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EMBRYONIC STEM CELLS |
Agonists Enhance Cardiomyogenesis of Mouse ES Cells by Utilization of a Reactive Oxygen Species-Dependent Mechanism
aDepartment of Physiology, Faculty of Medicine, Justus-Liebig-University, Giessen, Germany;
bDepartment of Internal Medicine I, Cardiology Division, Friedrich-Schiller-University, Jena, Germany
Key Words. Embryonic stem cells • Peroxisome proliferator-activated receptor • heart • Cardiomyogenesis • Reactive oxygen species
Correspondence: Correspondence: Heinrich Sauer, M.D., Department of Physiology, Justus-Liebig-University Giessen, Aulweg 129, 35392 Giessen, Germany. Telephone: +49-641-9947333; Fax: +49-641-9947219; e-mail: heinrich.sauer{at}physiologie.med.uni-giessen.de
Received on July 6, 2007;
accepted for publication on October 9, 2007.
First published online in STEM CELLS EXPRESS October 18, 2007.
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ABSTRACT |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionAcknowledgmentsReferences |
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, -β and -
) are nuclear receptors involved in transcriptional regulation of lipid and energy metabolism. Since the energy demand increases when cardiac progenitor cells are developing rhythmic contractile activity, PPAR activation may play a critical role during cardiomyogenesis of embryonic stem (ES) cells. It is shown that ES cells express PPAR, -β, and - mRNA during differentiation of ES cells towards cardiac cells. Treatment with PPAR agonists (WY14643, GW7647, and ciprofibrate) significantly increased cardiomyogenesis and expression of the cardiac genes MLC2a, ANP, MHC-β, MLC2v, and cardiac -actin. Furthermore, WY14643 increased PPAR gene expression and the expression of the cardiogenic transcription factors GATA-4, Nkx2.5, DTEF-1, and MEF 2C. In contrast, the PPAR antagonist MK886 decreased cardiomyogenesis, whereas the PPARβ agonist L-165,041 as well as the PPAR agonist GW1929 were without effects. Treatment with PPAR, but not PPARβ, and PPAR agonists and MK886, resulted in generation of reactive oxygen species (ROS), which was inhibited in the presence of the NADPH oxidase inhibitors diphenylen iodonium (DPI) and apocynin and the free radical scavengers vitamin E and N-(2-mercapto-propionyl)-glycine (NMPG), whereas the mitochondrial complex I inhibitor rotenone was without effects. The effect of PPAR agonists on cardiomyogenesis of ES cells was abolished upon preincubation with free radical scavengers and NADPH oxidase inhibitors, indicating involvement of ROS in PPAR, mediated cardiac differentiation. In summary, our data indicate that stimulation of PPAR but not PPARβ and - enhances cardiomyogenesis in ES cells using a pathway that involves ROS and NADPH oxidase activity. Disclosure of potential conflicts of interest is found at the end of this article.
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionAcknowledgmentsReferences |
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is highly expressed in tissues such as liver, muscle, kidney, and heart, where it stimulates the β-oxidative degradation of fatty acids. PPAR is predominantly expressed in intestine and adipose tissue where it triggers adipocyte differentiation and promotes lipid storage. PPARβ/
is ubiquitously expressed and activators exert anti-inflammatory and antihypertrophic effects in cardiomyocytes. The hypolipidemic fibrates and the antidiabetic glitazones are synthetic ligands for PPAR and PPAR, respectively. Furthermore, fatty acids and eicosanoids are natural PPAR ligands: PPAR is activated by leukotriene B4, whereas prostaglandin J2 is a PPAR ligand [2]. The synthetic PPAR ligands, besides their role as hypolipidemic and insulin-sensitizing agents, have been previously shown to exert beneficial effects in inhibiting the pathogenesis of heart failure, atherosclerosis, and hypertension although the underlying signaling cascades are not yet unraveled [2, 3]. In the course of cardiac hypertrophy, which is an adaptive response of the heart, during which terminally differentiated cardiomyocytes increase in size without undergoing cell division, PPAR expression is downregulated with the consequence of reduced fatty acid oxidation and increased glycolysis [4]. In patients with hypertensive heart disease, the expression of native PPAR protein was decreased and truncated PPAR protein was overexpressed [5]. Chronic reduction in fatty acid oxidation due to downregulation of PPAR leads to cardiac lipotoxicity and subsequent apoptosis. Furthermore, chronic downregulation of PPAR during cardiac hypertrophy may further impair cardiac function due to insufficient production of ATP, thereby leading to cardiac failure [6]. Animals with genetic inactivation of PPAR are viable and fertile and exhibit no detectable gross phenotypic defects [7]. However, short-term starvation caused hepatic steatosis, myocardial lipid accumulation, and hypoglycemia, with an inadequate ketogenic response in adult mice lacking PPAR [8]. PPAR is upregulated in the heart during embryogenesis which is presumably due to an increased energy demand arising from the increasing cardiac workload in the growing embryo [6]. Interestingly, PPAR expression during embryogenesis is transient, suggesting that a period of PPAR expression during embryonic heart development may be required for normal heart function [6].
Recently, it was shown that the PPAR agonist WY14643 stimulated cardiac differentiation from mouse ES cells [9] by a so far unknown signaling pathway. In the latter study it was suggested that PPAR activity is required during differentiation of stem cells towards cardiac cells with the emergence of contractile activity and concomitant increased ATP consumption. In the present study, we demonstrate that PPAR activation but not PPARβ and - are required for the stimulation of cardiac cell differentiation. It is furthermore demonstrated that PPAR stimulation results in activation of NADPH oxidase and subsequent ROS generation, which is, as previous studies from our group have suggested [10–13], a prerequisite for cardiovascular differentiation of ES cells.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionAcknowledgmentsReferences |
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Measurement of ROS Generation
Intracellular ROS levels were measured using the fluorescent dye 2'7'-dichlorodihydrofluorescein diacetate (H2DCF-DA) (Molecular Probes, Eugene, OR), which is a nonpolar compound that is converted into a nonfluorescent polar derivative (H2DCF) by cellular esterases after incorporation into cells. H2DCF is membrane impermeable and is rapidly oxidized to the highly fluorescent 2'7'-dichlorofluorescein (DCF) in the presence of intracellular ROS. For the experiments, embryoid bodies were incubated in serum-free medium, and 20 µM H2DCF-DA dissolved in dimethyl sulfoxide (DMSO) was added. After 30 minutes, intracellular DCF fluorescence (corrected for background fluorescence) was evaluated in 3,600 µm2 regions of interest using an overlay mask unless otherwise indicated. For fluorescence excitation, the 488 nm band of the argon ion laser of a confocal laser scanning microscope (Leica SP2 AOBS, Leica, Bensheim, Germany) was used. Emission was recorded at an emission band of 515–550 nm.
Immunohistochemistry
Immunohistochemistry was performed with whole mount embryoid bodies. As primary antibody, a monoclonal mouse anti -actinin antibody (Sigma) was used. The respective tissues were fixed in icecold methanol for 20 minutes at –20°C, and washed 3 times with PBS containing 0.01% Triton X-100 (PBST) (Sigma). Blocking against unspecific binding was performed for 60 minutes with 10% fetal calf serum (FCS) dissolved in 0.01% PBST. The tissues were subsequently incubated for 60 minutes at room temperature, with primary antibodies dissolved in PBS supplemented with 10% FCS in 0.01% PBST. The tissues were thereafter washed 3 times with PBST (0.01% Triton) and reincubated with a Cy5-conjugated goat anti mouse IgG (H+L) (Dianova, Hamburg, Germany) at a concentration of 3.8 µg/ml in PBS containing 10% FCS in 0.01% PBST. After washing 3 times in PBST (0.01% Triton), the tissues were stored in PBS until inspection. Fluorescence recordings were performed by means of a confocal laser scanning setup (Leica TCS SP2). The confocal setup was equipped with a 5 mW helium/neon laser single excitation 633 nm (excitation of Cy5). Emission was recorded at >665 nm. The pinhole settings of the confocal setup were adjusted to give a full width half maximum (FWHM) of 10 µm. Fluorescence was recorded in a depth of 80–120 µm in the depth of the tissue, and the fluorescence values in the respective optical sections were evaluated by the image analysis software of the confocal setup.
Analysis of the Size of Cardiac Cell Areas
Embryoid bodies plated on day 4 of cell culture to Petriperm Petri dishes (InVitro Systems Inc. Göttingen, Germany) were fixed on day 10 of cell culture, and stained with an antibody against -actinin. Immunofluorescence of -actinin-positive cell areas was assessed by confocal laser scanning microscopy. By means of the "area measure" option of the image analysis software of the confocal setup, the size of the -actinin-positive cell areas was measured after correction for background fluorescence. Mean values of all -actinin-positive areas of cardiac cells which remained either untreated or were treated as indicated, were collected from 60 embryoid bodies in individual experiments from at least 3 different cell cultures.
Real Time RT PCR
Total RNA from CCE embryoid bodies treated for 24 hours with the substances as indicated, was prepared using Trizol (Invitrogen) method followed by genomic DNA digestion using DNAse I (Invitrogen, Karlsruhe, Germany). Total RNA concentration was determined by OD260 nm method. cDNA synthesis was performed using 2 µg RNA with MMLV RT (Invitrogen). Primer concentration for qPCR was 10 pM. Primer (Invitrogen) sequences were as follows:
PPARrev: 5'-GAA GGT GTC ATC TGG ATG GGT-3'
PPARβ fwd: 5'-TTG AGC CCA AGT TCG AGT TTG-3'
rev: 5'-CGG TCT CCA CAC AGA ATG ATG-3'
PPAR
rev: 5'-CTC CAA GAA TAC CAA AGT GCG A-3'
cardiac fwd: 5'-CTC GAT TCT GGC GAT GGT GTA-3'
ANP fwd: 5'-CGT GCC CCG ACC CAC GCC AGC ATG GGC TCC-3'
rev: 5'-GGC TCC GAG GGC CAG CGA GCA GAG CCC TCA-3'
MLC2a fwd: 5'-TCA GCT GCA TTG ACC AGA AC-3'
rev: 5'-AAG ACG GTG AAG TTG ATG GG-3'
MLC2v fwd: 5'-AAA GAG GCT CCA GGT CCA AT-3'
rev: 5'-CCT CTC TGC TTG TGT GGT CA-3'
MHCβ fwd: 5'-CTA CAG GCC TGG GCT TAC CT-3'
rev: 5'-TCT CCT TCT CAG ACT TCC GC-3'
rev: 5'-TCCTTGAGGTTGTACAGCACA-3'
DTEF-1 fwd: 5'-CCC GAA CGC TTT CTT CCT TGT C-3'
rev: 5'-ACC TTG GTG GAG ACG CTG ATG-3'
GATA-4 fwd: 5'-TCA AAC CAG AAA ACG GAA GC-3'
rev: 5'-GTG GCA TTG CTG GAG TTA CC-3'
MEF 2C fwd: 5'-GTG GCA TTG CTG GAG TTA CC-3'
rev: 5'-TAT TCC TCT GCA GAG ACG GG-3'
Nkx2.5 fwd: 5'-CCA CTC TCT GCT ACC CAC CT-3'
rev: 5'-CCA GGT TCA GGA TGT CTT TGA-3'
CD68 fwd: 5'- TAAAGAGGGCTTGGGGCATA-3'
rev: 5' – CTCGGGCTCTGATGTAGGTC-3'
GAPDH fwd: 5'-TCC ATG CCA TGA CTG CCA CTC-3'
rev: 5'-TGA CCT TGC CCA CAG CCT TG-3'
mPolr2a fwd: 5'- GACAAAACTGGCTCCTCTGC-3'
rev: 5'- GCTTGCCCTCTACATTCTGC-3'
Amplifications were performed in an Icycler Optical Module (Biorad, Munich, Germany) using iQTM SYBR Green Supermix (Biorad). Following programs were used:
Cycle 1: Step 1: 95°C for 3 minutes (1X)
Cycle 2: Step 1: 95°C for 30 seconds (45X)
Step 2: specific annealing temperatures for 30 seconds (45X)
Step 3: 72°C for 30 seconds (45X)
Cycle 3: Step 1: 50°C for 10 minutes
Annealing temperatures were:
58.5°C for PPAR
60°C for
CT method.
Statistical Analysis
Data are given as mean values+ SD, with n denoting the number of experiments unless otherwise indicated. One-way ANOVA for unpaired data was applied as appropriate. A value of p < .05 was considered significant.
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RESULTS |
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, PPARβ, and PPAR mRNA During the Differentiation of ES cells; Correlation with -MHC and CD68 Expression, -β and - during the differentiation of ES cells, revealed that all PPAR species under investigation were expressed during differentiation, however with different time patterns (Fig. 1A–C, n = 4). Expression of PPAR increased from day 6 of differentiation on and reached maximum expression at day 12 of cell culture. A second maximum of PPAR expression was observed on day 20. PPARβ expression exerted a comparable time frame with maximum expression on day 12 and an elevated plateau of expression in late stages of differentiation. In contrast PPAR expression displayed 2 maxima of expression at days 8 and 18, respectively.
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-MHC and the macrophage marker CD68 (Fig. 2A, 2B, n = 3). Both -MHC and CD68 mRNA displayed a transient increase during ES cell differentiation with maximum expression achieved on day 10 of cell culture. This expression pattern well coincided with mRNA expression of PPAR, which showed maximum expression between days 10 and 12 of cell culture (see Fig. 1A).
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, but not PPARβ or PPAR Agonists, Stimulate Cardiomyogenesis of ES Cells agonists WY14643 (10 µM), GW7647 (10 µM), and ciprofibrate (10 µM), the PPARβ agonist L-165,041 (10 µM) or with the PPAR agonist GW1929 (10 µM) (Fig. 3A, 3B). In parallel experiments, embryoid bodies were treated with the PPAR antagonist MK886 (10 µM) (see Fig. 3A, 3B). Treatment with PPAR agonists increased the number of spontaneously contracting embryoid bodies by approximately 50% whereas the PPARβ agonist L165,041 and the PPAR agonist GW1929 were without effects (see Fig. 3A, n = 6). Furthermore, the PPAR antagonist MK886 depressed cardiomyogenesis of ES cells, however, without reaching statistical significance. The effect of PPAR agonists to increase the number of beating foci in ES cell-derived embryoid bodies was paralleled by formation of increased sizes of beating foci as evaluated by computer-assisted image analysis of -actinin-positive cardiac cell areas (Fig. 3B, n = 3). Sizes of spontaneously contracting cardiac areas were significantly increased by PPAR agonists, whereas PPARβ and agonists were without effects, and a depression in the size of cardiac areas was observed with the PPAR inhibitor MK886.
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-, Cardiac Gene-, and Cardiogenic Transcription Factor mRNA Expression through Stimulation of PPAR Receptors by WY14643 agonists should be paralleled by an increase in cardiac gene expression. Furthermore activation of PPAR receptors could induce a feed forward cycle of PPAR gene expression thereby enhancing cardiomyogenesis. To investigate these issues, PPAR mRNA expression was assessed in the absence and presence of WY14643 (10 µM) during the differentiation of ES cells (Fig. 4A, n = 3). It was apparent that presence of PPAR agonist in the cell culture medium significantly increased PPAR mRNA expression, the effect being considerably larger with longer incubation times. Treatment of embryoid bodies from day 4 to day 10 with WY14643 resulted in significantly increased expression levels of the cardiac genes ANP, MHC-β, MLC2v, MHC2a, and cardiac -actin (Fig. 4B, n = 3), thus corroborating our observation of a stimulation of cardiomyogenesis of ES cells by PPAR agonists.
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activation indeed stimulates cardiomyogenic differentiation of ES cells rather then enhancing cardiomyocyte survival it should be assumed that PPAR agonists increase the expression of cardiogenic transcription factors. To investigate this issue the mRNA expression of the cardiogenic transcription factors GATA-4, Nkx2.5, DTEF-1, and MEF 2C, was evaluated following treatment of 4-day-old embryoid bodies until day 8 of cell culture with WY14643 (10 µM) (Fig. 5A–5D). It was evident that already a 24 hour treatment with WY14643 (from day 4 to day 5) significantly increased GATA-4, Nkx2.5, and DTEF-1 gene expression, whereas a significant increase in MEF 2C mRNA expression was observed after 48 hours. Hence the data of the present study indicate that PPAR stimulation indeed stimulated the cardiogenic transcriptional program.
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Agonists in ES Cells, Role of NADPH Oxidase agonists on cardiomyogenesis of ES cells, 4-day-old embryoid bodies were incubated for 24 hours with the PPAR agonists WY14643 (10 µM), GW7647 (10 µM), and ciprofibrate (10 µM); the PPAR antagonist MK886 (10 µM); the PPARβ agonist L-165,041 (10 µM) and the PPAR agonist GW1929 (10 µM). Subsequently the tissues were incubated with the redox-sensitive ROS indicator H2DCFDA and ROS generation was monitored (Fig. 6AB, n = 6). Whereas, PPAR agonists significantly increased ROS generation, a significant downregulation of ROS generation was observed in the presence of the PPAR antagonist MK886. In contrast to the effects of PPAR agonists, no effects on ROS generation within embryoid bodies was observed upon stimulation of PPARβ and PPAR receptors. To investigate whether ROS generation induced by PPAR agonists was due to stimulation of NADPH oxidase, embryoid bodies were co-incubated with the NADPH oxidase inhibitors DPI (100 nM - 10 µM) and apocynin (10 µM) (Fig. 6C, n = 3). Both inhibitors of NADPH oxidase significantly reduced the increase in ROS generation achieved with WY14643, GW7647, and ciprofibrate suggesting NOX enzymes as the source of ROS generation. Furthermore, preincubation with the free radical scavengers vitamin E (trolox) (100 µM) and NMPG (100 µM) completely abolished ROS generation induced by the PPAR agonist WY14643 (Fig. 6D, n = 4). To exclude a contribution of the respiratory chain in PPAR-mediated ROS generation 4-day-old embryoid bodies were incubated for 24 hours with 2 µM rotenone which inhibits the mitochondrial electron transport complex I. As shown in Figure 6E, (n = 3) rotenone neither impaired the increase in ROS generation upon treatment with WY14643 nor inhibited the basal ROS generation in embryoid bodies which strongly suggests that the respiratory chain is not involved in ROS generation of differentiating ES cells.
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agonists stimulate cardiomyogenesis of ES cells by inducing ROS generation through NOX enzymes. The effect of PPAR agonists on cardiomyogenesis should therefore be consequently inhibited in the presence of free radical scavengers as well as inhibitors of NADPH oxidase. To investigate this issue embryoid bodies were treated from day 4 to day 10 of cell culture either in the presence of WY14643 (10 µM), WY14643 in the presence of vitamin E (100 µM) and NMPG (100 µM), or with free radical scavengers alone (Fig. 7A, 7B). In parallel embryoid bodies were treated with WY14643 in the presence of either DPI (100 nM), or apocynin (10 µM), or with NADPH oxidase inhibitors alone (Fig. 7C). In agreement with our working hypothesis, the effect of WY14643 on cardiomyogenesis as evaluated by counting spontaneous contracting foci of cardiomyocytes (see Fig. 7A, n = 4) or assessing the size of beating foci by immunohistochemical analysis of -actinin-positive cell areas (see Fig. 7B, n = 3), was totally abolished in the presence of free radical scavengers. Furthermore, cardiomyogenesis was significantly inhibited in the presence of DPI and apocynin (see Fig. 7C, n = 4), thus clearly indicating that PPAR agonists stimulate cardiomyogenesis of ES cells by a ROS-dependent mechanism that involves activity of NADPH oxidase.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionAcknowledgmentsReferences |
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agonists are stimulating cardiomyogenesis of ES cells comparably to a recent study by Ding et al. [9]. PPAR activation apparently induced the cardiogenic transcriptional program rather than promoting cardiac cell survival since WY14643 induced cardiogenic transcription factors prior to the appearance of spontaneously contracting (mature) cardiomyocytes in the cell culture. Previously it has been shown that inhibition of the respiratory chain in ES cells by antimycin A, totally abolished cardiomyogenesis, suggesting that proper mitochondrial function is a prerequisite for ES cell-derived cardiac differentiation [16]. The data of the present study demonstrate that PPAR stimulation resulted in generation of ROS presumably by NADPH oxidases, since pharmacological inhibitors abolished the response. A contribution of ROS generation by the respiratory chain could be excluded since the complex I inhibitor rotenone was without effects. This finding is in contrast to recent in vivo studies which showed that PPAR activation could diminish postischemic cardiomyocytic apoptosis and ROS-mediated injuries after global cardiac arrest under cardiopulmonary bypass, possibly via prevention of both caspase-dependent and caspase-independent apoptotic pathways [17]. On the other hand, it has been recently reported that activation of PPAR in mice leads to increased cardiac fatty acid oxidation and subsequent accumulation of oxidative stress intermediates in the heart, resulting in cardiomyocyte necrosis [18]. Furthermore, it has been shown that PPAR induces NADPH oxidase activity in macrophages [19]. In embryoid bodies derived from mouse ES cells macrophage differentiation has been demonstrated to occur on day 10 of cell culture [20], which is beyond the time frame of our experiments where incubations with PPAR agonists were undertaken between day 4 and day 10 in cell culture. Data from our group have recently shown that NOX-2, which is the NADPH oxidase isoform present in macrophages, is maximally expressed in embryoid bodies on day 12 of cell culture. NOX-2 expression is preceded by the transient expression of NOX-1 which has its maximum on day 8 of cell culture. NOX-4 was continuously expressed during the whole cell culture time but displayed elevated expression levels around day 15 of ES cell culture [13]. The peak in PPAR expression well coincided with the expression maximum of NOX-2, which is the NADPH isoform of macrophages. Since we found that the macrophage marker CD68 was expressed during this time, it may be assumed that PPAR-induced ROS generation may at least partially be mediated by macrophage activation. In the present study it is furthermore shown that the expression of the macrophage marker CD68 coincides with the expression of the cardiac marker -MHC suggesting that ROS derived from macrophages indeed may be involved in cardiac cell commitment. We can therefore not exclude that PPAR agonists induce differentiation of macrophages which may add to the global ROS generation observed by addition of PPAR agonists in the whole embryoid body tissue. Interestingly, ROS generation was only achieved with PPAR agonists, whereas the PPARβ agonist L-165,041 and the PPAR agonist GW1929 were without effect and consequently did not stimulate cardiomyogenesis of ES cells, even though our data demonstrate that PPAR as well as PPARβ and - are expressed in ES cells already at early stages of differentiation.
The involvement of ROS in cardiomyogenesis and cardiac cell proliferation of mouse ES cells has been extensively studied by us during recent years. It was demonstrated that low levels of ROS generated by either treatment of ES cells with electromagnetic fields, stimulation with the cardiogenic cytokine cardiotrophin-1 (CT-1) [21] or by exogenous addition of H2O2 to the cell culture medium [12, 13] significantly increased the expression of cardiac genes and transcription factors as well as the number and extension of spontaneously contracting cardiac foci. The effects were abolished in the presence of NADPH oxidase inhibitors, suggesting that the rapid elevation of ROS after stimulation was followed by a long-term enhancement of NADPH gene expression [13]. Besides stimulated ROS generation ES cells have been shown to endogenously generate ROS through NOX enzymes. This endogenous ROS generation apparently controls cardiomyogenesis since inactivation of NOX-4 by si-RNA technology totally abolished cardiomyogenesis of ES cells, thus suggesting that cardiac cell differentiation indeed requires NADPH oxidase activity [14]. The control of NOX enzyme expression and ROS generation during the differentiation of ES cells has not yet been unraveled. It may be assumed that during cardiovascular differentiation of ES cells, and with the onset of beating activity the energy demand of the growing tissue is increased thus initiating a switch from glycolytic ATP generation towards fatty acid oxidation. This switch in energy metabolism consequently requires increased expression and function of PPARs which may, besides their function in stimulating fatty acid metabolism, increase ROS generation thereby adjusting cardiomyogenesis and cardiac cell proliferation of ES cells to the increasing energy demand within the differentiating cardiac tissue. Whether a comparable mechanism of increased energy supply and cardiac hyperplasia exists in vivo remains elusive. Previous studies on PPAR homo-knockout (–/–) mice have demonstrated no severe impairment of viability and embryonic heart development, although these mice displayed reduced expression of lipid transporters, enzymes of β-oxidation as well as myocardial uptake and oxidation of long chain fatty acids. However, PPAR seemingly plays a role in cardiac function since PPAR–/– mice have an alteration of cardiac contractile performance under basal and under stimulation of β1-adrenergic receptors which are associated with myocardial fibrosis [22].
PPAR agonists are widely used in clinical practice for the treatment of dyslipidemia to reduce the cardiovascular risk. Furthermore, some studies suggested that PPAR ligands may be involved in the regulation of cardiac hypertrophy, inflammation, vascular function, and vascular remodelling, thus highlighting extra potential indications for these agents [2]. Since stem cell therapy within the heart appears to be a reasonable therapeutic regime to cure cardiovascular diseases in the future, a potential treatment of patients with PPAR agonists may be exploitable to enhance cardiomyogenic differentiation and cardiac cell proliferation within the area of transplantation, in parallel to exerting beneficial effects on the pathogenesis of heart failure [23], hypertrophy [24], and atherosclerosis [25].
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ACKNOWLEDGMENTS |
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