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Cardiovascular Research 2006 69(2):440-449; doi:10.1016/j.cardiores.2005.10.019
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Copyright © 2005, European Society of Cardiology

Peroxisome proliferator-activated receptor {delta} (PPAR{delta}) activation protects H9c2 cardiomyoblasts from oxidative stress-induced apoptosis

Matthieu Pesanta, Stéphanie Sueura, Patrick Dutartreb,1, Mireille Tallandierb, Paul A. Grimaldic, Luc Rochettea and Jean-Louis Connata,*

aLaboratoire de Physiopathologie et Pharmacologie Cardiovasculaires Expérimentales, Biologie Animale Cellulaire et Moléculaire, Faculté des Sciences Gabriel, Université de Bourgogne, IFR Santé 100, 6 Bd Gabriel, 21000 Dijon, France
bLaboratoires Fournier Pharma, Biologie Exploratoire, 50 rue de Dijon, 21121 Daix, France
cCentre de Biochimie, INSERM U636, UFR Sciences, Parc Valrose, 06108-Nice cedex, France

* Corresponding author. Tel.: +33 380 39 62 16; fax: +33 380 39 38 25. Email address: jean-louis.connat{at}u-bourgogne.fr

Received 14 March 2005; revised 11 October 2005; accepted 12 October 2005


   Abstract
Top
 Abstract
1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Acknowledgments
 References
 
Objective: Activation of peroxisome proliferator-activated receptor {alpha} (PPAR{alpha}) and PPAR{gamma} plays beneficial roles in cardiovascular disorders such as atherosclerosis and heart reperfusion. Although PPAR{alpha} and {gamma} have been documented to reduce oxidative stress in the vasculature and the heart, the role of PPAR{delta} remains poorly studied.

Methods and results: We focused on PPAR{delta} function in the regulation of oxidative stress-induced apoptosis in the rat cardiomyoblast cell line H9c2. Using semi-quantitative reverse transcriptase-polymerase chain reaction (RT-PCR), we showed that PPAR{delta} is the predominantly expressed isotype whereas PPAR{alpha} was weakly detected. By performing cell viability assays, we also showed that the selective PPAR{delta} agonist GW501516 protected cells from H2O2-induced cell death. The protective effect of GW501516 was due to an inhibition of H2O2-triggered apoptosis as shown by annexin-V labeling, DNA fragmentation analysis, and caspase-3 activity measurement. We demonstrated by transient transfection of a dominant negative mutant of PPAR{delta} that the protection induced by GW501516 was totally dependent on PPAR{delta}. Semi-quantitative RT-PCR and Western blotting analysis demonstrated that GW501516 treatment upregulated catalase. Moreover, forced overexpression of catalase inhibited H2O2-triggered apoptosis, as evidenced by annexin-V labeling.

Conclusion: Taken together, our results account for an important role of PPAR{delta} in inhibiting the onset of oxidative stress-induced apoptosis in H9c2 cells. PPAR{delta} appears to be a new therapeutic target for the regulation of heart reperfusion-associated oxidative stress and stimulation of enzymatic antioxidative defences.

KEYWORDS Nuclear receptors; Hydrogen peroxide; Cell death; Catalase; PPAR{delta}; Dominant negative mutant; Peroxisome proliferator-activated receptor {delta}


   1. Introduction
Top
 Abstract
 1. Introduction
2. Methods
 3. Results
 4. Discussion
 Acknowledgments
 References
 
Cardiovascular diseases represent one of the main causes of death in Western countries. High-fat diet, a sedentary way of life and genetic factors seem to account for the development of cardiovascular diseases including atherosclerosis and heart stroke. It is also well known that reactive oxygen species (ROS), mediators of the oxidative stress, play deleterious effects on vasculature and myocardium [1–3]. ROS and hydrogen peroxide (H2O2), a by-product of oxidative stress, have been reported to induce apoptosis in various cell types including primary cultured newborn rat cardiomyocytes [4–6]. The same apoptotic effect of H2O2 exposure was reported for the embryonic rat heart-derived H9c2 cardiomyoblasts [7]. Peroxisome proliferator-activated receptors (PPARs) are ligand-activated nuclear transcriptional activators belonging to the superfamily of nuclear receptors [8,9]. Three subtypes have been described, including PPAR{alpha}, {gamma} and β/{delta} or {delta} (also called NR1C1, NR1C3 and NUC-1, respectively). They form heterodimers with retinoid X receptors (RXRs), bind to the hexanucleotidic PPAR responsive element (PPRE) and thereby regulate the expression of target genes. Recent studies have documented that PPAR{alpha} and {gamma} exert antioxidant effects on vasculature by inhibiting the production of superoxide anion by NADPH oxidase [10,11]. Moreover, the PPAR{alpha} ligand Bezafibrate and the PPAR{gamma} agonists Troglitazone and Pioglitazone have been reported to induce the expression of Cu/Zn superoxide dismutase, thereby accounting for an important antioxidant role of these PPAR subtypes [12,13]. More recently, a functional PPRE was identified in the rat catalase promoter and PPAR{gamma} was shown to induce catalase expression via fixation to the PPRE [14]. The function in the heart of the less studied and ubiquitously expressed PPAR{delta} subtype remains, however, largely unknown. We therefore investigated the role of PPAR{delta} in regulating the apoptotic cell death induced by H2O2 exposure in the H9c2 rat cardiomyoblasts using GW501516 as a selective PPAR{delta} ligand. We next examined the specificity of action of PPAR{delta} by using transient transfection with a dominant negative mutant of PPAR{delta}. Finally, the effect of GW501516 treatments on catalase expression was assessed by semi-quantitative RT-PCR and Western Blotting. The results presented here argue for an important role of PPAR{delta} in protecting cardiac cells against oxidative stress. Indeed, PPAR{delta} activation led to inhibition of H2O2-induced apoptosis and upregulated the expression of the antioxidative enzyme catalase.


   2. Methods
Top
 Abstract
 1. Introduction
 2. Methods
3. Results
 4. Discussion
 Acknowledgments
 References
 
2.1 Cell culture and treatments
The rat embryonic-heart derived H9c2 cell line (American Type Culture Collection, Rockville, MD, USA) was cultured at 37 °C and 5% CO2 in 100-mm dishes containing Dulbecco's Modified Eagle Medium (DMEM) (Gibco) supplemented with 10% (v/v) Fetal Bovine Serum (FBS) (Sigma). Cells were maintained in exponential phase of growth and were subcultured when they reached about 70% confluence at a split ratio of 1:3. For the various experiments H9c2 cells were seeded at the density of 106 cells per 100-mm dish, 1 x 105 cells per 35-mm well of 6-well plates or 4 x 104 cells per 20-mm well of 24-well plates. After 24 h culture, cells were made quiescent by serum starvation (0.5% FBS) for 24 h before treatment with the PPAR{delta} agonist GW501516 dissolved in dimethyl sulfoxide (DMSO) (Sigma) or H2O2 (Sigma) diluted in phosphate buffered saline (PBS). For control experiments, cells received DMSO (0.1%) or PBS as vehicle.

2.2 Cell viability assay
Cells were seeded in 24-well plates and cultured as mentioned above. 3,(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium (MTT) (Sigma) was dissolved in PBS containing 0.5 g/l of glucose and 10 mg/l of CaCl2 at a concentration of 1 mg/ml. Cells were washed twice with PBS and then incubated with the MTT solution for 1.5 h at 37 °C. Cells were then lysed with propan-2-ol, HCl 0.1 N and the medium was transferred into 96-well plates. The absorbance of the reaction solution at 620 nm was measured with an ELISA-plate reader (Titertek Multiskan MCC/340).

2.3 Plasmids
We used a plasmid containing three copies of the PPRE site of the rat Acyl-CoA oxidase promoter (PPRE-TK-luc) derived from the pGL3 luciferase expression vector (Promega) and a β-galactosidase expression vector. Two plasmids were used to overexpress either the native form of PPAR{delta} (PPAR{delta}) or a dominant negative mutant of PPAR{delta} (PPAR{delta}-DN) where the glutamate residue at position 411 of PPAR{delta} cDNA is substituted by a proline residue. PPAR{delta} cDNAs were cloned at the BamHI site of the pcDNA3.1 vector (Invitrogen) [15]. A plasmid (kindly provided by Dr. Stephan Lortz, Institut für Klinische Biochemie, Hannover, Germany) was used to overexpress the human catalase cDNA cloned at an EcoRI site of the pcDNA3.1 vector [16].

2.4 Transient transfection
Transient transfections were carried out on 24-well plates or 6-well plates. Cells were transfected using JetPEI (Qbiogene), either with 0.5 µg of PPRE-TK-luc or with 50 ng of PPAR{delta} or with various amounts of PPAR{delta}-DN. In another set of experiments, cells were transfected with 500 ng of human catalase expression vector. Control of transfection efficiency was always realized using parallel transfection with 200 ng of the β-galactosidase expression vector and total amount of plasmid DNA per well was kept constant by adding the corresponding empty vectors.

After 6 h transfection, cells were washed with PBS and treated for 24 h with GW501516 in DMEM-10% FBS. Luciferase activity was measured with a luminometer (Victor2, Wallac) using the luciferase reporter assay kit (Promega) and normalized using a β-galactosidase reporter assay kit (Promega).

2.5 Reverse transcriptase-polymerase chain reaction analysis
Total RNA was extracted using the RNeasy mini kit (Qiagen) according to the manufacturer's instructions and a treatment with the RNase-Free DNase Set (QIAGEN) was subsequently performed. To ensure that genomic DNA-free RNA was purified, RNA was analyzed by ethidium-bromide-agarose gel electrophoresis using 1% agarose gels and also subjected to PCR without reverse transcription step. One µg of total RNA was subjected to reverse transcription using the iScript Reverse Transcriptase (BioRad) conformly to the manufacturer's instructions. Generated single strand cDNAs were double strand synthesized and amplified by PCR. Primer sequences used were as follows: PPAR{alpha} forward primer, 5'-GGTCAAGGCCCGGGTCATACTCG-3', reverse primer, 5'-AAGCATTGCCGTACGCGATCAG-3' (350 pb); PPAR{gamma} forward primer, 5'-CTCCCAGCTGTCGCAAGGTGC-3', reverse primer, 5'-GCAATCGATAGAAGGAACACT-3' (174 pb); PPAR{delta} forward primer, 5'-GCCCTTTTCATTGCCGCCATCAT-3', reverse primer, 5'-TGGGCGGGTCCTCTGAACAGTCC-3' (350 pb); catalase forward primer, 5'-TACTACCCCAACAGCTT-3' reverse primer, 5'-GCTAAGCCCTAATCTTTAA-3' (589 pb); glyceraldehyde phosphate dehydrogenase (GAPDH) forward primer, 5'-CGTCTTCACCACCATGGAGA-3', reverse primer, 5'-CGGCCATCACGCCACAGTTT-3' (395 pb). For amplification of the PPARs isotype fragments the following conditions were used: initial denaturation step at 94 °C (2 min), denaturation at 94 °C (15 s), annealing at 55 °C (1 min), extension at 68 °C (2 min) and final extension at 68 °C (5 min), over a total of 40 cycles. PCR amplification of the catalase fragment was conducted as follows: initial denaturation step at 94 °C (5 min), denaturation at 94 °C (45 s), annealing at 57 °C (45 s), extension at 72 °C (90 s) and final extension at 72 °C (5 min), over a total of 35 cycles. PCR amplification of the GAPDH fragment was conducted as follows: initial denaturation step at 94 °C (5 min), denaturation at 94 °C (1 min), annealing at 55 °C (1 min), extension at 72 °C (2 min) and final extension at 72 °C (5 min), over a total of 28 cycles. Amplification products were analyzed by ethidium-bromide-agarose gel electrophoresis using 2% agarose gels and visualized under UV illumination. Product bands were captured and analyzed by densitometry using a Gel-Doc and the QuantityOne software (BioRad). Relative intensities of the PCR products were normalized to GAPDH.

2.6 Western blotting
Cells were harvested by trypsinization and centrifuged at 470 x g at 4 °C for 10 min. The cell pellet was resuspended in lysis buffer (0.25% NP-40, 1 mM PMSF) and kept on ice for 30 min. The lysate was sonicated (10 s) and then centrifuged at 13,000 x g at 4 °C for 10 min. Protein concentration in the supernatant was determined by the Bradford assay (BioRad). Samples were diluted (1:1) in sample buffer (67 mM Tris–HCl, pH 6.8, 30% glycerol, 2% SDS, and 0.01% bromophenol blue) and boiled for 3 min. Thirty µg of protein were separated by SDS-PAGE using 10% acrylamide gels (Euromedex) and transferred to a nitrocellulose membrane (Schleicher and Schuell). To ensure quality of the transfer, the blots were stained with Ponceau Red and then washed with deionized water. Blots were probed with an anti-human erythrocyte catalase antibody (1:500) (Calbiochem) as the primary antibody and then with horseradish peroxidase conjugated to goat anti-rabbit IgG (1:1000) (Santa Cruz Biotechnology) as the secondary antibody. Detection of the bound antibodies was carried out using the ECL kit (Amersham Pharmacia Biotech).

2.7 Analysis of apoptosis
2.7.1 Terminal transferase nick-end labeling (TUNEL)
Cells grown in Petri dishes were fixed with methanol for 5 min at –20 °C and rehydrated with PBS. TUNEL labeling was performed using the ApopTag Peroxidase in situ Apoptosis Detection Kit (Oncor) according to the manufacturer's instructions.

2.7.2 Annexin V/propidium iodide/Hoechst 33342 staining
To detect early stages of apoptosis, an Annexin-V-FLUOS staining kit (Roche) was used according to the manufacturer's instructions. Briefly, cells were washed with PBS and incubated with Annexin-V/propidium iodide (PI)/Hoechst 33342 for 15 min at room temperature. Cells were visualized with a fluorescence microscope and Annexin-V positive and PI negative cells were assessed as apoptotic. The percentage of apoptotic cells was calculated by dividing Annexin-V positive cells by the total number of cells visualized by Hoechst staining. Three randomly chosen optical fields per preparation were digitized and each experimental condition was done in triplicate. The mean percentage of apoptotic cells was averaged over a total of three independent experiments.

2.7.3 DNA fragmentation analysis
Cells were homogenized in 1 ml of lysis buffer (20 mM Tris–HCl, pH 8.0, 5 mM EDTA, 0.5% SDS, 0.5 mg/ml proteinase K) and incubated for 15 h at 42 °C under constant agitation. Proteins were then precipitated by a 6 M NaCl solution and centrifuged at 2500 x g at 4 °C for 15 min. Supernatants containing genomic DNA were then treated with RNase A at 37 °C for 30 min. The genomic DNA was precipitated 3 h at –70 °C with 2.5 volumes of 100% ethanol and 0.2 volume of 3 M sodium acetate. Samples were then centrifuged at 20,800 x g at 4 °C for 30 min. Resulting pellets were washed with 70% ethanol and resuspended in 40 µl of nuclease-free water. Genomic DNA extracts (10–20 µl) were run on 1.8% agarose gels and visualized under UV illumination.

2.7.4 Caspase-3 activity measurement
Caspase-3 activity was measured using the CaspACE kit (Promega) according to the manufacturer's instructions. Briefly, cells were collected by trypsinization followed by centrifugation at 470 x g at 4 °C for 10 min. The cell pellet was washed with cold PBS, centrifuged at 470 x g at 4 °C for 10 min and then resuspended in 40 µl of lysis buffer. After 3 rapid freeze–thaw cycles, the lysate was incubated on ice for 15 min and then centrifuged at 15,000 x g at 4 °C for 20 min. The protein concentration in the supernatant was determined by the Bradford assay and 75 µg of proteins were incubated with the caspase-3 substrate at 37 °C for 4 h. The absorbance of the reaction was then spectrophotometrically measured at 414 nm.

2.8 Statistics
Each experiment was repeated 3 times. The results are presented as sample mean ± S.D. Statistical analysis was performed by ANOVA and the Bonferroni/Dunn post hoc test using the Statview software. Differences were considered significant at p<0.05.


   3. Results
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
4. Discussion
 Acknowledgments
 References
 
3.1 PPAR isotype expression in H9c2 cells
Prior to evaluate the effect of PPAR{delta} on H2O2-induced cytotoxicity, we first determined by RT-PCR the expression profile of the 3 PPAR isotypes in the left ventricle of 3-month-old Wistar Kyoto rats and then in our model of H9c2 cardiomyoblasts. PPAR{alpha} and PPAR{delta} were abundantly expressed in the left ventricle whereas PPAR{gamma} was detected at a lower level (Fig. 1A). The major isotype expressed in H9c2 cells was PPAR{delta} (Fig. 1B). PPAR{alpha} was only weakly detected and despite increasing the number of PCR cycles no PPAR{gamma} expression was detected.


Figure 1
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  		Fig. 1 Pattern of expression of PPAR isotypes in H9c2 cells. (A) Total RNA from left ventricles of 3-month-old Wistar Kyoto rats were extracted and subjected to RT-PCR for a total of 36 cycles. The electrophoresis gel presented is representative of 3 independent experiments giving similar results. (B) The electrophoresis gel is representative of 3 independent RT-PCR experiments performed on H9c2 cells with 40 PCR cycles.

 
3.2 The PPAR{delta} agonist GW501516 attenuates H2O2-induced cytotoxicity
Using the MTT cytotoxicity test, we tested the cytoprotective effect of GW501516 using H2O2 to generate an oxidative stress. We first treated the cells with concentrations of GW501516 ranging from 10 nM to 1000 nM. Reduction of cell viability was never observed (Fig. 2A). We then determined the cytotoxicity of H2O2 by treating the cells with increasing concentrations of H2O2 (50–200 µM). As shown in Fig. 2B, a significant decrease in cell viability was observed for the 3 tested concentrations reaching a maximum of 84% reduction for a treatment of 200 µM H2O2 (p<0.001). When cells were pretreated for 24 h with growing concentrations of GW501516 (10 nM–1000 nM) prior to a 100 µM H2O2 stress, significant increases in cell viability (22% and 43%; p<0.001) were observed (Fig. 2C).


Figure 2
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  		Fig. 2 GW501516 attenuates H2O2-induced cytotoxicity. (A) Effect of GW501516 on cell viability. Cell viability was estimated using the MTT test after a 24 h treatment with various concentrations of GW501516. (B) Effect of H2O2 on cell viability. Cell viability was estimated using the MTT test 15 h after treatment with various concentrations of H2O2. (C) GW501516 protects the cells from H2O2-induced cytotoxicity in a dose-dependent manner. Cells were pretreated for 24 h with increasing concentrations of GW501516 and then treated for 15 h with 100 µM H2O2. The results are presented as mean percentage of cell viability vs. control (CTL) ± S.D. For each group n=12. *p<0.001 vs. CTL, #p<0.001 vs. H2O2-treated cells, SND=statistically not different.

 
3.3 GW501516 inhibits H2O2-induced apoptosis
Using the TUNEL method we checked that nuclei of stressed cells (100 µM H2O2 for 15 h) appeared dark brown under the microscope indicating DNA fragmentation typical of apoptosis (Fig. 3A). We next investigated the effect of GW501516 on H2O2-induced apoptosis by performing an Annexin-V/PI/Hoechst staining. Very few Annexin-V positive cells were detected in control cultures or GW501516 treated cultures (0.87% and 0.84%, respectively) whereas cultures treated with 100 µM H2O2 for 5 h showed a large number of positively stained cells (33.7%). Pretreatment with 100 nM GW501516 for 24 h however led to a decrease in the number of apoptotic cells with only 3.1% of positive cells (Fig. 3B and C). We then quantified caspase-3 activity 15 h after induction of the oxidative stress. A 24 h 100 nM GW501516 treatment did not induce a statistically different caspase-3 activity from control cultures. However, 100 µM H2O2 led to a 6.5-fold increase in caspase-3 activity. This was totally abolished when the cells received the pan-caspase inhibitor Z-VAD-FMK. Pretreatment with GW501516 decreased significantly (4.25-fold, p<0.001) caspase-3 activity generated by H2O2 (Fig. 3D). The inhibition of H2O2-induced apoptosis was also studied by DNA laddering. DNA fragmentation induced by 100 µM H2O2 for 48 h was fully prevented by pretreatment with 100 nM GW501516 for 24 h. Three independent experiments were conducted and gave similar results (data not shown).


Figure 3
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  		Fig. 3 GW501516 inhibits H2O2-induced apoptosis. (A) TUNEL labeling. Cells were treated or not with 100 µM H2O2 for 15 h and then fixed with methanol. TUNEL was then performed and dark brown labeled apoptotic nuclei (right panel, arrows) were visualized under the microscope. Photos shown are representative of three independent experiments giving similar results. (B) Annexin-V/PI/Hoechst 33342 fluorescent staining. (1) Control cells (vehicle), (2) cells treated 24 h with 100 nM GW501516, (3) cells treated 5 h with 100 µM H2O2, (4) cells pretreated 24 h with 100 nM GW501516 and then 5 h with 100 µM H2O2. (C) Quantification of the percentage of apoptotic cells (Annexin-V positive and PI negative cells). Results are presented as mean percentage of apoptotic cells ± S.D. For each group n=3, see Methods. *p<0.001 vs. CTL, #p<0.001 vs. H2O2-treated cells, SND=statistically not different. (D) Measurement of H2O2-induced caspase-3 activity. Cells received the same treatments as indicated above. The pan-caspase inhibitor Z-VAD-FMK (50 µM) was added at the same time as H2O2. Three independent experiments were performed. Results are presented as mean optical density (414 nm) ± S.D. For each group n=3. *p<0.001 vs. CTL, #p<0.001 vs. H2O2-treated cells.

 
3.4 The protective effect of GW501516 is PPAR{delta} dependent
To determine whether GW501516 action was mediated by PPAR{delta}, we performed transient transfection with a dominant negative mutant of PPAR{delta} (PPAR{delta}-DN). We first validated this construction in our H9c2 model by testing its ability to inhibit the induction of a PPRE-TK-luc plasmid in cotransfection studies with a PPAR{delta} expression vector (PPAR{delta}). As shown in Fig. 4A, treatment of control cells or cells transfected with PPAR{delta}-DN with GW501516 did not induce luciferase activity. However, cells transfected with PPAR{delta} showed a 4.5-fold GW501516-dependent induction of luciferase. The dominant negative action of PPAR{delta}-DN was then analyzed by transfection of H9c2 cells with a constant amount of PPAR{delta} and increasing amounts of PPAR{delta}-DN. PPAR{delta}-DN inhibited in a dose-dependent manner the PPAR{delta}-mediated transactivation of the luciferase reporter. Luciferase induction was decreased by 45.5% and 55% when PPAR{delta}-DN was transfected in a 2- and 4-fold excess, respectively.


Figure 4
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  		Fig. 4 Expression of a dominant negative mutant of PPAR{delta} abolishes the protective action of GW501516. (A) Validation of the dominant negative effect of PPAR{delta}-DN in H9c2 cells. H9c2 cells were transfected with the PPRE-TK-luc and β-galactosidase reporter plasmids and the indicated amounts of PPAR{delta} and PPAR{delta}-DN expression plasmids. Cells were cultured in the absence (open bars) or presence (solid bars) of 100 nM GW501516 for 24 h prior to determination of luciferase and β-galactosidase activities. Results are presented by taking as 1 the value obtained for cells cultured in control medium and are the mean ± S.D. of three independent experiments. (B) Cells were transfected with the indicated amounts of PPAR{delta}-DN expression plasmid. They were pretreated for 24 h with 100 nM GW501516 before 100 µM H2O2 treatment. Cell viability was estimated by the MTT test 15 h after induction of the oxidative stress. Each transfection was performed in triplicate and the results are presented as the mean percentage of cell viability vs. control (CTL) ± S.D. For each group n=12. *p<0.001 vs. CTL, #p<0.001 vs. H2O2-treated cells, {dagger}p<0.001 vs. cells pretreated with GW501516 and then treated with H2O2, SND=statistically not different.

 
We next examined the effect of transfection with the PPAR{delta} dominant negative mutant on cell viability measured after the H2O2 treatment. Cells transfected with the pcDNA 3.1 empty vector and treated with 100 µM H2O2 showed a 60% decrease in cell viability whereas pretreatment with 100 nM GW501516 led to an increase of 32% in cell viability (p<0.001). However, increasing amounts of PPAR{delta}-DN dose-dependently prevented the GW501516-induced protection. Significant decreases in cell viability of 15% and 32.4% were observed when, respectively, 100 ng and 200 ng of PPAR{delta}-DN were transfected (p<0.001). Protection was totally abolished when 200 ng of PPAR{delta}-DN were transfected (Fig. 4B).

3.5 GW501516 induces catalase expression in H9c2 cardiomyoblasts
It has already been reported that the catalase promoter contains a functional PPRE and that PPAR{gamma} can induce the expression of this gene. To test whether PPAR{delta} can also regulate the expression of catalase, we determined the effect of GW501516 on catalase expression using semi-quantitative RT-PCR for transcripts levels (Fig. 5A and B). Cells were treated for 6 and 24 h with increasing concentrations of GW501516 and RT-PCR was then performed. The levels of catalase transcripts were significantly increased at 6 h and also 24 h, but to a lesser extent (p<0.001). The expression of catalase was regulated in a dose-dependent manner with significant increases at a concentration of 100 nM of GW501516 for both times of treatment (2-fold increase for 6 h and 1.6-fold increase for 24 h). The maximal increase was observed for a GW501516 concentration of 1000 nM (4.9-fold increase for 6 h and 3-fold increase for 24 h). Analysis by Western Blotting (Fig. 5C) showed also a significant increase in catalase protein expression induced after 24 h or 48 h GW501516 treatment.


Figure 5
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  		Fig. 5 GW501516 induces the expression of catalase transcripts and protein. (A) Cells were treated for the indicated times and with the indicated concentrations of GW501516. RNAs were submitted to RT-PCR for catalase (589 pb) and GAPDH (395 pb) cDNAs. The amplification products were separated on a 2% agarose gel and visualized under UV illumination. The gel presented is representative of three experiments giving similar results. (B) Densitometric analysis of PCR products presented in (A). Catalase intensity was normalized to GAPDH intensity and results are presented as the mean ratio of catalase/GAPDH intensities ± S.D. by taking the value 1 for untreated cells (CTL). Three independent experiments were performed. *p<0.001 vs. CTL. (C) Cells were treated for the indicated times and with the indicated concentrations of GW501516. Proteins were extracted and catalase expression was estimated by Western Blotting. The 42 kDa band, corresponding to the major muscular protein actin, was used as internal control and visualized on the nitrocellulose filter with Ponceau Red staining.

 
3.6 GW501516 increases catalase expression during oxidative stress
As previously, RT-PCR was performed on RNA from cells treated with H2O2 in the presence or absence of GW501516 (Fig. 6). H2O2 clearly increased (approximately 1.5-fold) the signal corresponding to catalase transcripts compared to control cells. Addition of GW501516 potentiated the H2O2-induced catalase upregulation. Basal level of catalase expression (without any treatment) was increased in the presence of H2O2 by 2- to 2.5-fold by 100 and 1000 nM GW501516, respectively.


Figure 6
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  		Fig. 6 GW501516 potentiates the expression of catalase transcripts in the presence of H2O2. Cells were treated for 24 h with the indicated concentrations of GW501516 and treated or not with 100 µM H2O2. Extracted RNAs were submitted to RT-PCR for catalase (589 pb) and GAPDH (395 pb) cDNAs. The amplification products were separated on a 2% agarose gel and visualized under UV illumination. The gel presented is representative of three experiments giving similar results.

 
3.7 Catalase overexpression protects cells from H2O2-induced apoptosis
Cells were transfected with a catalase expression vector and treated or not with H2O2. Apoptosis was assessed by Annexin-V/PI labeling (Fig. 7). Catalase overexpression noticeably reduced phosphatidylserines externalization triggered by oxidative stress.


Figure 7
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  		Fig. 7 Catalase overexpression inhibits H2O2-induced apoptosis. Annexin-V/PI/Hoechst 33342 fluorescent staining. (1) Control cells (empty pcDNA3.1), (2) cells transfected with 500 ng of catalase expression vector, (3) cells treated 5 h with 100 µM H2O2 (empty pcDNA3.1), (4) cells transfected with 500 ng of catalase expression vector and then treated 5 h with 100 µM H2O2.

 

   4. Discussion
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
Acknowledgments
 References
 
The present study demonstrates for the first time that the nuclear receptor PPAR{delta} exerts an antioxidative role in rat cardiac myoblasts by inhibiting H2O2-induced apoptosis. In the first instance we have shown by the use of semi-quantitative RT-PCR that PPAR{delta} was the predominant PPAR isotype in H9c2 cardiomyoblasts. PPAR{alpha} was slightly expressed in this cell line. PPAR{gamma}, however, could not be detected in H9c2 cells contrary to results obtained in ventricular tissue from Wistar Kyoto rats. A recent study showed that only PPAR{delta} is expressed in H9c2 cells [17]. Those authors, however, used a Northern Blotting approach which is less sensitive than RT-PCR.

It is now well established that H2O2-induced oxidative stress induces apoptosis in cardiomyocytes [7,18]. Apoptosis is also clearly induced during myocardial infarction [19]. We have shown that treatment of H9c2 cells with concentrations of H2O2 ranging from 50 µM to 200 µM induces a dose-dependent decrease in cell viability as measured by the MTT test. Moreover, typical features of apoptosis such as TUNEL labeling, phosphatidylserines externalization, a strong caspase-3 activity and DNA ladder indicate that the cell death observed with the cell viability studies is of apoptotic nature. The apoptosis observed in the present study remains classical. On the contrary, hypoxia-reoxygenation in neonatal rat cardiomyocytes appears to induce a less typical apoptotic cell death associated with membrane disintegrity [20]. We have demonstrated that the pretreatment of cells with the PPAR{delta} selective agonist GW501516 inhibited H2O2-induced apoptosis. Other studies have also demonstrated that PPARs are involved in the regulation of apoptotic processes. PPAR{alpha} was shown to have an antiapoptotic role in liver [21] whereas PPAR{gamma} was shown to be proapoptotic in several cell types [22,23] and antiapoptotic in others [24]. Recently, PPAR{delta} was shown to be implicated in the regulation of apoptosis in the keratinocyte HaCaT cell line [25]. The activation of PPAR{delta} indeed inhibited growth factor deprivation-induced apoptosis by upregulating ILK and PDK1, two members of the PI3K-Akt1 pathway. Moreover, it has been shown that H2O2 induces the degradation of Akt leading to a caspase-3-independent-apoptotic cell death [26,27]. We thus hypothesize that the antiapoptotic effect of PPAR{delta} observed in the present study is probably not due to an activation or upregulation of the PI3K-Akt signalling pathway since the cell death observed here is caspase-3 dependent.

GW501516 is a PPAR{delta} selective ligand when used at nanomolar concentrations in macrophages [28]. By using GAL4-PPARs ligand binding domain chimeras transfections the authors also demonstrated that GW501516 up to 1000 nM was unable to activate PPAR{alpha} and {gamma}. However, in our hands, some PPAR{alpha} activation is observed for GW501516 treatment concentrations of 100 and 1000 nM. We therefore tested the selectivity of GW501516 and the specificity of action of PPAR{delta} by transfecting H9c2 cells either with native PPAR{delta} or with the previously described dominant negative mutant (PPAR{delta}-DN) [15]. This dominant negative mutant of PPAR{delta} is able to bind the ligand and the PPRE but remains in a repressive form. Treatment of non-transfected cells with GW501516 could not induce luciferase-reporter activity. This could be explained in part by the fact that endogenous PPAR{delta} expression is too low to activate the overexpressed luciferase reporter. As expected, however, PPAR{delta}-DN dose dependently inhibited PPAR{delta}-induced luciferase activity. We then transiently transfected PPAR{delta}-DN to determine whether it could abolish the protection induced by GW501516 in cell viability studies. The fact that PPAR{delta}-DN dose-dependently reduced the protection against oxidative stress induced by GW501516 argues for a specific protective role of PPAR{delta} against an oxidative stress generated by H2O2.

It was recently shown that the promoter of the H2O2 detoxifying enzyme catalase contains a functional PPRE [14]. Moreover, PPAR{gamma} was shown to induce the expression of catalase by binding to this PPRE. We therefore tested the effect of PPAR{delta} on catalase expression by using two approaches, semi-quantitative RT-PCR and Western Blotting. RT-PCR analysis revealed that 6 h and 24 h GW501516 treatments induced a significant rise in catalase transcript levels reaching a 5-fold induction after 1000 nM GW501516 treatment during 6 h. Western Blotting protein analysis showed an increase in catalase protein content after treatment with the PPAR{delta} ligand thus demonstrating that the induced transcripts are properly translated into protein.

Although our study focused on the pretreatment effect of the PPAR{delta} agonist, demonstrating its major effect on catalase induction, our data also indicate that GW501516 also increases catalase expression in the presence of H2O2. The transcript level is however lower as expected, probably due to RNA degradation during oxidative stress. Moreover, the overexpression of catalase induced by GW501516 readily protects cells from H2O2-induced apoptosis as evidenced by our experiment using plasmid-induced catalase overexpression.

In conclusion, the present study demonstrates a new role for PPAR{delta} in cardiac myoblasts. In accordance with the well established antioxidative effects of PPAR{alpha} and {gamma} in the vascular system [29–31], PPAR{delta} appears to be a newly recognized but important actor of the PPAR family in stimulating the expression of the antioxidative enzyme catalase in the myocardium.


   Acknowledgments
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Acknowledgments
References
 
We kindly acknowledge Frédéric Thoreau at Fournier Pharma Laboratories for the help provided on PPAR RT-PCR analysis. We also thank Pr. Jean-Jacques Michaille (Université de Bourgogne, France) and Pr. Colin Green (University of Auckland, New Zealand) for critical reading of the manuscript.


   Notes
 
1 Present affiliation: Laboratoire de Biologie Moléculaire et Cellulaire, Université de Bourgogne, 6 Bd Gabriel, 21000 Dijon, France. Back

Time for primary review 21 days


   References
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Acknowledgments
 References
 

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