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Am J Physiol Heart Circ Physiol 293: H93-H102, 2007. First published March 16, 2007; doi:10.1152/ajpheart.00037.2007
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Role of oxidative stress in cardiac dysfunction of PPAR{alpha}–/– mice

Aziz Guellich,1 Thibaud Damy,1,2 Yves Lecarpentier,1,3 Marc Conti,4 Victor Claes,6 Jane-Lyse Samuel,1 Jeanine Quillard,5 Jean-Louis Hébert,3 Thierry Pineau,7 and Catherine Coirault1

1Institut National de la Santé et de la Recherche Médicale (INSERM) U689, Centre de Recherche Cardiovasculaire INSERM Lariboisière, Paris, France; Université Paris VII, Paris, France; 2INSERM U660; 4Faculté de Médecine Paris XII, Créteil Cedex France, 3Services d'Explorations Cardio-Respiratoires; 4de Biochimie et 5d'Anatomie-Pathologie, Hôpital de Bicêtre, Assistance Publique-Hôpitaux de Paris; Université Paris XI, Le Kremlin-Bicêtre, France; 6University of Antwerp, Centrum Technologie voor Gehandicapte Persone, Antwerp, Belgium; and 7Laboratoire de Pharmacologie et Toxicologie, Institut National de la Recherche Agronomique, Toulouse, France

Submitted 10 January 2007 ; accepted in final form 15 March 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This study was designed to determine the effects of PPAR{alpha} lack on cardiac mechanical performance and to identify potential intracellular mechanisms linking PPAR{alpha} pathway deficiency to cardiac contractile dysfunction. Echocardiography, ex vivo papillary muscle assays, and in vitro motility assays were used to assess global, intrinsic ventricular muscle performance and myosin mechanical properties, respectively, in PPAR{alpha}–/– and age-matched wild-type mice. Three-nitrotyrosine formation and 4-hydroxy-2-nonenal protein-adducts, both markers of oxidative damage, were analyzed by Western blot analysis and immunolabeling. Radical scavenging capacity was analyzed by measuring protein levels and/or activities of the main antioxidant enzymes, including catalase, glutathione peroxidase, and manganese and copper-zinc superoxide dismutases. Echocardiographic left ventricular fractional shortening in PPAR{alpha}–/– was 16% lower than that in wild-type. Ex vivo left ventricular papillary muscle exhibited reduced shortening velocity and isometric tension (three- and twofold, respectively). In vitro myosin-based velocity was {approx}20% slower in PPAR{alpha}–/–, indicating that myosin itself was involved in the contractile dysfunction. Staining of 3-nitrotyrosine was more pronounced in PPAR{alpha}–/–, and myosin heavy chain was the main nitrated protein. Formation of 3-nitrotyrosine myosin heavy chain was twofold higher in PPAR{alpha}–/– and 4-hydroxy-2-nonenal protein-adducts were threefold higher. The expression and activity of manganese superoxide dismutase were respectively 33% and 50% lower in PPAR{alpha}–/–, with no changes in copper-zinc superoxide dismutase, catalase, or glutathione peroxidase. These findings demonstrate that PPAR{alpha} pathway deficiency impairs cardiac function and also identify oxidative damage to myosin as a link between PPAR{alpha} deficiency and contractile dysfunction.

contractility; metabolism; myosin; cardiac failure

THE PEROXISOME PROLIFERATOR-ACTIVATED RECEPTOR (PPAR-{alpha}, -beta/{delta}, and -{gamma}) family of lipid-activated nuclear receptors plays a critical role in the gene regulation of cellular lipid metabolism (10). In cardiac muscle, PPAR{alpha} is expressed at relatively abundant levels and activates numerous genes involved in cellular fatty acid uptake and oxidation (10). Interestingly, PPAR{alpha} gene regulatory pathway activity is downregulated in the hypertrophied heart (2) and in the human failing heart (2, 40, 43). It is unclear whether this metabolic shift, with increased reliance on glucose metabolism rather than fatty acid oxidation, is a protective response allowing the heart to maintain contractile function or an initial step leading to progressive deterioration of contractile function (2, 33). The functional and biological roles of PPAR{alpha} in cardiac muscle have been widely investigated through the PPAR{alpha}–/– mouse model (5, 11, 47). Although PPAR{alpha}–/– mice have a normal life span, they develop progressive cardiac fibrosis and myofibrillar fragmentation associated with abnormal mitochondrial ultrastructure (47). Reduced cardiac contractile performance at baseline (26) or in response to increased workload (27) has been recently reported in PPAR{alpha}–/– mice. However, the precise consequences of chronic PPAR{alpha} deficiency on the cardiac contractile apparatus and muscle performance remain to be established.

There is increasing evidence that a proper balance between oxidants and antioxidant defenses is required to maintain normal cardiac function (3, 12, 30, 39). Depletion in antioxidant enzymes such as superoxide dismutase (SOD), catalase, and glutathione peroxidase (GPX) (39) or overproduction of reactive oxygen species (ROS) or reactive nitrogen species (RNS) may induce oxidative stress and cause cardiac functional disorders (12, 39). Recently, increased PPAR{alpha} activity and fatty acid oxidation have been associated with an increase in reactive oxygen intermediates (42). However, there is also strong evidence that activation of PPAR{alpha} is necessary to prevent cellular oxidative damage that may occur during physiological cellular metabolism or under conditions of inflammation and oxidative stress, probably through repressing nuclear factor-{kappa}B signaling and reducing inflammatory cytokine production (38, 44, 45). Therefore, it is possible that chronic deactivation of the PPAR{alpha} signaling pathway upsets normal equilibrium between oxidant production and antioxidant defenses and contributes to cardiac damage.

The first aim of our study was to identify the effects of chronic deficiency in the PPAR{alpha} pathway on cardiac dysfunction. To this end, echocardiography, in vitro papillary muscle mechanics, and motility assays with purified myosin molecules were performed in PPAR{alpha}–/– mice. We then aimed to identify the mechanisms involved in cardiac dysfunction. We tested the hypothesis that oxidative damage occurred in PPAR{alpha}–/– hearts by using markers of protein oxidation (4-hydroxy-2-nonenal protein adducts, HNE) and a marker of peroxynitrite and/or nitrosative stress (3-nitrotyrosine formation, 3-NT) (35). Finally, the cardiac levels and activities of several intracellular antioxidant enzymes were analyzed to assess the radical scavenging capacity in PPAR{alpha}–/– hearts. To this end, protein levels and/or activities of the H2O2 scavenging enzymes catalase and GPX and the two intracellular superoxide dismutating enzyme copper-zinc (Cu/Zn-SOD, cytosolic) and manganese superoxide dismutase (Mn-SOD, mitochondrial) were examined.

We found that the lack of PPAR{alpha} was primarily responsible for the development of intrinsic cardiac dysfunction associated with mechanical impairment of myosin, and we identified oxidative damage to myosin as a link between PPAR{alpha} deficiency and cardiac contractile dysfunction.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. Experiments were performed on male PPAR{alpha} wild-type (WT) and null mice (PPAR{alpha}–/–) mice (23) at 9 mo of age. WT C57 BL/6J mice were obtained from Janvier (Le Genest Saint Isle, France). PPAR{alpha}-deficient males on a C57 BL/6J genetic background were bred at the Institut National de la Recherche Agronomique transgenic animal facility in accordance with European Union guidelines for animal care. The PPAR{alpha} null mouse was chosen because it has proven to be an appropriate model to link the loss of PPAR{alpha} to cardiac dysfunction (5, 11, 47). Nine-month-old PPAR{alpha}–/– mice were chosen because they are expected to exhibit moderate global cardiac dysfunction (26, 47). The investigation conformed to the National Institutes of Health Guide for Care and Use of Laboratory Animals and was allowed by the Animal Ethics Committee of the Institut National de la Sauté et de la Recherche Médicale. Mice were fed standard rodent chow and had free access to water.

Echocardiography. Echocardiographic measurements including two-dimensional, M-mode, and tissular Doppler imaging were performed using a Vivid 7 machine (General Electric), equipped with 8- and 12-MHz linear probes. Measurements were performed under light isoflurane anesthesia on three consecutive cardiac cycles, blind to the genotype. Internal end-systolic (ESLVD) and end-diastolic (EDLVD) left ventricular (LV) diameters were determined, and the percentage of LV fractional shortening was calculated as [(EDLVD – ESLVD)/ EDLVD] x 100. Other calculations were performed as previously described (28, 37).

Electrocardiography recording. Long-term telemetric ambulatory ECGs were recorded by using implantable transmitters (ETA-F20, Data Sciences International). The transmitters were inserted subcutaneously in the abdominal region under isoflurane anesthesia. Telemetric ECG tracings were obtained in conscious mice over 24 h. RR interval analysis was performed by ECG Auto software (EMKA Technologies, Paris, France).

Blood pressure measurement. Noninvasive determinations of systolic blood pressure (SBP) were made in conscious mice using a tail-cuff system (Visitech BP-2000 Systems, Apex, NC). Ten consecutive measurements were performed after a training period. Artifactual and aberrant tracings due to movements of the mice were discarded, and a mean value was generated for each individual mouse.

Antibodies. Rabbit polyclonal anti-MnSOD and anti-Cu/Zn antibodies were purchased from Stressgen. Rabbit polyclonal anti-catalase and anti-HNE antibodies were from Calbiochem, and mouse monoclonal anti-3-NT antibody was purchased from Santa Cruz. Mouse monoclonal anti-{alpha}-actinin, anti-myosin, anti-vinculin antibodies, and rabbit polyclonal anti-actin antibody were purchased from Sigma. Secondary anti-rabbit IgG FITC-conjugated, anti-mouse, and anti-rabbit horseradish peroxidase-conjugated antibodies were from Amersham. Biotinylated anti-mouse IgG antibody was from Vector, and anti-mouse IgG Alexa Fluor 488 was purchased from Molecular Probes.

Histology and immunofluorescence analysis. Histology was performed on ventricular formalin-fixed tissue embedded in paraffin. Sections of 5-µm thickness were stained by hematoxylin-eosin-saffron and Masson trichrome. Additional hearts were snap frozen in liquid nitrogen and stained with Oil Red O or used for immunofluorescence analysis. LV cryosections of 5-µm width were air-dried, fixed in 4% formalin for 15 min, and saturated with 5% bovine serum albumin (BSA) in phosphate-buffered saline (PBS) for 30 min. Sections were incubated for 90 min at room temperature with primary antibodies (anti-vinculin, dilution 1:250; anti-{alpha}-actinin, dilution 1:200; anti-MnSOD, dilution 1:100; and anti-3-NT, dilution 1:50) diluted in PBS-2% BSA. After three rinses in PBS, the slides were incubated at room temperature in the dark for 30 min with secondary antibodies diluted in PBS (FITC-conjugated anti-rabbit IgG, dilution 1:50; biotinylated anti-mouse IgG dilution 1:50; or anti-mouse IgG Alexa Fluor 488, dilution 1:250). Incubation with the biotinylated antibody was followed by application of streptavidin fluorescein (Amersham, 1/50). Slides incubated with phalloidin Fluoprobes 547 (Interchim, 2.5 UI/ml) were postfixed in methanol for 5 min at –20°C. Slides were mounted in Vectashield medium (Vector). Sections exposed only to secondary antibodies were used as negative controls and showed very low background staining. Fluorescence was visualized using a DMRBE Leica microscope equipped with a x40 oil epifluorescence objective. Morphometry was performed using IPlab software. A minimum of 50 myocytes/field and 5–8 fields/heart were measured.

Papillary muscle mechanics. Mechanical experiments were performed on LV papillary muscle. After anesthesia with pentobarbital, the heart was rapidly excised and placed in a physiological solution. The LV was opened under a dissection microscope, and one papillary muscle was carefully dissected. The muscle was then placed in a circulating organ bath containing modified Krebs-Henseleit buffer solution (22). The solution was bubbled with 95% O2-5% CO2 and maintained at 29°C and pH 7.4 to ensure good mechanical stability. One extremity of the papillary muscle was held by a stationary clip, and the other was maintained with another clip attached to an isotonic electromagnetic force-transducer lever. In brief, a precision current source delivers the current through the coil and hence determines the force at the tip of the lever. The displacement of the lever is measured with an opto-electronic transducer. The length range is 1,000 µm with a maximal error of 1%, and the maximum force range corresponds to a load of 20 mN at the lever tip, with a noise floor of 0.01 mN. The force transducer we used enabled us to analyze mechanical performance over the whole load continuum. After an equilibration period of 30 min, muscle was supramaximally stimulated using two platinum electrodes at the optimal force-frequency response, i.e., at a frequency of 6/min. Experiments were performed at the initial muscle length at which active isometric tension was maximum (Lmax). Muscle force was normalized per cross-sectional area (CSA) at the middle part of the muscle using the following approximation: CSA = 0.75 x width x depth (24). The mechanical parameters were calculated from three different loaded contractions: contraction 1 was loaded with preload only and abruptly clamped to zero load just after stimulus according to the zero-load clamp technique (4); contraction 2 was loaded with preload only; and contraction 3 was carried out against a heavy load that the muscle could not overcome, so its contraction was fully isometric. During the contraction phase, we recorded the maximum unloaded shortening velocity (Vmax, in Lmax/s) of contraction 1, the maximum extent of muscle shortening of contraction 2 ({Delta}L, in %Lmax), the peak isometric tension, i.e., peak force normalized per cross-sectional area (Pmax, in mN/mm2) of contraction 3, and the positive peak of isometric tension derivative of contraction 3 (+dP/dt, mN·mm–2·s–1).

In vitro motility assays. Purified myosin obtained from LV was assessed by gel electrophoresis (19). F-actin was prepared from rabbit skeletal muscle by standard methods (36) and fluorescently labeled with phalloidin FluoProbes 547 (Interchim). Motility assays were carried out at 29°C as previously described (7). The movement of actin filaments was observed under a Zeiss epifluorescence microscope (Axiovert 200, 100/1.30 lens, Jena, Germany) equipped with an intensified camera (Hamamatsu C 2400, Hamamatsu City, Japan) and recorded on videotape. The mean velocities of each filament were analyzed using N. J. Carter's freeware RETRAC program.

Western blot analysis. Western blot analysis was performed on total protein extracts, and myosin fractions were obtained from LV tissues. Protein concentrations were determined using the BCA protein assay kit (Pierce) according to the manufacturer's instructions. The samples were stored at –20°C until use. Proteins were separated by 9% or 12% SDS-PAGE. Western blot analysis was performed using anti-MnSOD (1:1,000), anti-Cu/ZnSOD (1:1,000), anti-catalase (1:1,000), anti-HNE (1:1,000), and anti-3-NT (1:500) antibodies. Detection was performed using anti-mouse or anti-rabbit horseradish peroxidase-labeled antibodies (1/5,000). Membranes were revealed with ECL chemiluminescent substrate (Amersham). Light emission was detected with a highly sensitive imaging system (Fujifilm LAS-3000). Signals were quantified using Image Gauge software and normalized to the expression of actin or myosin.

Myosin isoform composition. Myosin isoform composition was determined on purified myosin stored in 50% glycerol. V1 and V3 myosin were separated on 8% polyacrylamide gels containing 10% glycerol (41). SDS-PAGE was performed in a Bio-Rad Mini-Protean II Dual Slab Cell electrophoresis system for 16 h at 4°C and 70 V. Gels were stained with Coomassie blue and quantification of V1 and V3 was performed using Image Gauge software.

Antioxidant enzymatic activities. To measure the activities of Cu/Zn-SOD, Mn-SOD, catalase, and GPX, frozen heart tissues were homogenized in 10 mM phosphate buffer, pH 7.8, EDTA 1 mM (8). The activities of SOD were measured using xanthine-xanthine oxidase to generate superoxide radicals, which react with 2-(4-iodophenyl)-3-(4-nitrophenol)-5-phenyltetrazolium chloride (INT) to form a red formazan dye with maximum absorbance at 550 nm. SOD activity was then measured by the degree of inhibition of this reaction. One unit of SOD was defined as the amount of enzyme causing a 50% inhibition of the rate of reduction of INT (Randox Laboratories). Total SOD activity was measured at pH 7.8, and Cu/Zn SOD activity, primarily located in the cytosol, was measured at pH 10.2. The activity of MnSOD, exclusively located in the mitochondria matrix, was obtained by subtracting Cu/Zn SOD activity from total SOD activity (29). GPX activity was determined using an indirect, coupled test procedure with terbutyl peroxide as a substrate (34). Catalase activity was measured according to Johansson and Borg's method (17). To minimize within-run variations, each sample was measured in duplicate. Activities are expressed in units normalized per milligram of protein (U/mg).

Statistics. Data are expressed as means ± SE. Comparisons were made using Student's unpaired t-test. Differences were considered significant at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Cardiac morphometry and structure. Body weight was about 20% higher in PPAR{alpha}–/– than that in WT mice (32.6 ± 0.7 vs. 40.6 ± 1.2 g, P < 0.05). The heart weight-to-body weight ratio did not differ between WT and PPAR{alpha}–/– mice (4.3 ± 0.1 and 4.0 ± 0.1 mg/g in WT and PPAR{alpha}–/–, respectively). Morphological data are presented in Fig. 1, with n being the number of animals. Gross examination of the heart of PPAR{alpha}–/– mice did not reveal abnormalities (Fig. 1, A and B). Morphometric analysis was performed on longitudinal (Fig. 1, C and D) and transversal (Fig. 1, E and F) sections. When compared with WT, LV knockout myocytes were wider (23.4 ± 1.6 vs. 14.6 ± 0.4 µm, P < 0.01) (Fig. 1, C and D). Although myocyte length did not change between groups (93.7 ± 5.4 vs. 112.3 ± 9.5 µm in WT and knockout mice, respectively, not significant), this resulted in a significantly greater calculated cell area (cell length x width) in PPAR{alpha}–/– compared with WT mice (+94%, P < 0.05). Immunofluorescence performed with antibody directed against {alpha}-actinin and actin labeling with phalloidin showed regular striation patterns in both groups. In addition, Oil Red O staining did not reveal neutral lipid accumulation within cardiomyocytes (data non shown).


Figure 1
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  		Fig. 1. Structure of ventricular myocardium in wild-type (WT) and peroxisome proliferator-activated receptor (PPAR{alpha}–/–) mice. (A, B). Light microscopy of 5-µm ventricular sections stained with hematoxylin and eosin saffron (x100). Double labeling with anti-vincilin antibody (in green) and phalloidin (in red) in longitudinal (C, D) and transversal (E, F) left ventricular (LV) sections (bar =10 µm). Nuclei appear in blue. Vinculin labeling of the sarcolemma allows measurement of cardiomyocyte width and length (white lines in C and D). Phalloidin labeled actin filaments within cardiomyocytes. When compared with WT, PPAR{alpha}–/– hearts exhibited cardiomyocyte hypertrophy with preserved striation pattern. n = 4/group in WT and PPAR{alpha}–/–.

 
In vivo assessment of cardiac function. Typical transthoracic echocardiograms are presented in Fig. 2, A and B. PPAR{alpha}–/– mice showed a significant decrease in in situ contractile performance, as attested by a 16% decrease in LV fractional shortening (P < 0.05) (Table 1). The early-to-late diastolic transmitral Doppler flow velocity (E/A) ratio, E wave deceleration times and E/Ea ratio did not differ between groups, thus suggesting preserved diastolic function in PPAR{alpha}–/– (Table 1). SBP was significantly lower in PPAR{alpha}–/– than that in WT mice (94.4 ± 3.1 vs. 104.5 ± 4.0 mmHg in PPAR{alpha}–/– and WT mice, respectively, P < 0.005; Fig. 2C). Heart rate at baseline did not differ between groups (Fig. 2D), and telemetric ambulatory ECG measurements did not evidence conduction or excitability defects.


Figure 2
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  		Fig. 2. Cardiac function in vivo. A and B: representative M-mode echocardiograms of WT and PPAR{alpha}–/– mice. PPAR{alpha}–/– mice exhibited a decrease in the LV fractional shortening percentage compared with control mice without significant changes in cardiac geometry. Systolic blood pressure (SBP, C) and heart rate (bpm, beats/min) in baseline conditions (D) in WT and PPAR{alpha}–/– mice. Values are means ± SE; n = 6 mice in WT and PPAR{alpha}–/–. *P < 0.005.

 

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  		Table 1. Echocardiographic characteristics

 
Intrinsic mechanical performance of in vitro LV papillary muscle. Intrinsic mechanical parameters were recorded from three different loaded contractions (Fig. 3A). No difference between groups was observed in the morphometric characteristics of the LV papillary muscles (Table 2). Vmax and {Delta}L were respectively 75% and 50% lower in PPAR{alpha}–/– than those in WT mice (Fig. 3, B and C), thus attesting to severe depression of shortening capacities in PPAR{alpha}–/– mice. In addition, peak isometric tension and +dP/dt were respectively 50% and 80% lower in PPAR{alpha}–/– than those in WT mice (Fig. 3, D and E).


Figure 3
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  		Fig. 3. Intrinsic LV muscle contractility. A: typical traces of length, velocity, and tension as a function of time were recorded in 3 different loading conditions: contraction 1 was loaded with preload only and abruptly clamped to zero load (4); contraction 2 was loaded with preload only; and contraction 3 was fully isometric. Mean mechanical parameters including maximum shortening velocity at zero load (Vmax) (B), maximum extent of shortening at preload ({Delta}L) (C), total isometric tension (D), and positive peak of tension derivative (+dP/dt) (E) were markedly reduced in PPAR{alpha}–/– when compared with WT. Values are means ± SE; n = 8 in WT and n = 10 in PPAR{alpha}–/– groups. *P < 0.05.

 

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  		Table 2. LV papillary muscle characteristics in WT and PPAR{alpha}–/–

 
Myosin molecular properties. In vitro motility assays were performed using purified LV myosin monomers. As shown in Fig. 4A, the mean velocity of actin filaments driven by myosin extracted from PPAR{alpha}–/– mice was 16% slower than that of the control rate (P < 0.01). Given that only myosin and not actin differed between groups under our experimental conditions, these data indicated that intrinsic alterations in myosin mechanical function occurred in LV from PPAR{alpha}–/– mice.


Figure 4
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  		Fig. 4. Myosin mechanical function and isoforms. A: distribution of velocities of actin filament sliding over cardiac myosins was shifted toward slower values in PPAR{alpha}–/– mice compared with those of WT mice. Individual frame-to-frame velocities from tracked filaments are plotted in a frequency histogram; n = 150 filaments in each group. B: relative proportion of {alpha}- and beta-myosin heavy chain (MHC) isoforms did not differ between WT and PPAR{alpha}–/– (n = 5/group). Inset: representative electrophoresis separation of {alpha}- and beta-MHCs in LV from WT and PPAR{alpha}–/–. LV with known {alpha}- and beta-MHCs was used to ascertain adequate separations of {alpha}- and beta-MHCs (lane 1).

 
Myosin isoforms. To determine whether reduced shortening capacities in PPAR{alpha}–/– were related to {alpha}- to beta-myosin isoform shift, myosin heavy chain (MHC) expression patterns were determined by electrophoresis (Fig. 4B). The amount of {alpha}-MHC and beta-MHC represented 86 ± 3% and 14 ± 3%, respectively, of total MHC isoform in WT. No significant shift in the {alpha}- to beta-MHC ratio was present in LV from PPAR{alpha}–/– mice (Fig. 4B).

Oxidative damage to contractile proteins. To determine whether oxidative damage to contractile proteins and particularly myosin contributed to reduced contractility in PPAR{alpha}–/–, we next examined protein tyrosine nitration and HNE-adduct formation by using both tissue samples and protein extracts. Immunohistochemistry studies revealed pronounced cytosolic and membrane-associated 3-NT staining in PPAR{alpha}–/– but not in control cardiomyocytes (Fig. 5, A and B). Importantly, in both myosin and total protein extracts, MHC was identified by Western blot analysis as the main nitrated protein and analysis revealed a twofold higher 3-NT-MHC in PPAR{alpha}–/– compared with that in WT mice (Fig. 6A, P < 0.01). In control and PPAR{alpha}–/– hearts, enhanced chemiluminescence revealed additional 3-NT bands of lower molecular masses ({approx}60, 47, and 43 kDa, respectively) but whose quantification did not differ between groups. HNE protein adduct, a marker of lipid peroxidation, was detectable in control and PPAR{alpha}–/– myocardium, the main HNE-conjugated band being at 75 kDa (Fig. 6B). The amount of this 75-kDa band was nearly threefold higher in PPAR{alpha}–/– than that in WT mice (P < 0.01). No evidence of myosin modification by HNE was detected in myosin extracts.


Figure 5
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  		Fig. 5. Typical examples of immunofluorescent stainings. Fixed LV sections were stained with antibodies against 3-nitrotyrosine (3-NT) (A and B) and MnSOD (C and D) in WT and PPAR{alpha}–/– (bar = 20 µm). Nuclei appear in blue. (n = 3/group).

 

Figure 6
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  		Fig. 6. Oxidative damage to sarcomeric proteins. Increased nitration of myosin heavy chain (3-NT MHC) (A) and increased 4-hydroxy-2-nonenal (HNE)-adducts of total protein extract (B) were observed in the PPAR{alpha}–/– heart. Quantifications are normalized to the expression of myosin (3-NT myosin) or actin (HNE-adducts) and reported in arbitrary units (a.u.). Values are means ± SE; n = 5–6/group. *P < 0.05.

 
Antioxidant enzyme protein expression. As shown in Fig. 5, C and D, LV myocytes of PPAR{alpha}–/– mice exhibited reduced MnSOD staining compared with WT. Quantification of MnSOD, Cu/ZnSOD, and catalase protein expressions are illustrated in Fig. 7. The amount of MnSOD was nearly 30% lower in PPAR{alpha}–/– mice than that in WT mice (P < 0.01) (Fig. 7A). In contrast, there were no significant differences in the protein expression of Cu/ZnSOD and catalase between PPAR{alpha}–/– and WT mice (Fig. 7, B and C).


Figure 7
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  		Fig. 7. Protein expression and/or activities of MnSOD (A), Cu/ZnSOD (B), catalase (C), and glutathione peroxidase (GPX) (D). Representative immunoblots obtained with specific antibodies directed against MnSOD (A), Cu/ZnSOD (B), and catalase (C). Protein quantifications are normalized to the expression of actin (bottom of each blot). Values are means ± SE; n = 5–6/group. *P < 0.05.

 
Antioxidant enzyme activities. Antioxidant enzyme activities are illustrated in Fig. 7. PPAR{alpha}–/– hearts displayed a 50% reduction in MnSOD activity compared with WT (P < 0.01, Fig. 7A). There were no significant differences in Cu/ZnSOD (Fig. 7B), catalase (Fig. 7C), and GPX (Fig. 7D) activities between WT and PPAR{alpha}–/– mice.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Our study demonstrated an important role of PPAR{alpha} in maintaining cardiac contractile performance and oxidative homeostasis. We found that cardiac PPAR{alpha} deficiency actually led to myosin dysfunction with a pronounced decrease in cardiac MnSOD expression and activity and an increase in oxidative damage. Furthermore, we identified myosin as a target of oxidative stress that may account for intrinsic myocardial contractile dysfunction in PPAR{alpha}–/– mice.

Functional consequences of cardiac PPAR{alpha} deficiency. The biological roles of PPAR{alpha} in cardiac muscle have been widely investigated through the PPAR{alpha}–/– mouse model. These roles include regulation of mitochondrial fatty acid (FA)-oxidation, FA uptake, and lipoprotein assembly and transport (10, 11, 20, 47). More recently, the functional significance of PPAR{alpha} deficiency has also been determined by analyzing cardiac substrate metabolism, myocardial energetics, and contractile function in PPAR{alpha}–/– hearts (26, 27). Thus the PPAR{alpha} null mouse has proven to be an appropriate model to demonstrate the link between lack of PPAR{alpha} and cardiac dysfunction. Furthermore, to date, a model of cardiac-restricted deletion of PPAR{alpha} has not been developed. Precise functional cardiac phenotyping of PPAR{alpha}–/– mice could reveal unsuspected physiological mechanisms of direct relevance to cardiac muscle diseases.

Our first aim was to identify the effects of chronic PPAR{alpha} deficiency on intrinsic cardiac muscle contractility. Echocardiography data revealed reduced systolic performance without ventricular dilatation, which is consistent with recently reported results in PPAR{alpha}–/– mice (26). However, histological investigations revealed significant cardiomyocyte hypertrophy in PPAR{alpha}–/– mice (Fig. 1). Given that the heart weight-to-body weight ratio did not differ between WT and PPAR{alpha}–/– mice, these data indicated a loss of cardiomyocytes in PPAR{alpha}–/– compared with WT, i.e., that apoptosis and/or fibrosis occurred in hearts from PPAR{alpha}–/–. Accordingly, we found that intrinsic papillary muscle performance was depressed in PPAR{alpha}–/– mice (Fig. 3). Several mechanisms could help explain why this occurred, despite a moderate reduction in fractional shortening on echocardiography. First, reduced LV fractional shortening in PPAR{alpha}–/– was not related to an increase in afterload because SBP was lower in PPAR{alpha}–/– mice than that in WT (Fig. 2C). Given that reduction in SBP is expected to induce an increase in LV fractional, in vivo cardiac contractility in PPAR{alpha}–/– mice could be more severely impaired than that evaluated by echocardiography. Second, the depression of intrinsic papillary muscle performance may involve a low rate of beta-oxidation and ATP production induced by the deletion of PPAR{alpha} (27, 47). These findings attest to energetic deprivation in the PPAR{alpha}–/– heart (27, 47), of which functional consequences may differ between in vivo and in vitro conditions. Indeed, it has been shown that the use of FAs in the perfusion medium prevents contractile abnormalities of isolated working heart preparation (26). In contrast, the ex vivo cardiac performance of PPAR{alpha}–/– mice was reduced when the perfusion medium contained only glucose as a substrate (26), i.e., in conditions where muscle energy was produced by glycolysis and glucose oxidation. In our isolated papillary muscle experiments, the bathing medium contained only glucose as a substrate. Although such bathing conditions are optimal in papillary muscle from WT mice, they could contribute to reduced intrinsic mechanical performance in PPAR{alpha}–/–. In addition, in our isolated papillary muscle experiments, the bathing medium was continuously bubbled with 95% O2-5% CO2. This results in a high oxygen partial pressure that may increase the production of ROS and oxidative stress within cardiomyocytes, and in turn could contribute to decreased in vitro muscle mechanical performance in PPAR{alpha}–/–. Finally, it is widely recognized that functional impairment does not necessarily correlate between in vivo and in vitro conditions, especially in mice, and that experimental in vitro conditions could unmask and exacerbate even moderate intrinsic dysfunction (16, 18).

We then explored the mechanical function of myosin, the molecular motor of the heart that generates force and motion by coupling its ATPase activity to its cyclic interaction with actin. We found that the in vitro capacity of myosin to propel actin filaments was reduced in PPAR{alpha}–/– heart (Fig. 4A), thereby strongly suggesting that pathological processes affecting the myosin molecule itself were involved in cardiac dysfunction in PPAR{alpha}–/– mice (7, 14, 19). We therefore sought to identify intracellular pathways and protein targets linking PPAR{alpha} deficiency to cardiac contractile dysfunction. Because {alpha}- and beta-MHC, the two MHC isoforms expressed in mammalian heart, have profound functional differences in terms of shortening velocity, force generation, and ATPase activity (14, 21), our first step was to analyze the relative expression of {alpha}- and beta-MHC in PPAR{alpha}–/–. In adult mice, {alpha}-MHC expression normally predominates in the ventricles (31), but a cardiac MHC isoform shift can be achieved by pleiotropic stimuli, such as hypertrophy or changes in hormonal status (15, 21, 31). No changes in MHC isoform content were observed in PPAR{alpha}–/– mice, so that changes in heart function cannot be ascribed to modifications in MHC isoform content.

Oxidative damage and PPAR{alpha} deficiency. PPAR{alpha} has an important role in the control of various types of inflammatory response (9). In noncardiac tissues, there is strong evidence that normal PPAR{alpha} function is necessary to protect cells from oxidative damage mediated by superoxide radicals generated during normal cellular metabolism or under conditions of inflammation and oxidative stress (38). Spleen and liver from PPAR{alpha}–/– mice express indicators of oxidative stress much earlier in their lifespan than WT mice, and PPAR{alpha} agonists help to prevent oxidative damage in aged WT mice (38). In the heart, myofibrillar (6, 30) as well as mitochondrial (6, 32) proteins are major targets of oxidative stress-derived effects, and oxidative and/or nitrosative changes in proteins have been shown to modulate cardiac function (3, 39). Therefore, we hypothesized that oxidative and/or nitrosative modification of contractile proteins contribute to cardiac dysfunction in PPAR{alpha}–/– mice. Among the different modifications induced by peroxynitrite and other RNS, that of protein tyrosine-nitration is one of the best characterized in several cardiomyopathies (3, 35, 39). Notably, we found that MHC was one of the major targets of protein tyrosine nitration in PPAR{alpha}–/– hearts (Fig. 6). Given that in vitro tyrosine nitration of myosin reduces sliding velocity (personal data), it is entirely conceivable that alterations caused in the conformation of myosin by its nitration account for the reduced myosin-based velocity observed in PPAR{alpha}–/– hearts. The other nitrated bands detected in WT and PPAR{alpha}–/– hearts had apparent molecular masses of {approx}58, 47, and 43 kDa, respectively. Given that quantification of these other lower molecular mass bands did not reveal a significant difference between groups, their identification was not systematically performed, except for the 43-kDa band, which was identified as actin by Western blot analysis. Mihm et al. (Ref 30) previously reported increased creatine kinase (CK) nitration in an experimental model of heart failure. Importantly, this previous study analyzes a specific protein fraction, namely, myofibrillar CK extracts obtained after immunoprecipitation of myofibrillar isolates, whereas total protein and myosin-enriched fractions were analyzed in our study. The molecular mass of CK is about 40 kDa, i.e., in the close range of the 43-kDa nitrated band found in our study and identified as actin. Thus we cannot exclude the possibility that actin nitration masked CK nitration and that increased CK nitration occurred in PPAR{alpha}–/– mice. In addition, PPAR{alpha}–/– hearts exhibited a substantial increase in HNE-protein adducts, thus attesting to increased lipid peroxidation, which may also contribute to contractile alterations (13). Taken as a whole, our results show that oxidative damage occurred in PPAR{alpha}–/– hearts and contributed, at least in part, to the depressed cardiac performance. Alternatively, peroxynitrite may also impair myocardial contractility via other mechanisms, including activation of matrix metalloproteinases, nuclear enzyme poly(ADP-ribose) polymerase, induction of apoptosis in myocytes, impairment of mitochondrial respiration, or abnormal calcium cycling (35). There are many possible sources of increased production of superoxide in PPAR{alpha}–/– hearts, including activation of several enzyme complexes such as NADPH oxidases (NADPHox) and xanthine oxidase. Increased myocardial nitric oxide (NO) may in turn result from different sources, including increased myocardial inducible NO synthase activity (35). Finally, it is important to remember that generation of peroxynitrite can trigger a wide range of cellular responses, from subtle posttranslational modifications of proteins such as S-glutathiolation or S-nitrosylation to severe oxidative injury leading to induction of cell death (35).

Protection against oxidative damage is accomplished by a complex defense system composed of antioxidant molecules and antioxidant enzymes that reduce the damaging effects of ROS by converting more reactive species to less reactive and less damaging forms (13). The SODs act as a first line of defense against oxygen free radical-mediated damage by catalyzing the dismutation of superoxide anions to H2O2. Subsequently, H2O2 is reduced to H2O and O2 by peroxidases (e.g., GPX) or catalase (13). Regulation of MnSOD is considered to play a crucial role in cardiac oxidative stress (25, 46). Our results clearly demonstrate a substantial decrease in MnSOD expression in PPAR{alpha}–/– hearts compared with WT hearts, associated with an even more pronounced decrease in enzymatic activity (Fig. 7), which would suggest enzyme inactivation (25, 46). These decreases were likely to increase the level of superoxide, which in turn may react with the NO to form the powerful oxidant peroxynitrite and induce protein nitration. Our data suggest that in addition to its metabolic regulatory effects, PPAR{alpha} was also involved in the maintenance of cardiac oxidant-antioxidant balance.

In conclusion, we demonstrated that lack of PPAR{alpha} was responsible for the development of reduced cardiac contractile function related at least in part to direct tyrosine-nitration of myosin and increased lipid peroxidation in the heart of PPAR{alpha}–/– mice. Oxidative and nitrosative stress appeared to be linked to profound alterations in MnSOD expression and activity. Our study thus provides further insight into the functional role of PPAR{alpha} in long-term regulation of cardiac muscle function and identifies oxidative damage to contractile proteins as a potential link between downregulation of the PPAR{alpha} pathway and the progression of cardiac dysfunction. Together, these data indicate that functional PPAR{alpha} is required to regulate cellular oxidant-antioxidant balance, prevent oxidative damage and preserve contractile function in cardiac muscle.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
This work was supported by the Fondation de France (Number 2005005345) and the Fédération Française de Cardiologie. A. Guellich is the recipient of a doctoral grant from the French Ministry of Education, Research and Technology.


   ACKNOWLEDGMENTS
 
We are thankful to Françoise Marotte and José Vilar for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. Coirault, CRCIL U689, 41 Bd de la Chapelle, 75475 Paris Cedex 10, France (e-mail: coirault{at}larib.inserm.fr)

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.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

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