We hypothesized the coordinate induction of mitochondrial regulatory genes in the hypertrophied right ventricle to sustain mitochondrial respiratory capacity and contractile function in response to increased load. Wistar rats were exposed to hypobaric hypoxia (11% O2) or normoxia for 2 wk. Cardiac contractile and mitochondrial respiratory function were separately assessed for the right and left ventricles. Transcript levels of several mitochondrial regulators were measured. A robust hypertrophic response was observed in the right (but not left) ventricle in response to hypobaric hypoxia. Mitochondrial O2 consumption was increased in the right ventricle, while proton leak was reduced vs. normoxic controls. Citrate synthase activity and mitochondrial DNA content were significantly increased in the hypertrophied right ventricle, suggesting higher mitochondrial number. Transcript levels of nuclear respiratory factor-1, peroxisome proliferator-activated receptor-γ-coactivator-1α, cytochrome oxidase (COX) subunit II, and uncoupling protein-2 (UCP2) were coordinately induced in the hypertrophied right ventricle following hypoxia. UCP3 transcript levels were significantly reduced in the hypertrophied right ventricle vs. normoxic controls. Exposure to chronic hypobaric hypoxia had no significant effects on left ventricular mitochondrial respiration or contractile function. However, COXIV and UCP2 gene expression were increased in the left ventricle in response to chronic hypobaric hypoxia. In summary, we found coordinate induction of several genes regulating mitochondrial function and higher mitochondrial number in a model of physiological right ventricular hypertrophy, linking the efficiency of mitochondrial oxidative phosphorylation and respiratory function to sustained contractile function in response to the increased load.
- gene expression
- hypobaric hypoxia
it is well described that chronic hypobaric hypoxia results in pulmonary vasoconstriction and an increase in pulmonary artery pressure, leading to the development of right ventricular (RV) hypertrophy. This trophic response is considered an adaptive mechanism to sustain RV cardiac output in response to increased load. For example, we recently reported a robust hypertrophic response in the RVs of rats exposed to chronic hypobaric hypoxia (1, 22). Moreover, gene expression of atrial natriuretic factor (ANF), a marker of cardiac hypertrophy, was increased in parallel. In addition, we also found that increased RV weight was not associated with a greater degree of fibrosis after 2 wk of hypobaric hypoxia, indicating a model of physiological RV hypertrophy (22).
Previous studies suggest that altered mitochondrial respiratory capacity may play a key role to sustain the contractile function of the hypertrophied heart. For example, in response to chronic hypobaric hypoxia, maintained mitochondrial respiration was found in the hypertrophied RV but not left ventricle (LV) (20). Furthermore, Nishio et al. (17) reported increased mitochondrial content in response to LV hypertrophy, while others found mitochondrial functional alterations in experimental models of volume overload- and pressure overload-induced cardiac hypertrophy (14). An emerging paradigm therefore suggests that, during the onset of moderate physiological cardiac hypertrophy, multiple adaptive pathways are triggered to sustain mitochondrial respiratory function, thereby allowing the hypertrophied heart to meet higher energetic demands in response to increased load. However, the functional significance and molecular mechanisms underlying such changes are poorly understood.
In this study, we hypothesized that exposure to chronic hypobaric hypoxia coordinately upregulates mitochondrial regulatory genes and mitochondrial content in the hypertrophied RV, as part of an adaptive response to sustain mitochondrial respiratory capacity and contractile function. To test our hypothesis, we exposed rats to 2 wk of hypobaric hypoxia (11% O2) and determined cardiac contractile and mitochondrial respiratory function for the RV and LV, respectively. Moreover, we performed real-time quantitative RT-PCR analysis to measure transcript levels of several mitochondrial regulators. Our data demonstrate the coordinate induction of several genes regulating mitochondrial respiratory function and increased mitochondrial DNA (mtDNA) content in the hypertrophied RV, linking the efficiency of mitochondrial oxidative phosphorylation and respiratory function to sustained RV contractile function in response to the increased load.
Six-week old male Wistar rats (weighing 190–230 g) were initially housed at room temperature on a 12:12-h reverse light-dark cycle [lights off at 4 AM zeitgeber time (ZT12); lights on at 4 PM (ZT0)], with access to a conventional laboratory diet and water ad libitum. The hypoxic groups were housed inside a lexan chamber (45 kPa, 11% O2) (SciTech, Cape Town, South Africa) for 14 days as described previously (22) and compared with age-matched normoxic controls. On day 14, animals were anesthetized with pentobarbital sodium (100 mg/kg ip) whereupon the hearts were isolated and perfused for functional assessment. For mitochondrial respiration, cardiac morphometrics, and gene expression determination, the RV was carefully separated from the LV plus the interventricular septum (LV+S). All rats were killed between ZT16 and ZT18, since they are most metabolically active during this period (33). The University of Cape Town's Animal Research Ethics Committee approved all animal experiments, and the investigation conformed to the National Research Council's Guide for the Care and Use of Laboratory Animals (National Institutes of Health publication no. 85-23, revised 1996).
Cardiac mitochondrial isolation and functional characterization.
Mitochondria were isolated according to the method of Sordahl et al. (24) with modifications. Briefly, RV and LV tissues were homogenized separately in 10 ml of ice-cold potassium-EDTA (KE) buffer (0.18 M KCl, 10 mM EDTA, pH 7.4); then, the homogenate was centrifuged at 755 g for 5 min. The supernatant was subsequently filtered through 41-μm nylon mesh (Spectrum), and the filtrate was centrifuged at 1,480 g for 5 min. The mitochondrial pellet was resuspended in 50 μl of KE buffer and was subsequently used for mitochondrial respiration measurements. Mitochondrial outer membrane integrity of normoxic and hypoxic samples was validated using a spectrophotometric cytochrome c oxidase assay according to the manufacturer's instructions (Sigma, St. Louis, MO).
Respiratory rates were polarographically measured using a Clark-type electrode (Hansatech Instruments, London, UK) at 25°C with constant stirring as previously described (6) with modifications. Briefly, isolated rat ventricular mitochondria (0.5 mg/ml) were added to the electrode chamber containing incubation medium (10 mM Tris·HCl, 0.25 M sucrose, 8.5 mM KH2PO4, pH 7.4). We employed a mixture of 5 mM malate and 40 μM palmitoyl-l-carnitine as oxidative substrates. State 3 respiration was determined by measuring mitochondrial oxygen uptake after the addition of ADP to a final concentration of 350 μM. State 4 respiration was determined by measuring mitochondrial oxygen uptake on complete phosphorylation of ADP to ATP. Basal proton leak in the isolated mitochondria was determined by measuring the rate of mitochondrial respiration after addition of 10 μg/ml oligomycin to state 4 mitochondria.
The ADP-to-O ratio (ADP/O), a measure of mitochondrial oxidative phosphorylation efficiency, was calculated as the ratio between the ADP added and oxygen consumed during ADP phosphorylation. The rate of ADP phosphorylation was calculated as nanomoles of ADP phosphorylated per minute during state 3 respiration as described before (2). Mitochondria were considered viable where the respiratory control index (RCI) (state 3/state 4) was ≥4. The RCI and the ADP/O ratios were calculated according to Estabrook (7) using 253 nmol O2 /ml as the value for the solubility of oxygen at 25°C. All mitochondrial polarographic studies were normalized to total mitochondrial protein content, determined using the Lowry assay (16).
Langendorff heart perfusions.
Isolated hearts were perfused in the Langendorff mode with ice-cold Krebs-Henseleit buffer (11 mM glucose, 118 mM NaCl, 25 mM NaHCO3, 4.7 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4·7H2O, 1.8 mM CaCl2·6H2O, pH 7.4). The aorta was located and cannulated on the Langendorff perfusion rig, and a retrograde perfusion of the coronary arteries via the aorta was immediately initiated. The perfusion was performed with oxygenated (95% O2-5% CO2) Krebs-Henseleit buffer at a constant pressure (104 cmH2O) and temperature (37°C). During perfusion, a latex balloon attached to a pressure transducer was first inserted into the LV cavity for stabilization and inflated to produce a diastolic pressure of 4–12 mmHg, whereupon the balloon was then inserted into the RV to determine the RV developed pressure, as described before (28). The RV and LV functional parameters were measured and included heart rate, systolic and diastolic pressure, developed pressure, coronary flow, and rate-pressure product (heart rate × developed pressure).
RNA isolation and real-time quantitative RT-PCR analysis.
RNA extraction and real-time quantitative RT-PCR of samples were performed using previously described methods (26). Specific quantitative Taqman assays were designed from rat sequences available in GenBank. We determined transcript levels of the following: cytochrome c oxidase subunits II and IV (COXII and COXIV, respectively); nuclear respiratory factor-1 (NRF-1), a well-described transcriptional modulator regulating expression of several mitochondrial proteins; peroxisome proliferator-activated receptor-γ-coactivator-1α (PGC-1α), a transcriptional coactivator controlling cellular energy metabolic pathways (31); and cardiac-enriched uncoupling proteins-2 and -3 (UCP2 and UCP3, respectively) (27, 12). Primer and probe sequences for PGC-1α, UCP2, and UCP3 have been published previously (26, 15), and primer and probe sequences for COXII, COXIV, and NRF-1 are presented in Table 1. Standard RNA was made for all assays by the T7 polymerase method (Ambion, Austin, TX), using total RNA isolated from rat hearts. The correlation between the Ct (the no. of PCR cycles required for the fluorescent signal to reach a detection threshold) and the amount of standard was linear over at least a 5-log range of RNA for all assays (data not shown). Gene expression data are represented as mRNA molecules per nanogram of total RNA.
Citrate synthase activity.
Citrate synthase activity was used as a marker for mitochondrial number and was determined according to an established spectrophotometric assay (25). Isolated RV and LV mitochondria (100 μg) were incubated in 1 ml of a Tris-based buffer [100 mM Tris (pH 7.4), 10 mM acetyl-CoA, 1 mM 5,5′-dithiobis-2-nitrobenzoic acid (DTNB)]. The assay is based on the reaction of citrate synthase with oxaloacetate and acetyl-CoA to produce coenzyme A (CoASH). DTNB reacts with sulfhydryls in CoASH, producing a free thionitrobenzoate ion (23). The absorbance of the suspension was measured at 412 nm (25°C). After a baseline setting, 100 μM oxaloacetate was added, and measurements were taken as before. The difference in absorbance before and after the addition of oxaloacetate was used as a measure of mitochondrial citrate synthase activity.
Mitochondrial DNA determination.
Mitochondrial DNA content was measured in pretreated homogenates by real-time PCR (primers sequences: 5′-TACACGATGAGGCAACCAAA-3′, 5′-GGTAGGGGGTGTGTTGTGAG-3′) as described before (14a). Data are expressed as mtDNA per gram of heart tissue.
Data are presented as means ± SE. Statistically significant differences between normoxic and hypoxic groups were calculated using Students t-test. Statistical significance was considered when P < 0.05.
Effects of hypoxia on morphometrics and cardiac contractile function.
The body weight (BW) was lower in the hypoxic group, but this difference did not reach statistical significance (Table 2). Heart weight (HW) was increased following exposure to hypobaric hypoxia (n = 6, P < 0.01 vs. normoxic control). The increase in HW reflects a robust hypertrophic response in the RV, as indicated by the higher RV/LV+S ratio (n = 6, P < 0.01 vs. normoxic control).
Exposure to chronic hypoxia did not significantly affect the heart rate (Table 3). However, coronary flow was increased in the RV in response to hypobaric hypoxia (n = 6, P < 0.05 vs. normoxic control). Chronic hypoxia increased RV developed pressure (n = 6, P < 0.01 vs. normoxic control) (Table 3). The increase in RV developed pressure was accompanied by an 80 ± 9.9% increase in RV rate-pressure product (n = 6, P < 0.01 vs. normoxic control). LV functional parameters did not significantly change following exposure to hypobaric hypoxia.
Cardiac mitochondrial respiration.
Following hypobaric hypoxia, RV mitochondrial state 3 respiration was increased by 32.9 ± 2.2% (n = 6, P < 0.05 vs. normoxic control) (Fig. 1A). State 3 mitochondrial respiration was not significantly increased in the LV in response to hypobaric hypoxia. The ADP/O ratio was not significantly different in the RV and LV after exposure to hypobaric hypoxia (Fig. 1B).
Following exposure to hypobaric hypoxia, there was no significant difference in the rate of mitochondrial ADP phosphorylation in the RV and LV (Fig. 1C). We next measured state 4 mitochondrial respiration as a marker for mitochondrial uncoupling. Here, we show that hypobaric hypoxia had no effect on state 4 respiration in the RV and LV (Fig. 1D). We further examined mitochondrial proton leak by adding oligomycin (inhibitor of mitochondrial electron transport chain complex V) to state 4 mitochondria. We found decreased proton leak in the RV in response to chronic hypoxia (n = 5, P < 0.05 vs. normoxic control) (Fig. 1E). In contrast, proton leak was not significantly altered in the LV following hypobaric hypoxia. Exposure to chronic hypoxia had no significant effect on the RCI (Fig. 1F).
Effects of hypoxia on cardiac metabolic gene expression.
Following exposure to chronic hypobaric hypoxia for 2 wk, COXII transcript levels were increased by 72 ± 10.6% in the RV (P < 0.05 vs. normoxic control) (Fig. 2A). RV COXIV transcript levels were not significantly increased, whereas LV COXIV levels were elevated (n = 6, P < 0.01 vs. normoxic control) following exposure to hypobaric hypoxia (Fig. 2B). However, both PGC-1α and NRF-1 transcript levels were significantly increased in the hypertrophied RV in response to hypoxia (Fig. 2, C and D). LV NRF-1 transcript levels were not altered for any of the experimental groups. UCP2 levels were significantly induced in the RV and LV following hypoxic exposure (Fig. 2E), while UCP3 expression was reduced by 47 ± 8.4% in the hypertrophied RV (Fig. 2F) (n = 6, P < 0.05).
Effects of hypoxia on citrate synthase activity and mtDNA content.
We spectrophotometrically measured citrate synthase activity as a marker of mitochondrial content. In agreement with our transcript data, citrate synthase activity was markedly increased in the RV in response to chronic hypoxia (n = 6, P < 0.01 vs. normoxic control) (Fig. 3A). Although LV citrate synthase activity was higher after hypoxia, this did not reach statistical significance. In agreement, mtDNA content was higher in the hypertrophied RV vs. control samples (n = 4, P < 0.05) (Fig. 3B). No significant changes were observed in the LV following the hypoxic exposure.
We hypothesized that mitochondrial regulatory genes are induced in response to hypoxia-mediated RV hypertrophy as part of an adaptive response to sustain mitochondrial respiratory capacity and contractile function in response to increased load. To test our hypothesis, we exposed rats to 2 wk of hypobaric hypoxia and determined cardiac contractile and mitochondrial respiratory function for RV and LV, respectively. The main finding of this study is the coordinate induction of several genes regulating mitochondrial function and increased mtDNA content in the hypertrophied RV, linking the efficiency of mitochondrial oxidative phosphorylation and enhanced respiratory function to increased RV contractile function.
Exposure to chronic hypobaric hypoxia results in pulmonary hypertension and increased RV load. Our data demonstrate a robust hypertrophic response in the RV in response to chronic hypobaric hypoxia. These data are in agreement with our earlier findings showing increased myocyte diameter size and ANF expression after 2 wk of hypobaric hypoxia (22). Moreover, we also reported a lack of fibrosis, suggesting adaptive physiological remodeling in the RV in response to pulmonary vasoconstriction. Since the LV is not challenged by increased load in this instance, higher hematocrit levels and/or neuroendocrine regulation may be key factors influencing adaptation.
Our respiration and gene expression studies show distinct remodeling in the RV and LV in response to the hypoxic stimulus. Here, we found that RV mitochondrial oxygen consumption was increased with chronic hypoxia. These data are in agreement with previous studies reporting sustained respiratory capacity in the RV in response to chronic hypobaric hypoxia (20, 18). Moreover, others have shown increased and/or sustained rates of ATP synthesis in response to chronic hypoxia (19, 5, 4). We also found that exposure to hypobaric hypoxia improved the efficiency of mitochondrial respiration (↓ proton leak) in the hypertrophied RV. In parallel, transcript levels of NRF-1, PGC-1α, COX II, and UCP2 were coordinately upregulated while UCP3 was downregulated in the RV. COX is the terminal enzyme of the mitochondrial electron transport chain catalyzing the transfer of electrons from cytochrome c to molecular oxygen. Its subunits are encoded by both nuclear (e.g., COXIV) and mitochondrial genomes (e.g., COXII) (11). NRF-1 and PGC-1α are central transcriptional regulators of mitochondrial biogenesis and are good candidate factors for further investigation (29). Interestingly, a recent study suggested that cyclic nucleotide regulatory element binding protein (CREB) may be a pivotal transcriptional modulator that regulates physiological hypertrophy by enhancing expression of genes important for efficient oxidative capacity and resistance to apoptosis (30). Moreover, CREB is proposed to mediate such effects via control of PGC-1 expression and subsequent mitochondrial biogenesis (32). In addition, other transcriptional modulators such as PGC-1β, NRF-2, Sp1 AP1, YY1, and TFAM are also likely candidates that may play a role in the induction of this gene program (reviewed in Refs. 9, 21).
In contrast, exposure to chronic hypobaric hypoxia had no significant effects on LV mitochondrial respiration or contractile function. However, COXIV and UCP2 expression was increased in response to chronic hypobaric hypoxia. Since the LV is not exposed to increased load in our experimental model, we propose that these genes are regulated by hypoxia-mediated transcriptional mechanisms that may result in a lesser degree of mitochondrial biogenesis.
Our data suggest that the adaptive mitochondrial phenotype in the RV is largely mediated via load-dependent mechanisms. In support, we found significantly increased citrate synthase activity and mtDNA content only in the hypertrophied RV. As the trophic response results in a larger cell volume, mitochondria must proliferate to ensure an adequate supply of mitochondrial energy required for cell maintenance and contractile purposes. In agreement, increased mitochondrial numbers have been found in hypertrophic hearts (reviewed in Ref. 10). However, it has been proposed that an imbalance between inadequate mitochondrial proliferation and increased energy demands (due to high workloads) may contribute to the onset of pathological hypertrophy. Here, we suggest that the hypertrophied RV adapts by increasing the expression of mitochondrial proteins leading to enhanced respiratory capacity and more efficient mitochondrial energy production.
What are the mechanisms responsible for increased efficiency of mitochondrial ATP production? We found reduced UCP3 expression in the hypertrophied RV, thus providing a potential mechanism whereby proton leak may be reduced. However, further studies are required to confirm this observation, since it has also been proposed that UCP3 does not necessarily act as a classic uncoupler of oxidative phosphorylation in the heart (12). Recently, it was reported that exposure to hypobaric hypoxia resulted in reduced opening of the mitochondrial permeability transition pore (MPTP) (34). Since opening of the MPTP will lead to the dissipation of the electrochemical proton gradient across the inner mitochondrial membrane, we propose that increased MPTP closure could also potentially contribute to improved efficiency of mitochondrial ATP production in the hypertrophied RV. In agreement with such an adaptive mitochondrial phenotype, we found a hypoxia-mediated induction of UCP2 transcript levels in the RV. Recent studies suggest that UCP2 is more likely to be part of a defense mechanism against damaging reactive oxygen species instead of a true uncoupler of mitochondrial oxidative phosphorylation in the heart (27, 13).
Relatively few studies have examined contractile function of the RV and LV in response to hypobaric hypoxia to assess the functional significance of described changes. In this study, we found that exposure to chronic hypobaric hypoxia increased contractile function in the hypertrophied RV. Moreover, we found that LV contractile function was sustained with chronic hypoxia. Taken together, these data indicate distinct responses by the RV and LV to sustain cardiac function in response to chronic hypobaric hypoxia. Here, the hypertrophied RV induces expression of several mitochondrial regulatory genes that are associated with increased respiratory function and contractility. On the other hand, the LV displayed sustained respiratory function and cardiac contractility. Moreover, expression of COXIV and UCP2 was increased while citrate synthase activity was not significantly altered, thus indicating that mitochondrial biogenesis largely occurred in the hypertrophied RV.
What are the broader implications of the findings of the present study? Since most changes were observed only in the hypertrophied RV (and not the LV), we believe our data may be extended to physiological hypertrophied hearts in general. Our data are in agreement with views expressed in a recent review article (8) where it was proposed that physiological hypertrophy is coupled with increased PGC-1α expression and greater mitochondrial oxidative capacity. Furthermore, the authors suggested that pathological hypertrophy is associated with reduced PGC-1α and dysfunctional mitochondria. These interesting possibilities require further investigation.
A potential limitation of this study is that our respiration studies were performed using isolated mitochondrial preparations. Ideally, intact mitochondrial preparations (e.g., saponin-permeabilized strips) should be employed to prevent the potential selection of artificial mitochondrial subpopulations.
In summary, we found coordinate induction of several genes regulating mitochondrial function and mitochondrial content in a model of physiological RV hypertrophy, linking the efficiency of mitochondrial oxidative phosphorylation and respiratory function to sustained RV contractile function in response to increased load.
This work was supported by the South African Medical Research Council and the South African National Research Foundation (M. F. Essop) as well as the National Heart, Lung, and Blood Institute (grant HL-074259-01) (M. E. Young).
We wish to thank Noel Markgraaff for technical assistance.
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.
- Copyright © 2007 by the American Physiological Society