Exercise training improves vascular mitochondrial function

The impact of exercise training on vascular mitochondrial respiratory function is unknown. Here we show that arteries from swim-trained mice exhibit increased mitochondrial respiratory capacity, redox balance, and nitric oxide bioavailability compared with sedentary animals. These data confirm and extend the evidence that chronic physical activity is vasculoprotective.

Abstract

Exercise training is recognized to improve cardiac and skeletal muscle mitochondrial respiratory capacity; however, the impact of chronic exercise on vascular mitochondrial respiratory function is unknown. We hypothesized that exercise training concomitantly increases both vascular mitochondrial respiratory capacity and vascular function. Arteries from both sedentary (SED) and swim-trained (EX, 5 wk) mice were compared in terms of mitochondrial respiratory function, mitochondrial content, markers of mitochondrial biogenesis, redox balance, nitric oxide (NO) signaling, and vessel function. Mitochondrial complex I and complex I + II state 3 respiration and the respiratory control ratio (complex I + II state 3 respiration/complex I state 2 respiration) were greater in vessels from EX relative to SED mice, despite similar levels of arterial citrate synthase activity and mitochondrial DNA content. Furthermore, compared with the SED mice, arteries from EX mice displayed elevated transcript levels of peroxisome proliferative activated receptor-γ coactivator-1α and the downstream targets cytochrome c oxidase subunit IV isoform 1, isocitrate dehydrogenase (Idh) 2, and Idh3a, increased manganese superoxide dismutase protein expression, increased endothelial NO synthase phosphorylation (Ser¹¹⁷⁷), and suppressed reactive oxygen species generation (all P < 0.05). Although there were no differences in EX and SED mice concerning endothelium-dependent and endothelium-independent vasorelaxation, phenylephrine-induced vasocontraction was blunted in vessels from EX compared with SED mice, and this effect was normalized by NOS inhibition. These training-induced increases in vascular mitochondrial respiratory capacity and evidence of improved redox balance, which may, at least in part, be attributable to elevated NO bioavailability, have the potential to protect against age- and disease-related challenges to arterial function.

NEW & NOTEWORTHY

The impact of exercise training on vascular mitochondrial respiratory function is unknown. Here we show that arteries from swim-trained mice exhibit increased mitochondrial respiratory capacity, redox balance, and nitric oxide bioavailability compared with sedentary animals. These data confirm and extend the evidence that chronic physical activity is vasculoprotective.

while mitochondria comprise ∼10% of adult human body weight, likely due to differing metabolic demands, mitochondrial density is tissue specific. For example, the heart has a high metabolic demand, with mitochondria making up ∼35% of cardiac tissue volume and being responsible for generating up to 90% of cardiac ATP requirements via β-oxidation (25, 37). In contrast, the vasculature has a relatively low metabolic demand, and mitochondria comprise only ∼2–5% of the tissue volume, and ATP generation results largely from glycolysis (50). Our group recently documented that, when normalized for these large differences in mitochondrial content, mitochondrial respiratory capacity in cardiac, skeletal, and smooth muscle is very similar (37). Of particular importance to the current study, we also reported that complex I + II state 3 respiration was similar in intact and denuded vessels, suggesting that mitochondria located within the vascular smooth muscle, rather than the endothelium, are responsible for the majority of vascular respiration (37).

Evidence is accumulating that, rather than contributing to cellular respiration, an important role of the relatively small volume of mitochondria within the endothelium might be to act as signaling organelles that respond to cues from the local environment (50). Nitric oxide (NO), generated by endothelial nitric oxide synthase (eNOS), is a locally produced cue with documented potential to influence mitochondrial respiration in a number of cell types (e.g., brown adipocytes, 3T3-L1, U-937, and HeLa) (34, 35). Miller et al. (33) reported that, when basal NO generation was disrupted genetically (i.e., eNOS^+/− and eNOS^−/− mice) or pharmacologically [i.e., nitric oxide synthase (NOS) inhibition in rats via N^G-nitro-l-arginine methyl ester (l-NAME)], the protein abundance of the electron transport chain (ETC) subunits was decreased in the aorta of these animals, relative to controls. These data support the potential role of basal NO generation in influencing mitochondrial capacity reported earlier in a variety of cell types (34, 35) and extend this important finding to include mitochondria located in the vasculature.

If mitochondrial respiration is suppressed by pharmacological and genetic approaches that limit basal NO production, but stimulated by NO donors (i.e., S-nitrosoacetyl penicillamine) (34), it is reasonable to hypothesize that a physiological intervention known to increase eNOS activity and NO generation (53), such as exercise training, might facilitate vascular mitochondrial respiration. Indeed, two investigations from the Reusch laboratory group (26, 33) reported that the content of one or more of the mitochondrial ETC subunits was greater in aortic lysates obtained from rats that performed treadmill running (8 days × 30–45 min/session) compared with their sedentary counterparts. Furthermore, the contribution from NOS was implicated by the finding that the adaptive mitochondrial response to exercise training was blunted in rats treated concurrently with l-NAME (33). While these insightful studies suggest that NO-mediated signaling is requisite for exercise to stimulate vascular mitochondrial adaptations, the impact of exercise training on mitochondrial respiratory capacity in the vasculature has not been directly assessed.

Consequently, the purpose of the present study was to test the hypothesis that exercise training concomitantly increases vascular mitochondrial respiratory capacity and vascular function. Arteries from both sedentary (SED) and exercise-trained (EX) male mice were compared in terms of mitochondrial respiratory function, vessel function, mitochondrial content, markers of mitochondrial biogenesis, redox balance, and NO signaling. Our results indicate that exercise training improves vascular mitochondrial respiratory capacity and redox balance, and evidence is provided that this adaptation is at least partly attributable to elevated NO bioavailability.

MATERIALS AND METHODS

Animals.

Male C57BL/6J/129Sv mice were housed under controlled temperature (22°C) and light conditions (12:12-h light-dark cycle) and were provided with food and water ad libitum. All experiments were performed in accordance with protocols approved by the Institutional Animal Care and Use Committee of the University of Utah.

Exercise training and tissue collection.

The 6-wk-old EX mice completed two swim-training sessions per day, each separated by at least a 4-h interval, for 36 days. The SED mice, of the same age, were also transported to the swim location two times per day, but remained in their cages. The duration of the initial session (10 min) was increased by 10 min each day until 90 min of continuous swimming was reached and maintained for an additional 28 days (24, 40). Mice were killed, and the entire aorta, both iliac arteries, and both femoral arteries were dissected free from adherent tissue while immersed in ice-cold dissection buffer that contained protease and phosphatase inhibitors (4, 45, 52). Vessel segments were used for the experiments described below.

Measurement of vascular mitochondrial respiration: aorta.

Immediately upon excision the entire aorta was placed in ice-cold buffer A containing (in mM) 2.77 CaK₂EGTA, 7.23 K₂EGTA, 6.56 MgCl₂, 0.5 dithiothreitol (DTT), 50 K-MES, 20 imidazole, 20 taurine, 5.77 Na₂ATP, and 15 phosphocreatine at pH 7.1 for 30 min. Next, being shaken mildly for 40 min in buffer A supplemented with 50 μg/ml saponin, the aorta was rinsed (2 × 10 min/rinse) in buffer B containing (in mM) 2.77 CaK₂EGTA, 7.23 K₂EGTA, 6.56 MgCl₂, 0.5 DTT, 50 K-MES, 20 imidazole, 20 taurine, 5.77 ATP, and 15 phosphocreatine at pH 7.0 (37). Mitochondrial respiration was assessed by measuring the oxygen consumption rate in buffer B, while being continuously stirred, at 37°C using a Clark electrode (Hansatech, Kings Lynn, UK), as previously described (37). Briefly, after the baseline respiration rate in the absence of substrate was recorded: 1) complex I state 2 respiration was assessed in the presence of glutamate + malate; 2) complex I state 3 respiration, the ADP-stimulated state of oxidative phosphorylation, was measured in the presence of glutamate + malate + ADP; and 3) complex I + II state 3 respiration was evaluated in the presence of glutamate + malate + ADP + succinate (37). In all experiments, the integrity of the outer mitochondrial membrane was confirmed by cytochrome c injection after the assessment of complex I and II state 3 respiration. None of the samples exhibited an increase in the rate of oxygen consumption following the addition of cytochrome c (data not shown). After respiration measurements, vessels were snap-frozen, and mitochondrial DNA content (mtDNA) and citrate synthase activity (CSA) were determined (29).

The respiratory control ratio (RCR) is defined as the ADP-stimulated flux divided by the flux without phosphorylation of ADP and was calculated as complex I + II state 3/complex I state 2 respiration. It should be noted that state 2 respiration was determined in the presence of glutamate + malate (in the absence of the complex II substrate succinate, since complex II does not release protons to the intermembrane space). Importantly, no difference was observed for state 2 respiration when comparing glutamate + malate + succinate vs. glutamate + malate as substrates (data not shown). For measurement of state 3 respiration, ADP together with succinate was supplemented to the respiration buffer to prevent depletion of metabolites from the mitochondrial matrix and to reconstitute the tricarboxylic acid cycle (14, 15, 37). Concentrations of each reagent in the vessel chamber were glutamate (2 mM), malate (10 mM), ADP (5 mM), succinate (10 mM), and cytochrome c (10 μM) (37).

Quantitative RT-polymerase chain reaction: Aorta.

Total RNA was extracted from aortic tissue with TRIzol reagent (Invitrogen, Carlsbad, CA) and was reverse transcribed (SuperScript III Reverse Transcriptase Kit; Invitrogen). Platinum Taq DNA polymerase (Invitrogen), primers, SYBR Green fluorescent dye (Invitrogen), and cDNA were transferred to a 384-well plate in triplicate, and real-time PCR was performed with an ABI Prism 7900HT instrument (Applied Biosystems, Foster City, CA) as previously described (39, 40). Data were normalized to ribosomal protein S16 (Rps16), and the results were expressed relative to the SED control mice. The following primers were used: cytochrome c oxidase subunit IV isoform 1 (Cox4i1): forward 5′-CGCTGAAGGAGAAGGAGAAG-3′, reverse 5′-GCAGTGAAGCCAATGAAGAA-3′; isocitrate dehydrogenase (Idh) 2: forward 5′-CCCTATTGCCAGCATCTTTG-3′, reverse 5′-TGTCCAGGAAGTCTGTGGTG-3′; Idh3a: forward 5′-CCCATCCCAGTTTGATGTTC-3′, reverse 5′-GCATCATCACAGCACTAAGCA-3′; peroxisome proliferative activated receptor-γ coactivator-1α peroxisome proliferative activated receptor-γ coactivator-1α (Ppargc1a) forward 5′-GTAAATCTGCGGGATGATGG-3′, reverse 5′-AGCAGGGTCAAAATCGTCTG-3′; and Rps16: forward 5′-TGCTGGTGTGGATATTCGGG-3′, reverse 5′-CCTTGAGATGGGCTTATCGG-3′.

Measurement of mtDNA content: Aorta.

DNA was extracted from aortic segments using a phenol-chloroform-isoamyl alcohol solution (25:24:1) followed by ethanol precipitation. Next, DNA was dissolved in 100 μl of Tris-EDTA, and 5 μl of a 50 times dilution was subjected to real-time PCR amplification using the QuantiTect SYBR Green PCR Kit (Qiagen, Valencia, CA) in a total reaction volume of 25 μl containing 0.5 μM of each primer. Real-time PCR was performed with an iQ real-time PCR detection system (Bio-Rad, Hercules, CA). The mtDNA content relative to nuclear DNA content was determined using the following primers: mtDNA-encoded cytochrome c oxidase 1: forward 5′-ACTATACTACTACTAACAGACCG-3′, reverse 5′-GGTTCTTTTTTTCCGGAGTA-3′; and nuclear DNA-encoded cyclophilin A: forward 5′-ACACGCCATAATGGCACTGG-3′, reverse 5′-CAGTCTTGGCAGTGCAGAT-3′ (5). Data were normalized relative to SED controls.

Measurement of CSA: Aorta and gastrocnemius.

Frozen vessels previously used for mitochondrial respiration measurements were homogenized (in mM: 250 sucrose, 40 KCl, 2 EGTA, and 20 Tris·HCl, pH 7.4). The homogenates were then supplemented with 0.1% Triton X-100 and incubated on ice for 60 min followed by centrifugation for 8 min at 10,000 g and a 20 times dilution (37). Similarly, gastrocnemius muscle was homogenized followed by two freeze-thaw cycles to release the citrate synthase from the mitochondrial matrix, followed by centrifugation for 10 min and a 10× dilution (40). CSA was determined in a total reaction volume of 200 μl for vessel homogenates and 1 ml for skeletal muscle homogenates. The reaction was performed in reaction buffer containing (in mM) 220 sucrose, 40 KCl, 20 HEPES, 1 EGTA, 0.1 5,5′-dithio-bis-2-nitrobenzoic acid (DTNB), and 0.1 acetyl-CoA, pH 7.4 at 25°C, and was started by the addition of 0.05 mM oxaloacetate. CSA was monitored at 412 nm to detect the reaction of sulfhydryl groups of CoA with DTNB for a total duration of 3 min using an Ultrospec 3000 spectrophotometer (Amersham Pharmacia Biotech).

Immunoblotting: Iliac and femoral arteries.

Protein isolation and immunoblotting analyses were performed using both iliac arteries and segments of femoral arteries not used for vascular function experiments using procedures that have been previously described (4, 45, 52). With the exception of manganese superoxide dismutase (MnSOD; Enzo Life Sciences, Farmingdale, NY), all primary antibodies were obtained from Cell Signaling (Danvers, MA). Alexa Fluor anti-Mouse 680 or anti-Rabbit 680 (Invitrogen) were used as secondary antibodies, and fluorescence was quantified using the LICOR Odyssey imager (Lincoln, NE).

Measurement of reactive oxygen species: Iliac and femoral arteries.

Reactive oxygen species (ROS) were quantified by 2′,7′-dichlorofluorescein diacetate (DCFDA) fluorescence. The assay is based on the conversion of nonfluorescent DCFDA to the highly fluorescent DCF in the presence of ROS. Tissue samples were homogenized in 0.05% Trypsin-EDTA (Invitrogen) and incubated for 30 min at 37°C. Following centrifugation (5 min at 14,000 g), pellets were incubated in DCFDA dissolved in DMEM (Invitrogen) at a final concentration of 5 μM for 30 min at 37°C. Samples were centrifuged for 5 min at 14,000 g, and pellets were resuspended in 200 μl of lysis buffer (50 mM HEPES, 150 mM NaCl, 10% glycerol, 1% Triton X-100, 1.5 mM MgCl₂, 1 mM EGTA, 10 mM sodium pyrophosphate, 100 mM sodium fluoride, 100 μM sodium vanadate, 1 mM phenylmethylsulfonyl fluoride, 10 μg/ml aprotinin, and 10 μg/ml leupeptin) and incubated for 10 min at 4°C under constant agitation. Next, samples were centrifuged (5 min at 14,000 g), and supernatants were transferred to a black 96-well plate. DCF was determined by spectrophotometry using a fluorescence plate reader (Synergy HT Multi-Detection Reader; Bio-Tek Instruments, Winooski, VT) at 485 nm excitation/530 nm emission. Data were normalized to protein content, and results were expressed as a fold change relative to SED controls.

Measurement of vascular function: Femoral arteries.

Vascular function was assessed in two femoral artery segments from each mouse using isometric tension techniques (4, 45, 52). First, both vessels completed a series of internal circumference-active tension curves to determine the vessel diameter that evoked the greatest tension development (L_max) to 100 mM KCl. Second, non-receptor-mediated and receptor-mediated vasocontractile responses to KCl (20–100 mM) and phenylephrine (PE, 10⁻⁸ to 10⁻⁵ M), respectively, were assessed. Third, after arteries were precontracted to ∼65% of maximal PE-induced contraction and tension was stable (1, 4, 9, 45, 52), responses to acetylcholine (ACh, 10⁻⁸ to 10⁻⁶ M) were assessed to determine stimulated endothelium-dependent vasorelaxation. In a fourth protocol, one vessel from each mouse completed a second PE dose-response curve after a 10-min wash and a 20-min incubation with N^G-monomethyl-l-arginine (l-NMMA, 10⁻³ M). Results were compared with the same vessel that was treated initially with PE in the absence of l-NMMA. Separate experiments revealed that repeatable contractions to PE are observed in femoral arteries when concentration-response curves are separated by 30 min. Specifically, maximal PE-evoked increases in tension development from baseline were not different when two dose-response curves were separated by 30 min (0.829 ± 0.058 vs. 0.911 ± 0.062 mg developed tension/μm vessel length). A fourth protocol completed on the second vessel obtained from each mouse involved administering 10⁻³ M l-NMMA to arteries that were stably precontracted with PE. l-NMMA-evoked vasocontraction in this context is a well-accepted functional estimate of basal eNOS activity (31) that we have used previously (45). The fifth and final protocol for both vessels involved administering sodium nitroprusside (10⁻⁹ to 10⁻⁴ M) to PE-precontracted arteries to determine endothelium-independent vasorelaxation. At least 30 min separated each protocol. When two vessel segments were treated identically, the results were averaged. All tension data were recorded continuously using an analog-to-digital interface card (Biopac Systems, Santa Barbara, CA) that allowed for subsequent off-line quantitative analyses.

Statistical analysis.

Data are presented as means ± SE. Significance was accepted when P < 0.05. The comparison of one time point between two groups was made using an unpaired t-test. The comparison of multiple time points between groups was made using a one-way or two-way repeated-measures analysis of variance. Tukey's post hoc tests were performed when significant main effects were identified.

RESULTS

Efficacy of the exercise training regimen.

CSA (nmol·min⁻¹· μg protein⁻¹) was greater in the gastrocnemius muscle obtained from the EX (44 ± 3) compared with the SED (32 ± 4) mice (Fig. 1A).

Fig. 1.

Vascular mitochondrial respiratory capacity increases in response to exercise training. A: citrate synthase activity (CSA) in gastrocnemius muscle (n = 6). B: mitochondrial oxygen consumption in permeabilized aortas from exercise-trained (EX) and sedentary (SED) mice normalized to vessel wet weight. C I State 2, complex I state 2 respiration; C I State 3, complex I state 3 (following ADP stimulation); and C I + II State 3, complex I and II state 3 respiration (following ADP stimulation). C: respiratory control ratio (RCR), complex I + II state 3 normalized to complex I state 2 (n = 8 animals, 16 vessels). D and E: mRNA expression of Ppargc1a (D) and genes involved in mitochondrial bioenergetics (E) following EX (n = 8). Data are presented as a fold change relative to SED, n = 8. F and G: mitochondrial DNA (mtDNA) copy number (F) and CSA (G) in aortas (n = 8). *P < 0.05 vs. SED; ns, Not significant.

Vascular mitochondrial respiratory capacity in the EX and SED mice.

Mitochondrial complex I, state 2 respiration was similar in the EX and SED mice (Fig. 1B). In contrast, mitochondrial complex I (11.9 ± 0.8 vs. 8.6 ± 1.2 pmol·mg⁻¹·s⁻¹) and complex I + II (20.8 ± 0.9 vs. 13.2 ± 1.3 pmol·mg⁻¹·s⁻¹; Fig. 1B), state 3 respiration, and RCR (2.5 ± 0.1 vs. 1.6 ± 0.1; Fig. 1C), were greater in vessels from EX compared with SED mice, respectively. Transcript levels of peroxisome proliferative activated receptor-γ coactivator-1α (PGC-1α; Ppargc1a) and the PGC-1α downstream targets involved in mitochondrial bioenergetics (Cox4i1, Idh2, Idh3a) were 1.3- to 1.5-fold greater in vessels from the EX compared with the SED mice (Fig. 1, D and E). Vascular mtDNA content (Fig. 1F) and CSA (Fig. 1G) were not different between groups.

Vascular redox balance in the EX and SED mice.

MnSOD protein expression was approximately threefold greater (Fig. 2, A and B) and DCFDA fluorescence was 26% lower (Fig. 2C) in the vessels from the EX compared with the SED mice.

Fig. 2.

Vascular indexes of antioxidant capacity following exercise training. A and B: manganese superoxide dismutase (MnSOD) protein expression normalized to GAPDH. C: reactive oxygen species (ROS) estimated by 2′,7′-dichlorofluorescein diacetate (DCFDA) fluorescence in iliac/femoral artery homogenates (n = 8). *P < 0.05 vs. SED.

Vascular function and kinase signaling to eNOS in the EX and SED mice.

There was no difference in vessel width (197 ± 6 and 221 ± 9 μm), vessel length (1,842 ± 23 and 1,846 ± 18 μm), resting tension at L_max (699 ± 10 and 713 ± 16 mg), or percent precontraction (68 ± 3 and 69 ± 4%), before the assessment of endothelium-dependent vasorelaxation, between the EX and SED mice, respectively. Neither endothelium-dependent (Fig. 3A) nor endothelium-independent (Fig. 3B) vasorelaxation was different when responses were compared among vessels from SED and EX mice. In contrast, non-receptor-mediated vasocontraction (Fig. 3C) tended to be greater in vessels from SED vs. EX mice at 80 mM (P = 0.09) and 100 mM (P = 0.06) KCl (Fig. 3C), and receptor-mediated vasocontraction in response to PE was greater in arteries from the SED vs. EX mice (P = 0.027; Fig. 3D). PE-induced vasocontraction in both groups was greater (P < 0.05) in the presence vs. the absence of NOS inhibition using l-NMMA. However, no difference was observed between groups concerning PE-induced vasocontraction when results were compared in the presence of NOS inhibition (P = 0.390; Fig. 3D). Furthermore, l-NMMA caused a 16 ± 4% vs. 8 ± 2% increase (P = 0.06) in tension development in vessels from EX vs. SED mice, respectively.

Fig. 3.

Vascular function and kinase-mediated signaling to endothelial nitric oxide synthase (eNOS) following exercise training. A and B: endothelium-dependent (EDR, A) and independent (EIR, B) vasorelaxation. C and D: non-receptor-mediated (NRC, C) and receptor-mediated (RC, D) vasocontraction in the absence (n = 6–7 animals each, 19–23 vessels each) or presence of nitric oxide synthase inhibition using N^G-monomethyl-l-arginine (l-NMMA) (n = 7 animals, 9 vessels). E and F: representative Western blots from iliac/femoral artery homogenates (E) and densitometric quantification as indicated (F) (n = 7–8). Data are presented as fold change relative to SED: *P < 0.05 vs. SED.

In terms of kinase signaling to eNOS, arteries from the EX mice exhibited greater vascular phospho (p)-eNOS^Ser1177/total eNOS, which was paralleled by greater phosphorylation of the upstream kinase p-Akt^Thr308/total Akt and p-AMPK^Thr172/total AMPK (Fig. 3, E and F).

DISCUSSION

The primary purpose of the present study was to determine the impact of exercise training on vascular mitochondrial respiratory function. However, afforded by a combination of vessel function studies, the assessment of markers of mitochondrial biogenesis, redox balance, and NO signaling, we provide insight into the potential mechanisms and consequences of exercise training-induced mitochondrial and vascular adaptations. Specifically, despite evidence of unaltered mitochondrial content, vascular mitochondrial respiratory capacity was significantly greater in the EX mice, and this was accompanied by a greater RCR. In terms of vascular function, although the EX mice did not exhibit enhanced ACh-induced endothelial-mediated relaxation, evidence of greater NO bioavailability in arteries from EX vs. SED mice is provided. Finally, vessels from the EX mice exhibited greater MnSOD protein expression and suppressed ROS generation, which can be interpreted as evidence that exercise-induced oxidant stress was balanced by the appropriate antioxidant defense. These exercise training-induced increases in vascular mitochondrial respiratory capacity and evidence of improved redox balance, which were associated with elevated basal NO bioavailability, have the potential to protect against age- and disease-related challenges to arterial function.

Vascular mitochondrial respiratory capacity and exercise training.

Important roles for the mitochondria, e.g., oxygen sensing, calcium handling, and ROS production, are emerging and underscore the importance of mitochondrial quality control in health and disease. Prior studies have documented the exercise training-induced increase in mitochondrial respiratory capacity in both skeletal and cardiac muscle (23, 40). However, with only a few exceptions (19), the preponderance of data suggests that this exercise-induced increase in mitochondrial respiratory capacity is simply the consequence of an increase in mitochondrial content and not an inherent change in function of the mitochondrion (8, 16, 18, 21). Of note, the current CSA data from the gastrocnemius of the SED and EX mice, with a higher value in the EX mice, both support the concept of an exercise-induced increase in mitochondrial content in skeletal muscle and document the efficacy of the training intervention (Fig. 1A). The underlying mechanisms responsible for these exercise-induced mitochondrial adaptations in cardiac and skeletal muscle are incompletely understood, although it is becoming increasingly recognized that both PGC-1α and NO/cGMP-dependent pathways likely play a critical role (34, 35, 40). Similarly, aortic tissue from Sprague-Dawley rats (26) revealed a greater protein abundance of the ETC subunits as a consequence of 8 days of treadmill exercise. The PGC-1α-mediated transcriptional response to exercise training observed in the arteries of the EX mice in the current study provides a potential mechanism for the increased mitochondrial respiratory capacity following training (Fig. 1, B–E). However, in this study there was no evidence that the relatively mild increase in Ppargc1a resulted in mitochondrial biogenesis, as assessed by CSA and mtDNA content, but it is possible that this upregulation might have been sufficient to increase the mRNA expression of ETC subunits and mitochondrial respiratory capacity, independent of mitochondrial content. Importantly, the current study is the first to document greater vascular mitochondrial respiratory capacity in response to exercise training using a functional ex vivo assay.

Vascular function, mitochondrial respiratory capacity, and exercise training.

Originally we hypothesized that endothelium-dependent vasorelaxation would be greater in arteries from EX vs. SED mice. Instead, ACh-evoked vasorelaxation was similar between groups after the arteries were precontracted to the same extent using PE. While ACh-evoked vasorelaxation represents stimulated eNOS enzyme activity, it should be noted that several indexes of increased basal eNOS enzyme activity were observed in arteries from EX vs. SED mice. First, p-eNOS^Ser1177, a positive regulatory residue on the eNOS enzyme, was greater in arteries from EX vs. SED mice (Fig. 3, E and F). Second, exaggerated PE-induced vasocontraction observed in arteries from SED vs. EX animals was negated when dose-response curves were repeated in both groups in the presence of eNOS inhibition using l-NMMA (Fig. 3D). Third, l-NMMA-evoked vasocontraction of submaximally contracted arteries, a well-accepted functional estimate of basal eNOS activity (31), tended to be greater (P = 0.06) in vessels from EX vs. SED mice. Specifically, l-NMMA administration to PE-precontracted arteries evoked a 16 ± 4% vs. 8 ± 2% increase in tension development in vessels from EX vs. SED mice, respectively. In combination, these findings support the statement that endogenous opposition to vasocontraction afforded by basal NO was greater in arteries from EX vs. SED mice.

Interestingly, mitochondrial respiration is suppressed by pharmacological and genetic procedures that limit basal NO production, but is stimulated by NO donors in brown adipocytes, 3T3-L1, U-937, and HeLa cells. Furthermore, because 8 days of treadmill running increases mitochondrial ETC subunits in aortic lysates in a manner that is sensitive to NOS inhibition, we reasoned that a training-induced increase in basal NO generation might facilitate vascular mitochondrial respiration. Results from the present study are the first to reveal that training-induced increases in basal arterial NO production occur concurrently with elevations in vascular respiratory capacity. The mechanism(s) whereby vascular NO production might influence vascular respiratory capacity is complex and incompletely understood. For example, in addition to reports that basal NO production increases the expression of ETC subunits (26, 33), studies indicate that elevated NO concentrations inhibit cytochrome c oxidase activity to thereby impair mitochondrial respiratory capacity (6, 7). Also, it has been shown that mitochondrial depolarization activates NO synthase in endothelial cells to an extent that might link metabolic demand of exercise to vasodilation (22). The apparent paradoxical regulation of mitochondrial respiratory capacity by NO might be secondary to the intensity and/or nature of the stimulus. For example, mitochondrial respiration can be inhibited by high, but not low, concentrations of NO, and the effect of NO can be inhibited by light and thiols (6). Training studies that use different intensities and durations coupled with procedures to limit or exaggerate vascular NO production will be necessary to rigorously investigate this important issue.

Vascular mitochondrial function, redox balance, and exercise training.

Mitochondrial dysfunction has been identified as a risk factor for vascular dysfunction and cardiovascular disease as a consequence of greater ROS production (28, 32, 47). Indeed, it has been suggested that attenuated mitochondrial content, in both vascular endothelial and smooth muscle cells, and the resultant increase in mitochondrial-derived ROS production may contribute to vascular dysfunction in aged rats (46). In the present study, we observed increased Ppargc1a and MnSOD expression and a decreased abundance of ROS following exercise training (Fig. 2, A–C), which, by either cause or effect, likely improve vascular redox balance and mitochondrial function. PGC-1α has been documented to regulate the expression of ROS-detoxifying enzymes, including MnSOD (43). Overexpression of PGC-1α in endothelial cells induced the expression of an antioxidant transcriptional program, resulting in increased mitochondrial detoxification capacity and decreased ROS levels. This was paralleled by increased mitochondrial membrane potential and decreased apoptotic cell death (42, 48). Furthermore, it has been determined that ROS formation plays a critical role for the positive effects of exercise training, including the induction of a PGC-1α-mediated transcriptional program and an adaptive endogenous antioxidant response. This has been identified in human muscle samples as evidenced by an attenuated increase in PGC-1α and MnSOD expression following antioxidant supplementation with exercise training (41). While one study reported increased superoxide concentrations following exercise training (2), it is important to note that these data were obtained following a short-term exercise training protocol, and no chronic training effect was reported. The present study used a training regimen of 36 days total, which might account for the differences in the ROS levels observed when comparing these studies. In this context, the induction of the antioxidant response might compensate for the acute increase in ROS resulting in an overall improved redox balance. The present study uses two independent measurements indicating increased antioxidant defense i.e., increased expression of MnSOD and decreased cellular oxidative stress. While these assessments were obtained at the end of the study, it would be of great interest to investigate the redox balance and antioxidant stress response at different time points throughout the training regimen, and this work is ongoing.

Acute exercise has been recognized to increase the phosphorylation and activation of protein kinase B (Akt), AMP-activated protein kinase (AMPK), and eNOS in the vasculature (53). Similar effects were observed in the present study following chronic exercise training, as also previously reported for cardiac tissue (51). In addition to the induction of an antioxidant response following acute training, increased ROS mediate the activation of Akt and AMPK signaling (2). We recently showed that AMPK is sufficient to activate p-eNOS^Ser1177 in the presence of the phosphatidylinositol 3-kinase inhibitor wortmannin following acute training (51). Of note, in addition to Akt and AMPK, changes in blood flow activate eNOS, as observed in mesenteric arteries obtained from rats. These mechanisms are both Ca²⁺ dependent (eNOS translocation from the plasma membrane to intracellular compartments) and Ca²⁺ independent (via Akt-mediated eNOS phosphorylation) (10).

Experimental considerations.

First, the current study was performed using healthy young mice, and it would be of great interest to ultimately investigate the impact of chronic exercise training on mitochondrial/vascular function and redox balance in the context of aging and/or disease. Second, our studies did not specifically address whether the mitochondrial and transcriptional changes we observed represent alterations within the vascular smooth muscle and/or endothelial cell compartment. Third, different segments of the vascular tree were used to investigate the endpoints reported. Because gene expression is differentially regulated throughout the vasculature (20, 36), we cannot exclude the possibility that exercise training might affect blood vessels in a heterogeneous manner, depending on their anatomical location. We acknowledge this limitation but highlight that comparisons between EX and SED mice in this study were performed in the same segment of the arterial tree. Finally, to definitively prove that NO increases vascular mitochondrial respiratory capacity, training studies are required using loss-of-function (e.g., eNOS null mice) and gain-of-function [e.g., eNOS transcription enhancers (12)] approaches. Ongoing studies in our laboratory are addressing each of these issues.

In conclusion, this study indicates that exercise training increases vascular mitochondrial respiratory capacity and improves vascular redox balance, and that these modifications correlate with elevated arterial NO bioavailability. These adaptations are relevant clinically because they have potential to protect against age- and disease-related challenges to arterial function, and our overall findings provide evidence that exercise training/physical activity is important to maintaining or improving vascular health.

GRANTS

Undergraduate student support was provided by the American Physiological Society (APS) Undergraduate Summer Research Fellowship Program, the APS Undergraduate Research Excellence Fellowship Program, the APS STEP-UP Program, the American Heart Association, Western States Affiliate, Undergraduate Student Summer Research Program, the American Diabetes Association Minority Undergraduate Internship Program, the Science without Borders Program of the Government of Brazil, the Native American Research Internship Program, and the University of Utah Undergraduate Research Opportunities Program. R. S. Richardson was supported, in part, by the National Institutes of Health (NIH) (PO1-HL-091830) and the United States Department of Veterans Affairs Rehabilitation Research and Development Service (E6910-R, E1697-R, E1433-P, and E9275-L). E. D. Abel was supported, in part, by NIH Grant RO1-DK-092065. J. D. Symons was supported, in part, by the American Diabetes Association (1-12-BS-208, ADA 7-08-RA-164), the NIH (2R15-HL-091493), and Seed Grants from the University of Utah Office of the Vice President for Research, the University of Utah College of Health, the University of Utah Center on Aging, and the University of Utah Diabetes and Metabolism Center. C. Riehle was supported by a Postdoctoral Fellowship from the German Research Foundation.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

S.-Y.P., J.D.S., and C.R. conception and design of research; S.-Y.P., M.J.R., J.R.G., L.P.B., J.D.S., and C.R. performed experiments; S.-Y.P., L.P.B., J.D.S., and C.R. analyzed data; S.-Y.P., J.D.S., and C.R. interpreted results of experiments; S.-Y.P., J.D.S., and C.R. prepared figures; S.-Y.P., J.D.S., and C.R. drafted manuscript; S.-Y.P., J.B., R.S.R., E.D.A., J.D.S., and C.R. edited and revised manuscript; S.-Y.P., M.J.R., J.R.G., L.P.B., J.B., R.S.R., E.D.A., J.D.S., and C.R. approved final version of manuscript.