TTFA and 3-nitropropionic acid alone did not increase ROS formation but completely prevented the oxLDL-dependent signal (Fig. 4B).



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  		Fig. 4. TTFA or 3-nitropropionic acid (3-NP) inhibits oxLDL-induced ROS generation as measured with coelenterazine probe. A: representative tracing of ROS production in BAECs incubated with or without oxLDL (75 µg/ml). B: data from A illustrated as relative light units (RLU) per 106 cells per minute, and data from BAECs after incubation with TTFA (10 µM) or 3-NP acid (10 µM) in the presence or absence of oxLDL (75 µg/ml); n = 3–5. #P < 0.05 compared with CTL cells; *P < 0.05 compared with treatment alone.

 
NOS contributes to oxLDL-induced ROS/RNS generation. To investigate whether endogenous sources of NO contribute to oxLDL-induced DCF fluorescence, cells were exposed to L-NNA, which is an inhibitor of NOS. Preincubation with L-NNA (250 µM) caused a decrease in oxLDL-induced DCF fluorescence, whereas L-NNA alone had no effect (Fig. 5). To rule out nonspecific effects of L-NNA, two structurally distinct NO donors, DETA-NONOate or cell-permeable DEA-AM, were used in the presence of the NOS inhibitor and oxLDL. Quantitative analysis indicated that both NO donors alone had minimal or no effect on DCF fluorescence but were able to significantly restore the oxLDL-dependent DCF fluorescence prevented by the NOS inhibitor (Fig. 5, B and C). Next, to examine whether the increase in mitochondrial ROS production was associated with peroxynitrite formation, BAECs were incubated in the presence or absence of urate (100 µM) for 30 min and subsequently exposed to oxLDL (75 µg/ml) for 30 min. Figure 6 shows a significant decrease (nearly 60%) in oxLDL-induced DCF fluorescence in cells treated with urate, whereas treatment with urate alone had no effect.



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  		Fig. 5. A: inhibitory effects of N{omega}-nitro-L-arginine (L-NNA) on nitric oxide synthase (NOS)- and oxLDL-induced ROS generation. Cells were incubated with DCFH-DA (20 µM) for 30 min before treatment with or without L-NNA (250 µM) for 30 min. B and C: to reverse the effect of L-NNA, cells were also treated with NO donors (Z)-1-[2-(2-aminoethyl)-N-(2-ammonioethyl)amino]diazen-1-ium-1,2-diolate (detaNO, 500 µM) and diethylamine NONOate-AM (DEA/AM, 2 µM). Next, cells were exposed to oxLDL (A and B, 150 µg/ml; C, 75 µg/ml); n = 3. *P < 0.01 compared with oxLDL alone; #P < 0.01 compared with L-NNA + oxLDL. Cells were then washed, and images were acquired (A) as described (see MATERIALS AND METHODS).

 


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  		Fig. 6. Effects of urate (UA) on oxLDL-induced DCF oxidation. BAECs loaded with DCFH-DA were preincubated with urate (100 µM) for 30 min before treatment with oxLDL (75 µg/ml) for 30 min. A: representative images acquired with epifluorescence microscopy show decreased oxLDL-induced DCF fluorescence in cells preincubated with urate compared with oxLDL treatment alone. B: level of DCF fluorescence was measured in images acquired from three randomly chosen fields and is expressed as pixel intensity per cell; n = 3–5. *P < 0.05 compared with cells treated with oxLDL alone.

 
OxLDL induces generation of ROS in mitochondria. To further confirm the mitochondrial localization of ROS generation by oxLDL, we used confocal fluorescence microscopy. BAECs were loaded with DCFH-DA and treated with GO (5 mU for 5–10 min) as a nonmitochondrial source of hydrogen peroxide or treatment with myxothiazol (a complex III inhibitor) or oxLDL for 30 min. Cells were also treated with the mitochondria-specific probe MitoTracker for 15 min before imaging. As shown in Fig. 7, the various treatments caused an increase in the level of DCF fluorescence compared with control cells. Merging of the DCF signal with the MitoTracker images (myxothiazol- or oxLDL-treated cells) indicates that the increase in DCF fluorescence colocalized to the mitochondria (Fig. 7, yellow areas). In contrast, in the case of GO, the DCF fluorescence was predominantly localized to the cytosol.



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  		Fig. 7. OxLDL-induced generation of ROS in mitochondria. BAECs grown in four-well chambers were loaded with DCFH-DA (20 µM) for 60 min before an exposure to GO (5 mU; 5–10 min), myxothiazol (5 µM; 30 min), and oxLDL (200 µg/ml; 30 min). Cells were incubated with MitoTracker (0.5 µM) for 15 min before being imaged. Images were acquired using a confocal scanning microscope (see MATERIALS AND METHODS). Exposure of BAECs to myxothiazol and oxLDL revealed an increase in intensity of DCF fluorescence (green) that colocalized with a mitochondrial marker (yellow). In contrast, mostly nonmitochondrial DCF fluorescence was present in cells exposed to GO.

 
Decrease of oxLDL-induced ROS formation in rho0 cells. To further evaluate the contribution of mitochondria to oxLDL-induced ROS formation, mitochondrial DNA was depleted in endothelial cells by chronic exposure to ethidium bromide. This treatment generates the equivalent of a rho0 cell phenotype as evidenced by a substantial decrease in the mitochondrially coded subunit I of cytochrome c oxidase (Fig. 8). In addition, the mitochondrial {Delta}{psi} in rho0 cells was measured using the JC-1 probe. JC-1 selectively enters mitochondria and aggregates when {Delta}{psi} values exceed 80–100 mV, thereby causing a shift in fluorescence from 530 (green) to 590 (red) nm (53). The JC-1 probe showed intense red fluorescence in control cells, and this was almost completely attenuated (>10-fold decrease) in rho0 cells (Fig. 8, A and B), which indicates a loss of mitochondrial {Delta}{psi} in the rho0 cells. On exposure to oxLDL, there was a significant decrease in DCF fluorescence that was evident after treatment with ethidium bromide for both 7 and 14 days (Fig. 9).



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  		Fig. 8. Characterization of mitochondrial DNA-depleted rho0 cells. A: representative images show the mitochondrial membrane potential ({Delta}{psi}) of CTL and rho0 cells visualized using 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazol-carbocyanine iodide (JC-1) with epifluorescence microscopy. Mitochondrial {Delta}{psi} is indicated by JC-1 red fluorescence (top). Relative change in mitochondrial {Delta}{psi} is demonstrated by the shift from red to green fluorescence shown in the merged image (middle). Nuclei of live cells were stained blue with Hoechst-33258 (bottom). B: JC-1 red fluorescence in rho0 and CTL cells was calculated as pixel intensity per number of nuclei obtained from images acquired from three randomly chosen fields. C: Western blot analysis demonstrates the decrease in complex IV subunit I protein level in total cell lysate of rho0 compared with CTL cells (top). Quantitation of complex IV subunit I is expressed as a percentage of CTL (bottom); n = 3. *P < 0.01 compared with CLT.

 


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  		Fig. 9. Effects of oxLDL on DCF oxidation in rho0 cells. CTL and ethidium bromide-treated (for 7–14 days) rho0 cells were loaded with DCFH-DA before incubation with or without oxLDL (75 µg/ml). Cells were then washed with serum-containing media and images were acquired (see MATERIALS AND METHODS). A: representative images demonstrate decreased oxLDL-induced DCF fluorescence in rho0 cells compared with oxLDL-treated CTL cells. B: levels of DCF fluorescence acquired from three random fields was expressed as pixel intensity per cell; n = 3. *P < 0.05 compared with oxLDL-treated CTL cells.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Various extracellular stimuli (including oxidized lipids) can cause an increase in the production of ROS/RNS within cells, which causes a shift in redox balance and subsequent initiation of downstream signaling cascades (18, 34, 56). It is now thought that mitochondria can contribute to redox cell signaling, and the mitochondrial ROS formation has the potential to be controlled (5, 6, 42). Several processes contribute to the regulation of mitochondrial ROS formation including S-thiolation of complex I and the activity of uncoupler proteins, which in turn may be regulated by oxidized lipids (4). The mitochondrial antioxidant enzyme manganese superoxide dismutase may also control intramitochondrial ROS formation. This is of particular interest, because the superoxide-dependent liberation of iron in the organelle from metalloproteins is an additional potential mechanism through which mitochondrial ROS may contribute to redox signaling (47, 57). A recent series of experiments shows compelling evidence for the regulation of metalloproteinase expression via a MAP kinase-dependent signaling pathway that is modulated by the production of superoxide in the matrix of mitochondria (29). Thus mitochondria are not only involved in the proapoptotic pathways but also in the more subtle adaptive responses that can be mediated through mitochondrial production of ROS.

OxLDL was shown to increase ROS generation in fibroblasts (1), and an oxLDL-dependent increase in hypoxia-inducible factor-1{alpha} in human fibroblasts was also dependent on ROS production (51). OxLDL has also been shown to induce apoptosis in vascular smooth muscle cells via generation of ROS (24), and binding of oxLDL to the lectin-like oxLDL receptor-1 on endothelial cells resulted in an increase in intracellular ROS generation (14). Interestingly, the mitochondrially targeted ubiquinone analog mitoQ has been shown to inhibit apoptosis caused by hydrogen peroxide, which suggests a role for secondary production of ROS by mitochondria in oxidant-induced apoptosis (3, 17, 28).

In this study, we investigated mitochondria as a potential source for ROS/RNS formed on exposure of cells to oxLDL. In the case of oxidized lipids, both the complex mixtures found in oxLDL and specific aldehydes are capable of inducing ROS/RNS formation by indirect methods such as DCF fluorescence. The use of DCF as an indicator of ROS/RNS formation is open to criticism, and the data obtained with such molecules must be interpreted with caution. The mechanisms of DCF fluorescence have been defined in considerable detail, and it is evident that the signals obtained from these experiments cannot by uniquely ascribed to a single mechanism. For example, recent studies (54) have defined a contribution from extracellular iron or the presence of peroxidases within cells and depletion of GSH. One important advantage of this technique, however, is that it offers the possibility of localizing the source of DCF fluorescence within the cell.

To test for a mitochondrial contribution to the increased ROS/RNS production after oxLDL treatment, cells were treated in conjunction with MitoTracker. OxLDL showed a strong mitochondrially localized formation of DCF fluorescence, whereas hydrogen peroxide generated from GO did not. A mitochondrial origin for this signal was confirmed by its inhibition using the complex II inhibitor TTFA. As an alternative approach to using DCF, we also utilized the structurally distinct chemiluminescent probe coelenterazine, which has a different mechanism of action for detection of ROS/RNS (55). The results from these experiments were entirely consistent with those obtained using DCF and indicate a mitochondrial contribution to ROS formation on exposure to oxLDL involving complex II. As an additional approach, we prepared rho0 cells by extended exposure to ethidium bromide treatment. These cells remained viable but showed decreases in both mitochondrial {Delta}{psi} and levels of a mitochondrially coded protein. When treated with oxLDL, the DCF fluorescence was significantly attenuated, which supports a major contribution of mitochondria to ROS formation.

In a previous study (20), we showed that oxLDL can activate eNOS, and this leads to activation of the MAP kinase c-Jun NH2-terminal kinase through the intermediate formation of peroxynitrite. Recently, an association of eNOS with mitochondria has been reported in endothelial cells, and this raises the possibility that DCF fluorescence at the mitochondrial level may be due to peroxynitrite (16, 19). Interestingly, inhibition of NOS using L-NNA abrogated the oxLDL-induced ROS formation. Subsequently, exposure of cells to an external source of NO in the presence of NOS inhibitor restored the oxLDL-dependent DCF fluorescence; this is consistent with peroxynitrite formation. In support of this conclusion, pretreatment with urate also caused a substantial decrease in oxLDL-induced fluorescence.

The precise site within mitochondria that causes the increased DCF fluorescence remains to be defined. Interestingly, complex II has been implicated in the generation of ROS in hyperglycemia (43, 58). The data obtained with the complex II inhibitor TTFA would be consistent with either a direct contribution from this respiratory complex to ROS formation or, alternatively, reversed electron transport through complex I of the respiratory chain. A number of proteins within mitochondria including complex II and aconitase could also act as sources of iron that could then contribute to DCF fluorescence.

In summary, we have shown that oxLDL is capable of inducing ROS/RNS formation from mitochondria through a mechanism that requires iron, eNOS activation, and complex II activity. The precise mechanisms that lead to increased ROS formation in mitochondria or the consequences for this interaction to adaptation or cell death remain to be defined.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by National Institutes of Health Grants ES-10167 and HL-58031 (to V. Darley-Usmar).


   ACKNOWLEDGMENTS
 
Present address of D. R. Moellering: Dept. of Nutrition, University of Alabama at Birmingham.

Present address of A. Ramachandran: Wellcome Trust Laboratory, Dept. of Gastrointestinal Sciences, Christian Medical College, Vellore 632004, India.


   FOOTNOTES
 

Address for reprint requests and other correspondence: V. Darley-Usmar, Dept. of Pathology, Univ. of Alabama at Birmingham, Biomedical Research Bldg. II, 901 19th St. South, Birmingham, AL 35294 (E-mail: Darley{at}path.uab.edu)

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.

* J. W. Zmijewski and D. R. Moellering contributed equally to this work. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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