Mitochondrial dysfunction has been implicated as a cause of energy deprivation in heart failure (HF). Herein, we tested individual and combined effects of two pathogenic factors of nonischemic HF, inhibition of nitric oxide synthesis [with l-NG-nitroarginine methyl ester (l-NAME)] and hypertension [with angiotensin II (AngII)], on myocardial mitochondrial function, oxidative stress, and metabolic gene expression. l-NAME and AngII were administered individually and in combination to mice for 5 wk. Although all treatments increased blood pressure and reduced cardiac contractile function, the l-NAME + AngII group was associated with the most severe HF, as characterized by edema, hypertrophy, oxidative stress, increased expression of Nppa and Nppb, and decreased expression of Atp2a2 and Camk2b. l-NAME + AngII-treated mice exhibited robust deterioration of cardiac mitochondrial function, as observed by reduced respiratory control ratios in subsarcolemmal mitochondria and reduced state 3 levels in interfibrillar mitochondria for complex I but not for complex II substrates. Cardiac myofibrils showed reduced ADP-supported and oligomycin-inhibited oxygen consumption. Mitochondrial functional impairment was accompanied by reduced mitochondrial DNA content and activities of pyruvate dehydrogenase and complex I but increased H2O2 production and tissue protein carbonyls in hearts from AngII and l-NAME + AngII groups. Microarray analyses revealed the majority of the gene changes attributed to the l-NAME + AngII group. Pathway analyses indicated significant changes in metabolic pathways, such as oxidative phosphorylation, mitochondrial function, cardiac hypertrophy, and fatty acid metabolism in l-NAME + AngII hearts. We conclude that l-NAME + AngII is associated with impaired mitochondrial respiratory function and increased oxidative stress compared with either l-NAME or AngII alone, resulting in nonischemic HF.
- heart failure
- nitric oxide
- angiotensin II
- oxidative stress
NEW & NOTEWORTHY
Inhibition of nitric oxide synthesis is an important pathogenic factor of nonischemic heart failure. Our study shows the importance of nitric oxide inhibition in the development of l-NG-nitroarginine methyl ester + angiotensin-mediated heart failure by characterizing the underlying mitochondrial, oxidative, and metabolic mechanisms.
heart failure (HF) affects 5.1 million people in the US, and, despite the medical progress in neurohormonal treatments, mortality rates are close to 59% (21). Broadly, HF has two main etiologies, ischemic, which includes myocardial damage after a myocardial infarct or arterial occlusion, and nonischemic, which includes hypertensive heart disease, myocarditis, arrhythmia, etc. (15). Mechanistically, with either ischemia or pressure overload, energy deprivation is a common characteristic that is attributed to myocardial mitochondrial dysfunction (22, 24, 33).
The renin-angiotensin II (AngII) aldosterone system plays a major role in the pathogenesis and progression of HF. Some of the present most beneficial HF therapies include drugs that inhibit the AngII-converting enzyme, AngII receptor, or aldosterone action (16). In mice, AngII treatment leads to nonischemic cardiomyopathy, wherein hypertension, hypertrophy, and cardiomyopathy result from increased vasoconstriction, fibrosis, and activation of the immune system (35). Conversely, AngII receptor blockers or inhibitors of AngII biosynthesis reverse ventricular hypertrophy, preventing HF (7). NG-nitro-l-arginine methyl ester (l-NAME) inhibits nitric oxide (NO) synthesis, which is important in maintaining vascular tone by inhibiting smooth muscle contraction and platelet aggregation, resulting in hypertension (4). HF is associated with uncoupling of NO synthase (NOS), such that there is greater production of superoxide than NO (4). The contributions of myocardial mitochondrial mechanisms underlying the individual and combined effects of l-NAME and AngII to HF are yet to be determined.
Mitochondrial dysfunction has been considered a primary cause of energy depletion and the resulting contractile dysfunction in HF (22, 24, 33). Our understanding of the mechanisms underlying HF comes primarily from animal models (28), such as those with myocardial ischemia resulting from either permanent coronary ligation or intermittent ischemia-reperfusion injury, pressure overload hypertrophy by transverse aortic constriction (TAC), and genetically induced models of cardiomyopathy. Such models of acute, ischemic HF have shown that the mitochondrial OXPHOS system is compromised at the levels of both expression and activity of the respiratory chain complexes (17, 46). Our group has recently shown that the combination of l-NAME and AngII results in nonischemic HF, and the model is independent of any major surgical trauma (6).
In this study, we determine the bioenergetic changes induced by l-NAME and AngII alone and in combination that underlie the development and progression of nonischemic HF. We hypothesized that the combined effects of hypertension and inhibition of NO would induce mitochondrial dysfunction and oxidative stress that would increase the severity of HF. Our results show that the combination of l-NAME and AngII, rather than l-NAME or AngII individually, is associated with more severe characteristics of HF, accompanied by mitochondrial functional aberrations and oxidative stress. An understanding of the bioenergetics of nonischemic HF may provide insight into new therapeutic opportunities that modify mitochondrial dysfunction and improve the redox state to prevent the progression from compensated hypertrophy to HF.
MATERIALS AND METHODS
All reagents for mitochondrial studies were purchased from Sigma-Aldrich (St. Louis, MO). Blood glucose was measured with a One-Touch glucometer. Microarray chips were purchased from Illumina (San Diego, CA). All other materials and the companies they were purchased from are referenced in the methods below.
Animals and treatments.
Twelve-week-old C57BL6 mice (n = 30 total) were randomly assigned to four groups: Group 1, control (n = 6); Group 2, l-NAME treatment (n = 8); Group 3, AngII treatment (n = 7); Group 4, l-NAME + AngII (n = 9) treatment. l-NAME (0.3 mg/ml with 1% NaCl) was administered in drinking water, whereas control and AngII-treated mice received plain drinking water. For the combination and AngII alone treatment Groups 3 and 4, 1 wk after ll-NAME treatment was continued. Groups 1 and 2 received pumps filled with PBS. All mice were housed in microisolation cages under a 12-h:12-h light/dark cycle and were fed a standard chow diet (Harlan Teklad, 2920). At the end of week 4 of the study, blood pressure was measured; at week 5, mice were anesthetized with isoflurane, and cardiac function was assessed with echocardiography. Body weight, body fat, and water content were assessed by EchoMRI-3in1 small animal Quantitative Magnetic Resonance. Mice were then fasted overnight and killed by cervical dislocation. Hearts were immediately excised, weighed, and dissected for left ventricle (LV) separation and division into pieces. The largest section of the LV tissue was maintained in ice-cold BIOPS buffer (2.77 mM CaK2EGTA, 7.23 mM K2EGTA, 5.7 mM Na2ATP, 6.56 mM MgCl2·6 H2O, 20 mM taurine, 15 mM Na2phospho-creatine, 20 mM imidazole, 0.5 mM dithiothreitol, 50 mM MES, pH 7.1; Oroboros Instruments, Innsbruck, Austria) for mitochondrial respiratory studies and H2O2 (reactive oxygen species, ROS) production assays. The other sections were either frozen for gene/protein expression and enzyme assays or fixed in 4% paraformaldehyde for fibrosis staining. To obtain enough sample material for each assay, the experiment was repeated in two cohorts. All animal protocols were approved by the Houston Methodist Research Institute's Institutional Animal Care and Use Committee and carried out in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
Paraffin-embedded heart tissue was sectioned and stained with Masson's Trichrome stain (Sigma-Aldrich) and quantified as area percentage with blue staining.
Cross-section slices of the mouse hearts were stained with wheat germ agglutinin (Sigma-Aldrich). Images were views on a Nikon fluorescence microscope for FITC (excitation 488/emission 510) (Tokyo, Japan). Cardiomyocyte size was manually quantified by measuring the shortest diameter of all cells in cross-sectional orientation using the Nikon NIS elements acquisition program. At least 150 cells were measured per field, three fields per sample for four samples per group, which were blinded to the two individuals performing the measurements. The average cardiomyocyte size is reported in microns for each group.
Cardiac function measurements.
Blood pressure was measured using Visitech Automated Blood Pressure Monitoring System (Apex, NC) by tail cuff, wherein mice were acclimated for 2 days and readings were obtained on the third day. Transthoracic echocardiography was performed and analyzed as previously described (26, 32, 38) using Vevo770 high-resolution imaging system equipped with a 30-MHz transducer (Visualsonics, Toronto, Ontario, Canada). Mice were anesthetized with 0.5–1.0% isoflurane for the duration of the recording. Two-dimensional targeted M-mode echocardiographic images were obtained at the levels of the papillary muscles from the parasternal short-axis view for measurements of LV wall thickness, LV end-diastolic diameter (LVEDD), and LV-end-systolic diameter (LVESD) using at least three consecutive heart beats. All M-mode parameters were read from the same positions on the LV. LV fractional shortening (FS) was calculated as FS (%) = [(LVEDD-LVESD)/LVEDD] × 100. With the use of the LVEDD and LVESD values, the LV volume at diastole and systole were determined as volume = (7.0/2.4 + d)(d3), where d is diameter. Ejection fractions (EF) were calculated as EF (%) = [(LVEDV − LVESV)/LVEDV] × 100 as measures of systolic function. Hypertrophy was determined from heart weight to tibia length ratio.
Mitochondrial functional analyses.
Mitochondrial respiratory function was assessed with Oroboros high-resolution respirometry using isolated mitochondrial subpopulations (0.2 mg/ml) and permeabilized cardiac fibers. Mitochondria were isolated from freshly excised hearts as described before the use of a differential centrifugation procedure (18, 20). Briefly, hearts were washed with ice-cold isolation buffer (buffer A, 220 mmol/l mannitol, 70 mmol/l sucrose, 5 mmol/l MOPS, pH 7.4), and tissue was cut into small pieces, homogenized in buffer A, and centrifuged (800 revolution/min for 10 min at 4°C). The pellet was treated with trypsin (5 mg/g wet weight of tissue) for 10 min on ice, followed by brief homogenization to separate the interfibrillar mitochondria (IFM). The reaction was stopped with double the volume of ice-cold buffer B (2 mM EGTA, 0.2% free fatty acid-free-BSA in buffer A) and centrifuged (800 revolution/min for 10 min at 4°C). The supernatants from the first centrifugation and from the pellet treatment were centrifuged separately at 13,000 revolution/min for 10 min. The resulting pellets were resuspended in cold buffer B and again centrifuged at 13,000 revolution/min for 10 min. The pellets were rinsed with buffer A, centrifuged for the same time and speed, after which the pellet was resuspended in 30 μl of cold buffer E (0.05 mM of EGTA in buffer A). The mitochondrial isolate obtained without trypsin treatment contained primarily subsarcolemmal mitochondria (SSM). Cardiac fibers were teased out from small pieces of heart tissue in cold buffer A and permeabilized with saponin (50 μg/ml) for 30 min on ice before the use of 10 mg of fibers for respiratory analyses.
For isolated mitochondrial studies, a single substrate protocol with unsaturating levels of ADP [for palmitoyl carnitine malate (PC), pyruvate malate (PM)], and for permeabilized fibers, a multiple substrate-protocol using saturating levels of ADP [for glutamate malate (GM), succinate] was used, modified from previously described protocols (20, 29). Mitochondria or fibers were suspended in MiRO5 medium (0.5 mM EGTA, 3 mM MgCl2·6 H2O, 60 mM K-lactobionate, 2 mM taurine, 10 mM KH2PO4, 20 mM HEPES, 110 mM sucrose, 1 g/l fatty acid-free BSA, pH 7.1) in the oxygraph chambers. For the single substrate protocols, either PM (5 mM each), PC (5 mM each), GM (5 mM) or succinate (5 mM) was added, followed by 100 μM ADP. State 2 respiration rate (ADP independent) was determined as respiration with substrate alone and state 3 as ADP-dependent respiration. Respiratory control ratios (RCR), an index of respiratory efficiency, were calculated as the ratio of state 3 (ADP dependent) to state 2 (ADP independent) respiration. All readings were normalized for mitochondrial protein content, as determined by Lowry assay. For the multisubstrate protocol, PM (5 mM), ADP (4 mM for saturation), succinate (5 mM), and oligomycin (1.5 μg) were sequentially added to measure maximum oxidative phosphorylation capacity and coupled respiration. All data for fibers were normalized for milligrams of permeabilized fibers.
DNA was isolated from LV samples (DNA extraction kit; Qiagen, Valencia, CA) and analyzed by quantitative real-time PCR to assess the DNA ratio of the mitochondrial-encoded gene Nd5 to the nuclear-encoded gene Gapdh.
Protein carbonyls were measured as stable markers of chronic oxidative stress in whole LV tissue using the OxiSelect Protein Carbonyl ELISA from Cell Biolaboratories (San Diego, CA). H2O2 production was determined in mitochondrial isolates using the Amplex Red kit from Life Technologies (Carlsbad, CA) in response to PM, antimycin A, and rotenone.
Heart RNA was isolated using RNeasy kits (Qiagen), reversed-transcribed into cDNA with High-Capacity cDNA Reverse Transcription kits (Applied Biosystems, Grand Island, NY), and analyzed for candidate gene expression using TaqMan PCR Core reagent kits and gene-specific primer/probe sets (Applied Biosystems). Candidate gene expression was normalized to an average of the expression of Ppia, Tptl, and Prl13a (44) because these genes were found to be the most stable across the four treatment groups compared with other housekeeping genes Gapdh, 18S, and Hprt (13).
Enzyme activity assays.
Pyruvate dehydrogenase (PDH) and complex I enzyme activities were measured with microplate assay kits from Abcam (Cambridge, MA). Briefly, the enzymes were immunocaptured from whole tissue lysates in microplate wells using respective antibodies. PDH activity was determined by following the reduction of NAD+ to NADH, coupled to the reduction of a reporter dye to yield a corresponding increase in absorbance of a colored reaction product at 450 nm. The activity for Complex I was determined by following the oxidation of NADH to NAD+ and the simultaneous reduction of a dye, which leads to increased absorbance at 450 nm.
Microarray analyses were performed as previously described (14). Briefly, mouse WG-6 v2 whole-genome expression arrays were purchased from Illumina (San Diego, CA), and cRNA synthesis and labeling were performed using Illumina TotalPrep-96 RNA Amplification Kit (Ambion, Austin, TX), with the labeling in vitro transcription reaction performed at 37°C for 14 h. Biotinylated cRNA samples were hybridized to arrays at 58°C for 18 h according to the manufacturer's protocol. Arrays were scanned using BeadArray Reader (Illumina). Unmodified microarray data obtained from GenomeStudio were background subtracted and quantile normalized using the Lumi package (11) and analyzed with the Limma package (37) within R (30). All analysis was corrected for multiple hypotheses testing (2), and effects were determined to be significant when there was a ≥1.5-fold increase/decrease relative to the control and they had an adjusted P value <0.01 and FDR <0.25. Raw microarray data sets have been deposited in the NCBI Gene Expression Omnibus (GEO) database under accession number GSE72132. Microarray data was also validated through qPCR (TaqMan) analysis, showing excellent correlation R2 = 0.87 (n = 69).
One-way ANOVA followed by Tukey's multiple-comparison test were performed to identify differences between groups. A two-tailed α = 0.05 was used as the significance cutoff for all tests. Data are presented as means ± SE, and sample sizes are reported in figure legends. GraphPad Prism 5.0 software was used for all statistical analyses.
Loss of body weight, body fat but greater water retention in l-NAME + AngII-treated mice.
Body weight and composition changes were determined for mice treated with vehicle (n = 6), l-NAME (n = 8), AngII (n = 7) and l-NAME + AngII (N = 9). The mice treated with l-NAME + AngII lost significant body weight compared with controls (Fig. 1A). Additionally, the percentage of body fat content (Fig. 1B) decreased in AngII- and l-NAME + AngII-treated mice after 5 wk of treatment, potentially attributable to reduced appetite and vigor associated with HF. There was no difference in percentage of lean mass between any groups (Fig. 1C). Percentage of free water, indicative of edema, was significantly increased compared with control only in the l-NAME + AngII-treated mice (Fig. 1D). These data indicate that l-NAME + AngII treatment had the greatest systemic effect with development of HF-related edema, change in body composition, and weight loss among the four treatment groups.
l-NAME + AngII-treated mice exhibit the most severe HF characteristics.
To characterize the cardiac function of the four treatment groups, echocardiography, blood pressure, pulse rate, heart weight and fibrosis assessment were performed in control (n = 6), l-NAME-treated (n = 8), AngII-treated (n = 7), and l-NAME + AngII-treated (n = 9) mice (Fig. 2). Systolic blood pressure, measured via tail cuff, was greater in AngII and l-NAME + AngII groups compared with controls, with the highest in l-NAME + AngII-treated mice (Fig. 2A). Diastolic pressure was greater in AngII group and significantly greater in the l-NAME + AngII group (Table 1). The calculated mean arterial pressure was greater than controls in AngII and l-NAME + AngII groups (Table 1). Heart weight:tibia length ratio, an indicator of cardiac hypertrophy, was significantly increased only in the l-NAME + AngII group compared with control (Fig. 2B). The size of the cardiomyocytes was the largest in the l-NAME + AngII group, indicative of hypertrophy (Fig. 2, C and D). Cardiomyocyte size tended to be greater in the l-NAME + AngII group than AngII group (P = 0.09). Heart rates measured in awake mice during blood pressure measurement were suppressed in both l-NAME-treated groups (Table 1). Surprisingly, hearts rates of mice anesthetized with isoflurane did not exhibit selective l-NAME-induced bradycardia; instead isoflurane suppressed hear rates to about 400 beats/min in all four groups (Table 1). Echocardiography performed in control (n = 4), l-NAME-treated (n = 8), AngII-treated (n = 6), and l-NAME + AngII-treated (n = 7) mice showed that FS (Fig. 2E) and EF (Fig. 2F) were significantly suppressed in all three treatment groups vs. controls, and l-NAME + AngII-treated mice had the lowest FS of 18.72 ± 0.03% and lowest EF of 38.52 ± 0.05% among the four groups. LV posterior wall thickness tended to be greater in the l-NAME + AngII group for diastole and systole (Table 1). The LV internal diameter and volume were significantly greater than control only in the l-NAME + AngII group and tended to be greater than control in the AngII group (Table 1). Fibrosis was dramatically increased in hearts from AngII- and l-NAME + AngII-treated mice compared with control and l-NAME-treated mice (Fig. 2, G and H). Although the changes in HF characteristics were the largest and most significant in the l-NAME + AngII-treated group, all were not statistically significantly different from the AngII group. Taken together, the combination of l-NAME and AngII tended to have exaggerated cardiac dysfunction than either l-NAME or AngII alone.
l-NAME + AngII-treated mice have mitochondrial functional aberrations.
Mitochondrial function was assessed in SSM and IFM subpopulations, as well as permeabilized fibers from hearts of control (n = 6), l-NAME-treated (n = 8), AngII-treated (n = 7), and l-NAME + AngII-treated (n = 9) mice. In SSM (Fig. 3), ADP-independent state 2 respiration rates were greater than control for l-NAME and l-NAME + AngII mice only when GM was used as substrate (Fig. 3, A, D, G, and J), whereas state 3 rates were significantly lower in the l-NAME + AngII SSM than those for control and AngII groups for PM and PC substrates (Fig. 3, B, E, H, and K). RCRs (state 3/state 2) for PM were also lower in l-NAME + AngII SSM than control and AngII. The RCRs for l-NAME + AngII SSM were lower than control for PC and GM substrates and lower than l-NAME for GM (Fig. 3, C, F, I, and L). Respiration rates did not differ for succinate among the four groups. Taken together, among the four study groups, l-NAME + AngII SSM showed the most consistent downregulation of mitochondrial function for primarily complex I-associated substrates.
For IFMs, state 2 rates did not differ between groups for any substrate (Fig. 4, A, D, G, and J). State 3 rates were surprisingly higher for AngII IFM than controls and lower for l-NAME + AngII IFM than AngII with PM (Fig. 4B). State 3 rates were also lower than controls and AngII for l-NAME + AngII IFM with PC and GM substrates (Fig. 4, E and H). However, none of the RCRs or succinate-associated respiration rates were significantly different among the four groups.
Whereas SSM and IFM analyses reveal the functional aspects of the individual isolated populations, permeabilized fibers can identify the overall energetic changes within the architecture of the cardiomyocyte fibrils. In a multisubstrate protocol, respiration rates with PM alone were significantly lower in l-NAME, AngII, and l-NAME + AngII compared with controls (Fig. 5A). With the addition of ADP, the respiration rates remained lower than those for controls for the other three groups; however, statistical significance was seen only in the l-NAME + AngII fibers (Fig. 5B). Furthermore, with addition of succinate, the rates continued to remain lower than control, again significantly reduced only for l-NAME + AngII fibers (Fig. 5C). Upon treatment with oligomycin to inhibit complex V, fibers from all three treatment groups had significantly lower respiration than controls (Fig. 5D). The respiratory control ratios for PM (state 3/state 2) and for PM + succinate (state 3/oligomycin-induced state 4) were suppressed significantly in l-NAME + AngII group (Fig. 5, E and F). Thus, although data with PM alone and oligomycin may indicate that the three treatment groups are more coupled with less leak than controls, the overall reduction in ADP-dependent respiration suggests that there is downregulation of the maximal energy-generating capacity of hearts from all three treatment groups, with the most significant changes in the l-NAME + AngII fibers.
Mitochondrial dysfunction is associated with impaired PDH and complex I activities and increased oxidative stress in the l-NAME + AngII mice.
Mitochondrial DNA content, a measurement of mitochondrial content, was significantly reduced in hearts from AngII- and l-NAME + AngII-treated mice (Fig. 6A). Whole LV tissue protein carbonyls, a measurement of chronic tissue oxidative stress, were significantly increased in AngII and l-NAME + AngII mice compared with controls and l-NAME. l-NAME + AngII-treated mice also had higher protein carbonyls than AngII-treated mice (Fig. 6B). Mitochondrial production of ROS (H2O2) was the greatest in AngII and l-NAME + AngII groups upon exposure to PM, complex I inhibitor rotenone, and complex III inhibitor antimycin A (Fig. 6C). Activity levels of PDH and complex I, but not PDH quantity, were suppressed in l-NAME + AngII-treated mice (Fig. 6, D–F). Therefore, energy depletion in the hearts of l-NAME + AngII mice results from reduced mitochondrial content as well as inefficient and dysfunctional mitochondria. All measurements were performed in control (n = 6), l-NAME-treated (n = 8), AngII-treated (n = 7), and l-NAME + AngII-treated (n = 9) mice.
Transcriptomic profiling of hearts reveal distinct gene regulation patterns for each treatment.
To determine the mechanism of action of each treatment, microarray gene expression profiling was performed on the four study groups, followed by bioinformatics analyses. Samples were randomly pooled within each group to give n = 4/group for the microarray chip. Ingenuity Pathway Analysis (IPA) was conducted with the gene expression datasets to identify pathways and biological processes differentially regulated by the three treatments compared with controls. Figure 7 shows a gene heat map with fold changes from controls of selected canonical metabolic pathways associated with HF. Primary metabolic pathways that regulate cardiac energy generation, such as oxidative phosphorylation, mitochondrial function, fatty acid oxidation, AMPK signaling, and tricarboxylic acid (TCA) cycle, were most consistently altered in the l-NAME + AngII group. HF-related pathways, such as hypertrophy signaling, oxidative stress, fibrosis, and calcium signaling pathways, were also altered most consistently in the l-NAME + AngII group. Table 2 shows differences in the fold changes and number of genes (molecules) for each of the selected canonical metabolic pathways. The l-NAME + AngII group had the most number of molecules differentially expressed from controls for each pathway. In addition, pathways regulating cardiac hypertrophy, hypoxia signaling, oxidative stress, calcium signaling, etc. were also most robustly altered both in terms of significance and number of genes in the l-NAME + AngII group. Among the selected metabolic pathways, l-NAME differentially expressed for NO/ROS, oxidative stress, and calcium signaling pathways. AngII group differentially expressed all selected pathways similar to the l-NAME + AngII group but with much fewer gene changes. Mitochondrial dysfunction pathway showed gene downregulation primarily in the l-NAME + AngII group in complexes I, II, IV, and V, but not in complex III (Fig. 8). The top 30 most significant disease pathways identified by IPA included pathways mainly associated with cell death, cell proliferation, connective tissue development, angiogenesis, etc., all processes with implications in LV remodeling, which changed the most in the l-NAME + AngII group (Table 3). These data highlight the unique gene profile of the combination treatment of l-NAME + AngII that is associated with HF.
Furthermore, of the 1,953 genes that were considered significantly differentially expressed, 1,606 genes changed (744 upregulated, 862 downregulated) exclusively in the mice with the combined treatment of l-NAME + AngII (Fig. 9A). Of the genes differentially expressed by AngII, 313 genes (135 upregulated, 178 downregulated) also changed in the l-NAME + AngII group. Only five genes were differentially expressed in both l-NAME and l-NAME + AngII compared with control. All three treatments shared only 29 genes (11 upregulated, 18 downregulated). These data indicate that, although the majority of the gene regulatory events in HF are associated with l-NAME + AngII, individually they may have only a small effect on gene regulation. Thus l-NAME in combination with AngII is crucial for some of the effects of the HF phenotype in this model. Microarray gene changes were validated with quantitative RT-PCR for representative genes, which increased (Angptl7, Nox4) and decreased (Ccl7, Ucp3) in the l-NAME + AngII group (Fig. 9B). Furthermore, pathway-specific gene expression consistently showed altered expression of HF genes, most dramatically in the l-NAME + AngII group (Fig. 9C). Expression of endothelial NO synthase (eNOS) and neuronal NOS (nNOS) was unchanged across treatment groups (Fig. 9C). Expression of OXPHOS genes was variable, with significantly decreased expression of Ndufs4 and Cox7b in l-NAME and l-NAME + AngII group from control levels. Expression of all complex genes was maintained or increased (Uqcrc2) with AngII treatment. Atp5s expression was decreased in the l-NAME + AngII group (Fig. 9D). Genes for fatty acid oxidation (Hadh, Acaa2) and glucose oxidation (PDHB) decreased in in the l-NAME + AngII group only, whereas Ldha did not change in any group (Fig. 9E).
The mechanisms of HF are complex, and the underlying bioenergetic changes are incompletely understood. In this study, we determined the myocardial bioenergetics that are associated with impaired cardiac function resulting from NO inhibition and hypertension by administering l-NAME and AngII individually and jointly. Although AngII treatment alone had a significant detrimental impact on cardiac function with increased fibrosis, oxidative stress (45), hypertrophy, and edema, the combination of l-NAME + AngII had a more deleterious effect in most HF measurements, including those of mitochondrial dysfunction and altered gene expression. Microarray analyses revealed that the largest portion of differentially expressed genes was exclusive to the combined treatment rather than the individual treatments. Taken together, l-NAME + AngII-induced pressure overload results in nonischemic HF with an impaired free-energy transfer associated with mitochondrial dysfunction and reduced mitochondrial content. These results reveal mechanistic information regarding hypertension-associated nonischemic HF and support a model that could be potentially useful for testing therapeutic interventions.
This mouse model of l-NAME + AngII-induced nonischemic HF may serve well for studying mechanisms and treatments for nonischemic HF in a highly reproducible manner (6). AngII treatment induces vasoconstriction and fibrosis and subsequent hypertension and cardiomyopathy (40). l-NAME treatment causes hypertension attributable to disruption of NO-mediated vascular tone. In patients, a combination of factors, including dysregulation of the renin-angiotensin system and NOS activity, occur, and therefore this model with the combined treatment of l-NAME + AngII closely mimics the human nonischemic HF condition (12, 39).
Mitochondrial dysfunction is one of the earliest events of AngII action leading to pathologic remodeling (27). Although IFMs have been described as the primary mitochondria that regulate ATP synthesis in the heart, our detailed mitochondrial functional data show small differences among groups only in state 3 with PM, PC, and GM for IFMs, whereas we see a more distinct downregulation of function in SSM in the l-NAME + AngII group. Oxygen consumption of permeabilized fibers give information of the overall energy-transfer capacity of the fibers, which is a result of the combined activities of SSMs, IFMs, the relative abundance of the two populations, and the total mitochondrial content. Therefore, with consideration of all these factors together, the overall oxidative energy-transfer capacity or respiratory capacity of the l-NAME + AngII cardiac fibers is significantly reduced from that of the control group. In addition, we find that mitochondrial content is reduced in l-NAME + AngII hearts, and thus fewer mitochondria with suboptimal function result in an energy-deficit state that could translate into the reduced EFs observed in these mice. Surprisingly, we did not see robust mitochondrial respiratory dysfunction in the AngII-treated group, contrary to previous findings (8–10). State 3 respiration was preserved in SSM and increased in IFM, and expression of representative OXPHOS genes was unchanged or increased with AngII treatment, suggestive of an early adaptive phase. However, the microarray data suggested overall changes in OXPHOS and mitochondrial function genes. Although the differences in methodologies of assessment of mitochondrial respiratory function, choice of tissue, and mitochondrial subpopulations make it difficult to directly compare studies, it is likely that differences in AngII doses or duration of treatment may underlie this discrepancy. A longer treatment of AngII or use of a higher daily weight-adjusted AngII dose, e.g., 1.1 mg/kg per day vs. 0.7 mg/kg per day in our model, is likely to cause mitochondrial dysfunction because its beginnings are suggested by the microarray data. Furthermore, in our analysis, we chose to use state 2 instead of state 4 to determine leak because state 4 is accompanied by high substrate and ATP levels. Damaged mitochondria in the preparation, if any, may uptake ATP and convert to ADP by ATPases, contributing to the ADP pool. Intact mitochondria may use this ADP and show oxygen consumption although it is not due to leak.
Dysfunctional mitochondria are the primary source of ROS (1, 42), and our data show increased chronic oxidative stress (increased protein carbonyls) and mitochondrial H2O2 production in l-NAME + AngII hearts. These data are in agreement with previous studies showing that mice that overexpress the mitochondria-targeted catalase but not peroxisomal catalase are protected from AngII-induced oxidative stress and associated cardiomyopathy (8). Although our data showed the largest increase in protein carbonyls in the l-NAME + AngII group, mitochondrial ROS production was equally increased in AngII and l-NAME + AngII groups. One likely explanation would be that cytoplasmic ROS production by l-NAME-impaired NOS contributed to the tissue protein carbonyls (4), whereas mitochondrial ROS was induced by AngII only (8). Although the decrease in mitochondrial OXPHOS function in this model is consistent with most other animal models of HF, it is different from our observations in human HF (5, 19). In human HF, although metabolic pathways upstream of the mitochondria are impaired (19), there is preserved mitochondrial OXPHOS capacity, as measured ex vivo (5). It is likely that HF in humans develops over many years unlike the acute HF in these animal models. Nevertheless, energy depletion attributable to reduced mitochondria numbers and/or inefficient substrate metabolism is an important characteristic of HF in humans and animal models.
Our HF mouse model exhibited edema, as measured by increased free water content. However, body weight and percentage of body fat were lower than that of the controls. Lean body mass, however, remained unchanged. This might be due to decrease in food intake, from poor blood perfusion, peripheral lipolysis, and reduced energy levels in HF. Furthermore, lean mass typically reduces in patients with HF (3). This may be developing over the years in chronic human HF and is not seen in this mouse model of acute HF. Pulse rates typically increase to compensate for reduced cardiac output; however, l-NAME may reduce the heart rate, as seen by reversal of effects of β-adrenergic stimulants, such as isoproterenol by l-NAME (31). l-NAME has been shown to decrease heart rate in dogs (14) and rats (43). It may be a combined result of impaired baroreceptor reflex response and elevated vagal parasympathetic tone evoked by the rise in blood pressure (25, 36). Interestingly, isoflurane suppressed heart rates of mice in all four groups. This anesthetic agent has been documented to initially raise then reduce heart rates to the range that we report and enable the most stable echocardiography readings for FS and end-diastolic dimensions compared with other anesthetics (34).
By determining the effects of l-NAME and AngII individually in our study, we found that each compound was exclusively responsible for certain aspects of the HF phenotype. Indeed, l-NAME suppresses pulse rate in this mouse model when subjects are awake, an effect that is exclusive to l-NAME-treated groups. On the other hand, AngII has an exclusive effect on fibrosis, whereas l-NAME has virtually no impact. Microarray analyses identified few gene changes exclusively associated with l-NAME, whereas 313 gene changes were exclusively contributed by AngII. Although l-NAME had a smaller exclusive impact, it was crucial for the development of HF because a large number of gene changes occurred in the combination treatment that were not seen with the individual drugs. This is representative of human nonischemic HF in which NO inhibition and hypertension are two key risk factors.
IPA analyses served to robustly distinguish the effect of the combination treatment of l-NAME + AngII. In contrast to the recent report by Lai et al. (23) on ischemic HF, we found a large number of gene changes in oxidative phosphorylation, but, in agreement with Lai et al., we found few gene changes in the TCA pathway in the l-NAME + AngII group. In this model, the microarray data suggest that cellular remodeling pathways, which included hypertrophy signaling, cell proliferation, TGF-β signaling pathways, and metabolic pathways that included OXPHOS, fatty acid oxidation, and mitochondrial function, determined the deterioration of metabolic and cardiac contractile function in the l-NAME + AngII group.
Our study, however, is not without limitations. One limitation of the measurements performed under isoflurane anesthesia is that l-NAME reduces the level of minimum alveolar concentration in rodents, thereby potentially influencing our echocardiography read outs. Therefore, although several measurements that were independent of isoflurane treatment, including gene expression, oxidative stress, mitochondrial function, cardiomyocyte size, and microarray data, show more distinct differences in groups, we do not see large differences in FS or EF, which were done while subjects were under anesthesia. Metabolic alterations that have been shown to change early and progressively lead to systolic dysfunction (27, 41); however, the longer duration of treatment in our study may lead to better distinction of cardiac contractile function among the four groups. Additionally, diastolic pressure assessed with tail cuff measurements has its limitations in accuracy compared with invasive blood pressure assessments. Finally, AngII and l-NAME were administered systemically and may have effects on several organ systems, including the nervous system, that cumulatively affect the cardiac remodeling.
In conclusion, using mitochondrial, metabolic, contractile, and microarray measurements, we show that a 5-wk treatment of l-NAME combined with AngII results in severe nonischemic HF in mice. This model emphasizes the role of mitochondria in nonischemic HF, primarily from respiratory impairment of SSM and mitochondrial oxidative stress mediated by downregulation of important mitochondrial pathways.
This study was supported by the American Heart Association Award (12POST9020018) to A. Gupte and generous support from Methodist DeBakey Heart & Vascular Center Texans Grant, Houston Methodist Foundation grants from Elaine and Mary Finger, Patrick Studdert, Charif Souki, and Stedman-West Foundation to D. Hamilton.
No conflicts of interest, financial or otherwise, are declared by the authors.
Author contributions: D.J.H., A.Z., A.M.C.-R., K.A.Y., G.T.-A., and A.A.G. conception and design of research; D.J.H., A.Z., S.L., and A.A.G. analyzed data; D.J.H., A.Z., S.L., K.A.Y., and A.A.G. interpreted results of experiments; D.J.H. and A.A.G. drafted manuscript; D.J.H., A.Z., S.L., G.T.-A., and A.A.G. edited and revised manuscript; D.J.H., A.Z., S.L., T.N.C., K.A.Y., G.T.-A., and A.A.G. approved final version of manuscript; A.Z., S.L., T.N.C., J.A.S., I.V., A.M.C.-R., and A.A.G. performed experiments; A.Z., S.L., and A.A.G. prepared figures.
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