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(Circulation Research. 2000;86:960.)
© 2000 American Heart Association, Inc.

Cellular Biology

Hydrogen Peroxide– and Peroxynitrite-Induced Mitochondrial DNA Damage and Dysfunction in Vascular Endothelial and Smooth Muscle Cells

Scott W. Ballinger, Cam Patterson, Chang-Ning Yan, Richard Doan, David L. Burow, Christal G. Young, F. Michael Yakes, Bennett Van Houten, Carol A. Ballinger, Bruce A. Freeman, Marschall S. Runge

From the Sealy Center for Molecular Cardiology and Division of Cardiology (S.W.B., C.P., C.-N.Y., R.D., D.L.B., C.G.Y., C.A.B., M.S.R.) and Sealy Center for Molecular Science (B.V.H.), University of Texas Medical Branch, Galveston, Tex; Department of Biochemistry (F.M.Y.), Vanderbilt University, Nashville, Tenn; and Department of Anesthesiology (B.A.F.), University of Alabama at Birmingham, Ala.

Correspondence to Marschall S. Runge, Cardiology Division and Sealy Center for Molecular Cardiology, University of Texas Medical Branch, Galveston, TX 77555-1064. E-mail mrunge{at}utmb.edu

	Abstract

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Abstract—The mechanisms by which reactive species (RS) participate in the development of atherosclerosis remain incompletely understood. The present study was designed to test the hypothesis that RS produced in the vascular environment cause mitochondrial damage and dysfunction in vitro and, thus, may contribute to the initiating events of atherogenesis. DNA damage was assessed in vascular cells exposed to superoxide, hydrogen peroxide, nitric oxide, and peroxynitrite. In both vascular endothelial and smooth muscle cells, the mitochondrial DNA (mtDNA) was preferentially damaged relative to the transcriptionally inactive nuclear ß-globin gene. Similarly, a dose-dependent decrease in mtDNA-encoded mRNA transcripts was associated with RS treatment. Mitochondrial protein synthesis was also inhibited in a dose-dependent manner by ONOO^-, resulting in decreased cellular ATP levels and mitochondrial redox function. Overall, endothelial cells were more sensitive to RS-mediated damage than were smooth muscle cells. Together, these data link RS-mediated mtDNA damage, altered gene expression, and mitochondrial dysfunction in cell culture and reveal how RS may mediate vascular cell dysfunction in the setting of atherogenesis.

Key Words: reactive oxygen species • reactive nitrogen species • atherosclerosis • oxidative damage

	Introduction

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Although reactive species (RS) serve as second messengers in vascular cells to mediate short-term signaling events,¹ the long-term effects of these radicals induce cellular damage and may potentiate atherosclerotic lesion formation. Numerous studies have linked excess RS generation with vascular lesion formation and functional defects.^{2 3} Increased levels of oxidatively modified proteins and lipoproteins, including oxidized LDL, lipid peroxidation products, nitrated and chlorinated tyrosines, and advanced glycation end products, have been identified in patients with coronary artery disease.^{4 5 6} A role for RS in atherogenesis is also supported epidemiologically by the association between the common risk factors for myocardial infarction—hypertension, hypercholesterolemia, cigarette smoking, and diabetes mellitus—and increased levels of RS.^{7 8 9}

The RS thought to be important in atherosclerosis include superoxide (O₂^·-), peroxynitrite (ONOO^-), hydrogen peroxide (H₂O₂), and nitric oxide (^·NO).¹⁰ Within the arterial wall, metabolic processes, cytokines, and inflammatory responses generate RS in different cell types through multiple and, in some cases, cell type–specific pathways. For example, the oxidative modification of LDL occurs in monocytes only if these cells are activated to express lipoxygenase or induce a respiratory burst with concomitant generation of RS.¹¹ In this case, the cytoplasmic membrane-associated NADPH oxidase of monocytes/macrophages generates RS derived from O₂^·-, H₂O₂, and ^·NO. The plasmalemmal NADH/NADPH oxidase and mitochondria are key sources of RS in heart¹² and vascular tissues.^{13 14 15} Whereas RS generation is possible at multiple sites within the mitochondrial respiratory chain,¹⁶ O₂^·- generation occurs preferentially at the level of coenzyme Q (UQ), through the accumulation of free radical semiquinone anion species (UQ^·-) and electron donation from UQ^·- to O₂.¹⁷ Consistent with this hypothesis, studies have shown that mitochondrial RS generation continues in the presence of rotenone (a complex I inhibitor) as a result of the supply of electrons via complex II (succinate). Antimycin A, which blocks the formation of the UQ^·- at the matrix face and results in the accumulation of UQ^·- on the cytoplasmic face, causes increased O₂^·- formation^{16 18} ; H₂O₂ generated via O₂^·- dismutation (superoxide dismutase [SOD]) reacts in the presence of transition metals (via the Fenton reaction) to yield reactive perferryl and hydroxyl radicals. Alternatively, O₂^·- can react with ^·NO at near diffusion-limited rates (10⁹ [mol/L]^-1 seconds^-1)¹⁹ to form ONOO^- that is directly oxidizing or, on protonization, can react similarly to hydroxyl radical.²⁰

In the present study, human umbilical vein endothelial cells (HUVECs) and human aortic smooth muscle cells (HASMCs) were treated with RS to test the hypothesis that they induce mitochondrial damage in vascular cells. Mitochondrial injury was reflected by DNA damage, changes in gene expression, protein synthesis, and redox function. It was observed that mitochondrial DNA (mtDNA) was preferentially damaged in both cell types, although HUVECs appeared most sensitive to RS treatment. Decreases in mitochondrial RNA (mtRNA) transcripts, mitochondrial protein synthesis, cellular ATP levels, and mitochondrial redox potential were also observed. These results demonstrate RS-mediated mitochondrial damage in vascular cells and suggest a mechanism by which RS induce vascular cell dysfunction, leading to atherogenesis and other chronic vascular diseases.

	Materials and Methods

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Cell Culture
HASMCs were purchased from Clonetics and were maintained as described previously.²¹ HUVECs were cultured under conditions similar to HASMCs, except in M199 and 20% heat-inactivated FCS. Monolayer cultures in 60-mm plates were treated with various RS at 70% to 80% confluence in serum-free, phenol red–free MEM at 37°C, between passages 5 and 7. After treatment, cells were washed with cold PBS and harvested immediately.

RS Treatments
H₂O₂ and ONOO^- were prepared as described.^{22 23} Xanthine oxidase (XO; 5 mU/mL) plus lumazine (LZ; 100 µmol/L) and spermine NONOate (85 µmol/L) were used to generate O₂^·- (2 µmol/L min^–1) and ^·NO (0.5 µmol/L min^–1), respectively. 3-Morpholinosydnonimine hydrochloride (SIN-1; 1 mmol/L, Molecular Probes) was used to generate both O₂^·- and ^·NO (3.7 to 7 µmol/L min^–1).²⁴

DNA Isolation and Quantitative Polymerase Chain Reaction (QPCR)
DNA extraction, QPCR conditions, and calculation of DNA lesion frequencies have been previously described.²² QPCRs were performed in a GeneAmp PCR system 2400 with the GeneAmp XLPCR kit (Perkin-Elmer-Cetus).

Northern Blot Analyses
Total cellular RNA and Northern analyses were performed using standard techniques.²⁵ mtRNA probes were made by PCR using purified mtDNA template (16S rRNA, sense primer, 2005 to 2022, and antisense primer, 2982 to 3001; NADH dehydrogenase 2 [ND2], sense primer, 4831 to 4847, and antisense primer, 5464 to 5481; and cytochrome b [cyt b], sense primer, 14730 to 14749, and antisense primer, 15845 to 15863) and were gel purified before random [-³²P]dCTP labeling. The levels of RNA in each lane were normalized by hybridization with a ß-actin probe.

Mitochondrial Protein Synthesis
Mitochondrial protein synthesis analyses were conducted as previously described.²⁶ Equal amounts of total protein (50 µg) were run on 10% to 20% gradient SDS-PAGE gels. The percentage labeling of translation products was determined by densitometry of all bands for treated and untreated samples. The sum of the labeled bands for each sample was used to calculate relative levels of incorporation.

MTT and ATP Assay
Cells were seeded in 96-well plates at a density of 8000 cells per well and incubated at 37°C for 48 hours. Cells were then treated with RS for 1 hour, washed with PBS, and incubated in conditioned medium for 1 hour with 2.0 µg/mL MTT and lysed, and absorbance was measured at 570 nm.²⁷ Total cellular ATP content was determined by using an ATP determination kit (Molecular Probes, A-6608) and a MicroLumat Plus LB (EG&G Berthold) microinjector luminometer.

Rhodamine (R123) Fluorescence and O₂^·- Production
After 1 hour of RS treatment, cells were trypsinized, washed, pelleted, and resuspended in culture medium containing 1 µmol/L R123 (37°C). R123 fluorescence was determined using a Becton Dickinson FACS flow cytometer. To determine O₂^·- production, aconitase activity was performed,²⁸ with minor modifications for cultured cells. Cells were treated with H₂O₂ for 1 hour and washed with PBS, and aconitase activity was measured for 1 to 60 minutes. NADPH production was normalized to total protein.

	Results

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RS Induce Vascular mtDNA Damage
Vascular cell DNA damage was assessed using a well-validated, sensitive QPCR assay that relies on the principle that oxidative DNA lesions inhibit DNA polymerases (eg, Taq and rTth DNA polymerase). DNA damage was quantified by comparing the relative efficiency of amplification of large fragments of DNA (16.2 kb for the mtDNA; 17.7 kb for the ß-globin gene) and normalizing this to the amplification of smaller (<250-bp) fragments, which have a statistically negligible likelihood of containing damaged bases.

Treatment with H₂O₂ (0.2 mmol/L) for 1 hour resulted (see Table and Figure 1A) in damage to both genomes in HUVECs (P<0.005). Similarly, mtDNA damage occurred in H₂O₂-treated HASMCs (P<0.005); however, no significant nDNA damage was observed. Time course analysis in HUVECs and HASMCs demonstrated that mtDNA damage accumulated rapidly, within 10 and 15 minutes, respectively. The nDNA ß-globin locus accumulated DNA damage less rapidly (Figure 1B), requiring 1 hour of exposure before nDNA damage was observed in HUVECs.

View this table: [in this window] [in a new window]   Table 1. Lesions After Oxidant Exposure

View larger version (24K): [in this window] [in a new window]   Figure 1. RS-induced mtDNA and nDNA damage. A, HUVECs were exposed to 0.2 mmol/L H2O2 for 1 hour and QPCR was performed. Control cultures were incubated in serum-free medium alone. B, Time course of H2O2-induced HASMC and HUVEC injury. Cells were treated with 0.2 mmol/L H2O2 for 0 to 60 minutes, and lesions per 10 kb were estimated. Data are mean levels±SEM. *Significantly different (P<0.05) damage relative to untreated cells.

ONOO^- treatment (0.1 mmol/L and 0.5 mmol/L) caused significant mtDNA damage in HUVECs (P<0.005), whereas only 0.5 mmol/L ONOO^- resulted in HASMCs mtDNA damage (P<0.005, Table). Lower doses of ONOO^- (0.05 mmol/L; data not shown) caused mtDNA damage 30 minutes after treatment in HUVECs (0.17 lesions per 10 kb, P=0.041), which dissipated by 60 minutes (0.01 lesions per 10 kb, P=0.71), implying that the damage had been repaired. HASMCs did not sustain significant damage with low-dose ONOO^- treatment (0.13 lesions per 10 kb at 30 minutes, P=0.069; 0.00 lesions per 10 kb at 60 minutes, P=0.97). No significant levels of nDNA damage were observed in either cell type (Table).

RS-Generating Systems Induce DNA Damage
To examine the effects of sustained, low-dose generation of RS, HUVECs and HASMCs were treated with 5 mU/mL of XO plus LZ (generates 2 µmol/L min^–1 O₂^·-) and/or 85 µmol/L spermine NONOate (generates 0.5 µmol/L min^{–1 ·}NO) to reveal the contributions of O₂^·-, H₂O₂, ^·NO, and ONOO^- to damage. Finally, cells were treated with platelet-derived growth factor (PDGF), which causes endogenous RS production.¹

HUVECs exposed to 5 mU/mL XO plus LZ had increased mtDNA damage (P<0.05, Table), which was inhibited by SOD or catalase (not shown), implicating both H₂O₂ and O₂^·- (XO plus LZ generates both radicals) in mtDNA damage. To assess the influence of ^·NO, cells were treated with a combination of 85 µmol/L spermine NONOate and 5 mU/mL XO plus LZ. ^·NO exposure alone did not cause DNA damage (not shown), but the addition of XO-derived O₂^·- and H₂O₂ caused mtDNA damage in HUVECs (P<0.05, Table). Similarly, SIN-1 treatment (1 mmol/L; generates 3.7 to 7 µmol/L min^–1 O₂^·- and ^·NO) caused significant mtDNA damage in HUVECs (P<0.001, Table and Figure 2). As with other treatments, HASMCs were less affected by RS than HUVECs were (Table, Figure 2).

View larger version (42K): [in this window] [in a new window]   Figure 2. Peroxynitrite-induced mtDNA and nDNA damage in HUVECs and HASMCs. A, Cells were treated for 1 hour with 0.1 or 0.5 mmol/L ONOO-, and QPCR was performed. Control cultures were incubated in serum-free medium alone. B, HUVECs or HASMCs were treated with 1 mmol/L SIN-1, in absence or presence of 300 U/mL SOD.

To assess the effects of endogenous RS, we determined the levels of mtDNA damage in HUVECs and HASMCs at 30 and 60 minutes after 5 ng/mL PDGF treatment. Similar to the 0.05 mmol/L ONOO^- treatment, HUVEC mtDNA damage was increased 30 minutes after PDGF treatment (Table; P<0.001). However, no damage was observed at 60 minutes, indicating that the damage was transient (Table). No significant differences were observed in PDGF-treated HASMCs. Together, these data reveal that HUVECs mtDNA appear more susceptible to both exogenous and endogenous RS-mediated damage than do HASMCs.

Because RS increased mtDNA damage in both HUVECs and HASMCs, their influence on mtRNA transcript levels, mitochondrial protein production, and mitochondrial redox potential was investigated. ONOO^- was chosen on the basis of the data reported here and its documented production in the vasculature and atherosclerotic lesions.²⁹ Treatment with 0.5 mmol/L ONOO^- resulted in up to a 55% decrease in ND2 and cyt b transcripts in HASMCs, and a 45% to 50% reduction in ND2 and cyt b transcripts in HUVECs (Figure 3). Treatment with 0.1 mmol/L ONOO^- reduced ND2 and cyt b transcript levels by 25% to 30% in HUVECs and 5% to 15% in HASMCs (Figure 3).

View larger version (36K): [in this window] [in a new window]   Figure 3. mtRNA transcript levels in ONOO--treated cells. Data are mean relative transcript levels (±SEM) normalized to ß-actin for 16S rRNA, ND2, and cyt b after exposure to 0.1 or 0.5 mmol/L ONOO- for 1 hour, compared with untreated cells. Transcript levels were determined by Northern analysis and hybridization to the appropriate radiolabeled probe. *Transcript levels significantly different (P<0.05) from controls.

RS Alter Mitochondrial Protein Synthesis
Having demonstrated that ONOO^- exposure decreased mtRNA levels, the influence of ONOO^- on mitochondrial protein synthesis was assessed by measuring [³⁵S]methionine incorporation in the presence of emetine, an inhibitor of nuclear, but not mitochondrial, protein synthesis. Exposure of cells to ONOO^- (0.5 mmol/L) resulted in a substantial decrease in mitochondrial protein synthesis (Figure 4). Lower doses of ONOO^- (0.1 mmol/L) produced a 12% decrease in [³⁵S]methionine incorporation in HASMCs (not performed in HUVECs). Treatment with H₂O₂ (0.2 mmol/L) resulted in a 23% and 33% decrease in overall mitochondrial protein synthesis in HUVECs and HASMCs, respectively.

View larger version (40K): [in this window] [in a new window]   Figure 4. [35S]Methionine incorporation in mitochondrial synthesized proteins. A, Protein labeling after exposure to 0.1 or 0.5 mmol/L ONOO- in HASMCs (left). Right, Same gel stained with Coomassie blue (total protein). B, Relative percentages, compared with controls, of [35S]methionine incorporation in both HUVECs and HASMCs exposed to 0.2 mmol/L H2O2 and 0.1 or 0.5 mmol/L ONOO-. ND indicates no data.

RS Inhibit Mitochondrial Function
The influence of RS on mitochondrial redox potential and cellular ATP level was evaluated by measuring MTT reduction and cellular ATP content. Analysis of MTT reduction by succinate dehydrogenase (mitochondrial complex II) reflects net cellular respiratory and redox functions (Figure 5).³⁰ Exposure to 0.1 mmol/L ONOO^- did not significantly decrease MTT reduction or cellular ATP, whereas 0.5 mmol/L ONOO^- induced a significant decrease in MTT reduction and cellular ATP in each cell type (P0.05). Treated HUVECs had greater declines in cellular ATP and MTT reduction compared with HASMCs (ATP, P=0.002; MTT, P=0.006). To verify that the decreases in MTT reduction and ATP production were not due to cell death, cells were stained with trypan blue 1 hour after oxidant exposure, showing essentially no cell death (<5% trypan blue staining) at times of MTT reduction and ATP level assessment.

View larger version (19K): [in this window] [in a new window]   Figure 5. Inhibition of mitochondrial complex II reduction and cellular ATP content. HASMCs and HUVECs were exposed to 0.1 or 0.5 mmol/L ONOO- for 1 hour and analyzed for relative levels of MTT reduction (A) and cellular ATP content (B). *Levels significantly different (P<0.05) from controls. Values are expressed relative to untreated cells (100%).

To evaluate whether mitochondrial membrane potentials () were altered by RS treatments, R123 uptake was determined in cells 1 hour after RS treatment. R123 selectively accumulates in the mitochondria, and rate of uptake is an indication of . HUVECs treated with 0.2 mmol/L H₂O₂ or 0.5 mmol/L ONOO^- had significantly lower rates of R123 uptake (Figure 6). In contrast, no differences were observed in R123 uptake in HASMCs (Figure 6).

View larger version (30K): [in this window] [in a new window]   Figure 6. R123 uptake in HUVECs (A) and HASMCs (B). Cells were treated with 0.2 mmol/L H2O2 or 0.5 mmol/L ONOO- in serum-free medium for 1 hour, trypsinized, washed, and resuspended in conditioned medium containing 1 µmol/L R123. Fluorescence was assessed at 1 minute and 30 minutes after R123 addition. Increased fluorescence is expressed relative to the 1-minute time point. *Significantly different (P<0.05) level of fluorescence.

To assess secondary RS formation associated with the treatments, we determined aconitase activity in HUVECs 1 to 2 hours after 0.2 mmol/L H₂O₂ treatment. Aconitase is specifically inactivated by O₂^·- and ONOO^-28 and is unaffected by H₂O₂ (Figure 7), so that aconitase activity is inversely related to the generation of O₂^·- and ONOO^-. HUVECs treated with 0.2 mmol/L H₂O₂ had a 3.4-fold decrease in aconitase activity compared with controls (Figure 7), indicating an increased production of secondary O₂^·- and ONOO^- in the treated cells.

View larger version (22K): [in this window] [in a new window]   Figure 7. Aconitase activity in H2O2-treated HUVECs. A, Control (no RS, 1 hour; •) and treated (0.2 mmol/L H2O2, 1 hour; ) cells were washed in PBS and lysed in assay buffer. Aconitase activity was measured for 1 hour. Decreased activity indicates increased O2·- production. B, Purified aconitase (5 mU, , ; 2 mU, , •) treated with serum-free medium (•, ) or serum-free medium plus 0.2 mmol/L H2O2 (, ).

	Discussion

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The RS used in this study mediate mitochondrial damage and dysfunction in HUVECs and HASMCs in vitro by increasing mtDNA damage, decreasing mtRNA transcripts, inhibiting mitochondrial protein synthesis, reducing cellular ATP, and altering redox function. These data suggest that RS, in pathophysiologically relevant concentrations, mediate mitochondrial damage and dysfunction that affect vascular cell function and can contribute to chronic vascular diseases.

With the exception of exposure to ^·NO, the various RS used, at their selected rates of production and/or concentration, yielded similar extents of mtDNA damage. This is likely due to the often similar target molecule reactions of different RS (eg, O₂^·- + H₂O₂ versus ONOO^-) as well as to their broad and potent reactivities. This is of relevance in atherogenesis, as individual RS will be generated at different rates by vascular cells during disease progression. The concentrations of ONOO^- used within these studies were well within physiological ranges.²⁰ Administration of a bolus concentration of ONOO^- produces effects comparable with longer, sustained low-dose exposures³¹ ; the additions of ONOO^- in the present study were equivalent to exposure to 1 µmol/L ONOO^- for 3 to 13 minutes. The half-life of ONOO^- at pH 7.4 is 1.6 seconds; thus, remotely located secondary oxidative reactions likely contribute to the sustained effect of this relatively long-lived oxidant.

mtRNA transcripts declined subsequent to ONOO^- exposure in HASMCs and HUVECs. This reduction was transient; 2 hours after treatment, mtRNA levels were restored to the same level as controls, indicating that mtRNA transcript reduction was directly related to RS exposure and reversible cell injury. The observation that the mitochondrial 16S rRNA transcript is minimally affected by exposure to RS may be the result of its previously noted differential expression³² or transcript stability.³³

Consistent with alterations in mitochondrial function, protein synthesis, and redox potential (complex II), mitochondrial membrane potentials were reduced in RS-treated HUVECs. In contrast, R123 uptake was not significantly affected in HASMCs, potentially because of increased resistance to RS-mediated damage (due to potentially greater antioxidant capacity or repair processes), and/or because of unique cell characteristics. Studies in liver tissues have shown that whereas ONOO^- treatment inhibits electron transport, it also promotes Ca²⁺ release from rat liver mitochondria under certain conditions, with maintenance of ³⁴ ; it may be that HASMCs respond similarly to RS-mediated stress. Further studies of the antioxidant and repair capacities of HASMCs relative to HUVECs will be required to determine why HASMCs appear more resistant to RS-mediated damage. Nevertheless, taken together, these data indicate that oxidative injury can mediate a cascade of events in HUVECs and HASMCs that ultimately result in mitochondrial damage and function.

A consistent observation for HUVECs and HASMCs was that nDNA is less affected by RS than mtDNA. It is possible that the concentrations of exogenously added RS achieved in the nucleus were lower than in the mitochondria. However, it is more likely that differences in structure, protein content, and repair make the nDNA more resistant to oxidative damage. The mtDNA is sensitive to damage because it lacks both protective histone and nonhistone proteins and has a limited DNA repair capacity (relative to the nucleus). For example, the mtDNA accumulates significantly higher levels of the DNA oxidation product 8-hydroxydeoxyguanosine than does the nDNA.³⁵ In addition, the high lipid-to-DNA ratio in mitochondria makes them especially susceptible to lipophilic species, whereas the attachment of the mtDNA to the matrix side of the inner membrane also increases sensitivity to membrane disturbances and makes mitochondria a target for electrophiles generated within the membrane.³⁶ Although endogenous rates of mitochondrial and nuclear radical generation are similar,³⁷ the added capacity of mitochondria to generate ^·NO³⁸ or differences in antioxidant defense system activities may contribute to the sensitivity of these organelles. Because the amount of intact DNA present at any point in time reflects a balance between damage and repair, it is possible that the extensive and persistent mtDNA damage is due to lesser repair and/or increased generation of secondary RS due to mitochondrial injury.

Although our studies have focused on the effects of RS on mitochondrial function, mitochondria themselves serve as a major source of RS. Mitochondria consume more than 90% of the oxygen available to the cell; 1% to 3% is used in the production of O₂^·-.³⁹ In addition to O₂^·-, mitochondria produce ^·NO⁴⁰ and CO₂⁴¹ and, thus, are capable of generating both intramitochondrial reactive oxygen species and reactive nitrogen species (ie, ONOO^- and ONOOCO₂^-), which efficiently mediate secondary oxidation and nitration reactions.⁴² Several reports reveal that ONOO^- inhibits mitochondrial respiration and enhances rates of respiratory chain O₂^·- and H₂O₂,⁴³ implicating these species in chronic mitochondrial damage. Consistent with this notion, numerous studies have shown an age-dependent decrease in mitochondrial respiratory enzyme activities,⁴⁰ increased lipid peroxidation,⁴⁴ mtDNA mutation,^{45 46} and damage. Studies have also shown that oxidized LDL inhibits mitochondrial function⁴⁷ and induces production of mitochondrial antioxidants⁴⁸ and that mitochondrial RS generation is involved in LDL oxidation.¹⁴ Finally, RS are capable of inhibiting specific mitochondrial enzymes,^{49 50} affecting cellular antioxidant and energetic capacities. Consequently, chronic and age-related increases in RS production will cause mitochondrial damage and dysfunction that perpetuate a catastrophic cycle of cellular injury, further RS generation, mitochondrial decline, and cell death. We are currently testing this hypothesis utilizing in vivo models of atherosclerosis.

	Acknowledgments

 
This work was supported by NIH grants to M.S.R. (HL59652), S.W.B. (ES09318), C.P. (HL03658), and B.V.H. (ES07038, 2PO1AG1O514) and by grants from the John Sealy Foundation (to S.W.B.), the Clayton Foundation (to M.S.R. and B.V.H.), and the Burroughs Wellcome Fund (to B.V.H.).

Received January 31, 2000; accepted March 21, 2000.

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