Arjunolic acid, a peroxisome proliferator-activated receptor α agonist, regresses cardiac fibrosis by inhibiting non-canonical TGF-β signaling

Abstract

Cardiac hypertrophy and associated heart fibrosis remain a major cause of death worldwide. Phytochemicals have gained attention as alternative therapeutics for managing cardiovascular diseases. These include the extract from the plant Terminalia arjuna, which is a popular cardioprotectant and may prevent or slow progression of pathological hypertrophy to heart failure. Here, we investigated the mode of action of a principal bioactive T. arjuna compound, arjunolic acid (AA), in ameliorating hemodynamic load-induced cardiac fibrosis and identified its intracellular target. Our data revealed that AA significantly represses collagen expression and improves cardiac function during hypertrophy. We found that AA binds to and stabilizes the ligand-binding domain of peroxisome proliferator-activated receptor α (PPARα) and increases its expression during cardiac hypertrophy. PPARα knockdown during AA treatment in hypertrophy samples, including angiotensin II-treated adult cardiac fibroblasts and renal artery-ligated rat heart, suggests that AA-driven cardioprotection primarily arises from PPARα agonism. Moreover, AA-induced PPARα up-regulation leads to repression of TGF-β signaling, specifically by inhibiting TGF-β-activated kinase1 (TAK1) phosphorylation. We observed that PPARα directly interacts with TAK1, predominantly via PPARα N-terminal transactivation domain (AF-1) thereby masking the TAK1 kinase domain. The AA-induced PPARα-bound TAK1 level thereby shows inverse correlation with the phosphorylation level of TAK1 and subsequent reduction in p38 MAPK and NF-κBp65 activation, ultimately culminating in amelioration of excess collagen synthesis in cardiac hypertrophy. In conclusion, our findings unravel the mechanism of AA action in regressing hypertrophy-associated cardiac fibrosis by assigning a role of AA as a PPARα agonist that inactivates non-canonical TGF-β signaling.

Keywords: TGF-β-activated kinase 1 (TAK1); angiotensin II; arjunolic acid; cardiac hypertrophy; collagen; fibroblast; fibrosis; heart failure; peroxisome proliferator-activated receptor α (PPARα); transforming growth factor-β (TGF-B).

Figure 1.

AA regresses collagen expression and improves cardiac function during cardiac hypertrophy. A, RT-PCR analyses showing significant increase in collagen-1 (col-1) and collagen-3 (col-3) gene expressions in AngII-treated adult cardiac fibroblasts in vitro compared with control fibroblasts. AA-infused AngII-treated cells showed significant decrease in col-1 and col-3 gene expressions compared with hypertrophied fibroblasts. Rpl-32 was used as internal loading control. Control cells treated with either DMSO or AA yielded similar results. AngII-treated cells were also treated with equivalent amounts of DMSO. n = 10 for each experimental group. All the results were expressed as ±S.E. of three independent experiments. Representative graphs showing relative alterations in collagen gene expressions among different experimental groups. ***, p < 0.001 with respect to DMSO-treated control cells; ###, p < 0.001 with respect to AngII-treated cells. B, RT-PCR analyses showing significant increase in col-1 and col-3 gene expressions in right renal artery-ligated rat heart compared with sham-operated control rat group. AA treatment in ligated animals showed significant decrease in col-1 and col-3 gene expressions compared with ligated animals. Rpl-32 was used as internal loading control. Sham-operated control animals and renal artery-ligated animals were also treated with equivalent amounts of DMSO. n = 7 for each experimental group. All the results were expressed as ±S.E. of three independent experiments. Representative graphs showing relative alterations in collagen gene expressions in different in vivo experimental groups. **, p < 0.01 with respect to sham-operated control rat group; ***, p < 0.001 with respect to sham-operated control rat group; ##, p < 0.01 with respect to renal artery-ligated rat group; ###, p < 0.001 with respect to renal artery-ligated rat group. C, graphical representation of in vitro hydroxyproline assay showing significantly increased collagen content in AngII-treated fibroblast culture supernatant compared with control cells. AA treatment in AngII-treated fibroblasts showing significant decrease in the collagen content in the culture supernatant compared with AngII-treated cells. Control cells treated with either DMSO or AA yielded similar results. AngII-treated cells were also treated with equivalent amounts of DMSO. n = 10 for each experimental group. Results were expressed as ±S.E. of three independent experiments. *, p < 0.05 with respect to control cells; #, p < 0.05 with respect to Ang II-treated cells. D, graphical representation of in vivo hydroxyproline assay showing significantly increased total left ventricular collagen content in ligated rat hearts compared with sham-operated control hearts. AA treatment in ligated rat hearts showed decreased total left ventricular collagen content compared with ligated rat heart. Equivalent amounts of DMSO were administered into sham-operated and renal artery-ligated rat group. n = 7 for each experimental group. Results were expressed as ±S.E. of three independent experiments. **, p < 0.01 with respect to sham-operated control rat group; #, p < 0.05 with respect to renal artery-ligated rat group. E, graphical representation of HW/BW ratio (in milligrams/gram) ratio showing significant increase in renal artery-ligated rat group compared with sham-operated control rat group. AA treatment in ligated animals showed significant down-regulation of HW/BW ratio compared with hypertrophied rat group. Sham-operated and renal artery-ligated rat group were also treated with equivalent amounts of DMSO. n = 7 for each experimental group. Results were expressed as ±S.E. of three independent experiments. **, p < 0.01 with respect to sham-operated control rat group; ##, p < 0.01 with respect to renal artery-ligated rat group. F, graphical representation of CSA measured from H&E-stained heart tissue sections showing significant up-regulation in renal artery-ligated rat heart compared with sham-operated control rat group. AA treatment in ligated animals showed significant down-regulation of CSA compared with ligated rat group. Sham-operated and ligated rats were also treated with equivalent amounts DMSO. n = 7 for each experimental group. Results were expressed as ±S.E. of three independent experiments. ***, p < 0.001 with respect to sham-operated control rat group; ###, p < 0.001 with respect to renal artery-ligated rat group. G, RT-PCR analyses showing significant increase in hypertrophy marker gene expressions, namely anf, β-mhc, and acta1 in renal artery-ligated rat hearts compared with sham-operated control hearts. AA treatment in ligated animals showed significant down-regulation of all these gene expressions compared with ligated rat hearts. Sham-operated control and renal artery-ligated animals were also treated with an equivalent amounts of DMSO. Rpl-32 was used as internal loading control. n = 7 for each experimental group. Results were expressed as ±S.E. of three independent experiments. Representative graphs showing relative alterations in the hypertrophy marker gene expressions among different experimental groups. ***, p < 0.001 with respect to sham-operated control rat group; **, p < 0.01 with respect to sham-operated control rat group; ###, p < 0.001 with respect to renal artery-ligated rat group; ##, p < 0.01 with respect to renal artery-ligated rat group. H, M-mode echocardiographic analyses from parasternal short axis view at papillary muscle level graphically representing decreased %FS and increased LvIDd in renal artery-ligated rat group compared with sham-operated control rat group. AA treatment in ligated animals showed restored %FS and lowered LvIDd in ligated rats compared with the hypertrophied rat group. Sham-operated control and renal artery-ligated animals were also treated with equivalent amounts of DMSO. n = 7 for each experimental group. Results were expressed as ±S.E. of three independent experiments. **, p < 0.01 with respect to sham-operated control rat group; ##, p < 0.01 with respect to renal artery-ligated rat group; #, p < 0.05 with respect to renal artery-ligated rat group.

Figure 2.

Analyses of interaction between AA and PPARα. A, fluorescence and CD titrations of AA with PPARα. Panel i, titration of 8 μm PPARα with increasing AA concentrations up to 55 μm, monitored by fluorescence spectroscopy. Panel ii, titration of 8 μm PPARα with increasing AA concentrations up to 55 μm, monitored by CD spectroscopy. All titrations were carried out in 20 mm Na₃(PO)₄, 150 nm NaCl, pH 8.0, buffer at 298 K. B, overlaid FTIR spectra of free AA, free PPARα, and AA-PPARα complex. C, thermal melting study. Panel i, FTS assay to determine melting temperature of PPARα alone (black) and in the presence of AA (red). Panel ii, CD thermal melting of PPARα alone (black) and in presence of AA (red) probed by monitoring changes θ₂₁₈ (α-helical transitions) and θ₂₂₂ (inset, β-transitions). Normalized data are plotted as percent unfolding induced by temperature increments. All experiments were carried out in 20 mm Na₃(PO)_4, 150 nm NaCl, pH 8.0, buffer at 298 K. D, tertiary structure alignment of PPARα (modeled, black), bound to AA (red), iloprost (green), TIPP-703 (blue), and fenofibrate (purple). Zoomed part shows superposition of AA with other agonists occupying identical ligand-binding pocket in PPARα. E, projections of MD simulations and thermal melting profiles for PPARα alone (black) and in bound state with AA (red). Panels i–ii, distributions of backbone RMSD and R_g values averaged over whole 100-ns trajectory for PPARα alone (panel i) and in the presence of AA (panel ii). Arrows indicate major areas of distribution over two components. Panel iii, RMSF analyses of PPARα alone and in the presence of AA averaged over last 10 ns of trajectory. Panel iv, H-bond network for whole trajectory showing persistence of H-bonds between PPARα and AA. Inset shows three H-bonds between AA and Thr-283, Leu-331, and Tyr-334 residues.

Figure 3.

AA treatment during hypertrophy increases PPARα expression in an autoregulatory loop. A, ChIP assay with anti-PPARα antibody followed by qRT-PCR analyses of the PPRE showing relative binding of PPARα to the PPRE within PPARα promoter among different experimental groups both in vitro and in vivo are represented graphically on logarithmic scale. Hypertrophy samples showed significantly down-regulated-fold enrichment in binding of PPARα to PPRE within the PPARα promoter compared with respective controls. AA treatment in hypertrophy samples further showed significantly increased-fold enrichment of the same, compared with hypertrophy samples. PPARα siRNA treatment in AA-treated hypertrophy groups showing down-regulated-fold enrichment in binding of PPARα to the PPRE compared with the AA-treated hypertrophy samples were used as negative controls. Control and hypertrophy samples in vitro and in vivo were treated with equivalent amounts of DMSO and non-specific (NS) siRNA. AA-treated hypertrophy samples were treated with equivalent amounts of NS siRNA. Chromatin from each experimental group immunoprecipitated with anti-IgG antibody were used for normalization. n = 3 both in vitro and in vivo. Results were analyzed by one-way ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. **, p < 0.01 with respect to control samples; ###, p < 0.001 with respect to hypertrophy samples; ↑↑, p < 0.01 with respect to AA-treated hypertrophy samples in vitro and/or in vivo. Corresponding changes in pparα mRNA expressions between different experimental groups as observed by qRT-PCR are represented graphically on logarithmic scale. Rpl-32 was used as internal loading control. n = 3 both in vitro and in vivo. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. **, p < 0.01 with respect to control samples; #, p < 0.05 with respect to hypertrophy samples; ##, p < 0.01 with respect to hypertrophy samples; ↑, p < 0.05 with respect to AA-treated hypertrophy samples; ↑↑, p < 0.01 with respect to AA-treated hypertrophy samples in vitro and/or in vivo. B, Western blot analyses revealed significant decrease in PPARα protein expression during hypertrophy compared with respective control groups that again showed significant recovery in AA-treated hypertrophy samples compared with respective hypertrophy groups in vitro and in vivo. Successful knockdown of PPARα protein expression was also confirmed by PPARα siRNA pretreatment in AA-treated hypertrophy samples. GAPDH was used as internal loading control. Control and hypertrophy samples were treated with equivalent amounts of DMSO and NS siRNA. AA-treated hypertrophy samples were also treated with NS siRNA. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 10 in vitro, n = 7 in vivo for each experimental group. Representative graphs showing relative changes in expression of PPARα among different experimental groups on logarithmic scale. **, p < 0.01 with respect to control samples; #, p < 0.05 with respect to hypertrophy samples; ##, p < 0.01 with respect to hypertrophy samples; ↑↑, p < 0.01 with respect to AA-treated hypertrophy samples in vitro and/or in vivo. C, Western blot analyses showing time-dependent increase in PPARα expression in AA-treated hypertrophied fibroblasts compared with AngII-treated cells at respective time points under study. GAPDH was used as internal loading control. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 5 for each experimental group. Representative graphs showing relative changes in PPARα expressions at respective time points among different groups under study. C, control fibroblasts; A, AngII-treated fibroblasts at different time points; A + AA, AngII-treated fibroblasts at different time points, treated along with AA. C and A cells were also treated with equivalent concentration of DMSO. ρρ, <0.01 with respect to A samples at the 3-h time point; øø, <0.01 with respect to A + AA samples at the 3-h time point.

Figure 4.

AA-mediated up-regulation of PPARα results in regression of cardiac fibrosis and improvement of cardiac function. A, graphical representation of qRT-PCR data showing significant down-regulation of col-1 and col-3 gene expressions in AA-treated hypertrophy samples compared with hypertrophy groups. PPARα siRNA-pretreated hypertrophied groups treated along with AA showed significant recovery in these gene expressions compared with AA-treated hypertrophy samples both in vitro and in vivo. Control and hypertrophy samples were treated with equivalent amounts of DMSO and NS siRNA. AA-treated hypertrophy samples were also treated with equivalent amounts of NS siRNA. Rpl-32 was used as internal reference control. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 10 in vitro and n = 7 in vivo for each experimental group. **, p < 0.01 with respect to control samples; ***, p < 0.001 with respect to control samples; ##, p < 0.01 with respect to hypertrophy samples; ↑, p < 0.05 with respect to AA-treated hypertrophy samples; ↑↑↑, p < 0.001 with respect to AA-treated hypertrophy samples in vitro and/or in vivo. B, graphical representation of in vitro hydroxyproline assay showing significantly restored collagen content in the culture supernatant of PPARα siRNA-pretreated hypertrophied fibroblasts treated along with AA compared with AA-treated hypertrophied cells in vitro. Control and AngII-treated cells were treated with equivalent amounts of DMSO and NS siRNA. AA-treated hypertrophied cells were also treated with equivalent amounts of NS siRNA. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 10 for each experimental group. **, p < 0.01 with respect to control cells; #, p < 0.05 with respect to AngII-treated cells; ↑, p < 0.05 with respect to AngII-treated cells treated along with AA. C, graphical representation of in vivo hydroxyproline assay showing significant recovery in total left ventricular collagen content in PPARα siRNA-pretreated AA-infused hypertrophied heart compared with AA-treated renal artery-ligated rat heart in vivo. Sham-operated control rats and renal artery-ligated rats were also treated with equivalent amounts of DMSO and NS siRNA. AA-treated ligated rats were also treated with equivalent amounts of NS siRNA. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 7 for each experimental group. **, p < 0.01 with respect to sham-operated control rat group; ##, p < 0.01 with respect to renal artery-ligated rat group; ↑, p < 0.05 with respect to AA-treated renal artery-ligated rat group. D, micrographs of Massons' trichrome staining showing significantly decreased %CVF in AA-treated hypertrophy samples compared with renal artery-ligated rat heart, which again showed significant recovery in AA-treated ligated samples pretreated with PPAR siRNA. S, sham-operated control group; L, ligated rat group; L + AA, AA-treated ligated rat group; L + AA + PPARα si, PPARα siRNA-infused AA-treated ligated rat group. S and L groups were treated with equivalent amounts of DMSO and NS siRNA. L + AA group animals were also treated with equivalent amounts of NS siRNA. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 7 for each group. (Scale bar, 40 μm, magnification = ×60). **, p < 0.01 with respect to sham-operated control samples; ##, p < 0.01 with respect to renal artery-ligated hypertrophy samples; ↑↑, p < 0.01 with respect to renal artery-ligated samples treated with AA. E, M-mode echocardiographic analyses of PPARα siRNA-pretreated ligated rats treated with AA showing significant decrease in %FS and significant increase in LvIDd compared with AA-treated ligated rats. Sham controls and renal artery-ligated rats were treated with equivalent amounts of DMSO and NS siRNA. AA-treated ligated rats were also treated with equivalent amounts of NS siRNA. n = 7 for each group. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. **, p < 0.01 with respect to sham-operated control rat group; ##, p < 0.01 with respect to renal artery-ligated rat group; ↑↑, p < 0.01 with respect to AA-treated renal artery-ligated rat group.

Figure 5.

AA-mediated up-regulation of PPARα specifically targets TAK1 for regression of TGF-β signaling during cardiac hypertrophy. A, Western blot analyses showing significantly reduced TβRI, TβRII, phospho/total levels of SMAD 2, SMAD 3, and TAK1 in AA-treated hypertrophy samples compared with hypertrophy groups both in vitro and in vivo. Significantly restored levels of all these proteins were observed in AA-treated hypertrophy groups pretreated with PPARα siRNA compared with AA-treated hypertrophy samples. Control and hypertrophy samples were treated with equivalent amounts of DMSO and NS siRNA. AA-treated hypertrophy samples were also treated with equivalent amounts of NS siRNA. GAPDH was used as internal loading control. n = 10 in vitro and n = 7 in vivo for each experimental group. B, Western blot analyses showing significantly decreased phospho/total TAK1 level during AA treatment in TGF-β-treated cardiac fibroblasts compared with only TGF-β-treated cells. PPARα knockdown in TGF-β- and AA-infused cells showed significant recovery in phospho/total TAK1 level compared with AA-treated fibroblasts pretreated with TGF-β. Expressions of TβRI, TβRII, and phospho/total levels of SMAD 2 and SMAD 3 remained unaltered in these experimental groups. Control and TGF-β-treated fibroblasts were also treated with equivalent amounts of DMSO and NS siRNA. AA-treated fibroblasts pretreated with TGF-β were also treated with equivalent amounts of NS siRNA. GAPDH was used as internal loading control. n = 5 for each group.

Figure 6.

AA-mediated regression of collagen gene expression involves PPARα-dependent inactivation of non-canonical TGF-β signaling. A, Western blot analyses showing significantly reduced phospho/total levels of IKKβ, NF-κBp65, p38 MAPK, and JNK in AA-treated hypertrophy samples compared with hypertrophy groups both in vitro and in vivo. Significantly restored levels of all these proteins were observed during PPARα knockdown in AA-treated hypertrophied groups compared with AA-treated hypertrophy samples. Control and hypertrophy groups were also treated with equivalent amounts of DMSO and NS siRNA. AA-treated hypertrophy samples were treated with equivalent amounts of NS siRNA. GAPDH was used as internal loading control. n = 10 in vitro, n = 7 in vivo for each experimental group. B, graphical representation of qRT-PCR analyses showing significant down-regulation of col-1 and col-3 gene expressions in AngII-induced fibroblasts during knockdown of either TAK1 or NF-κBp65 or p38 MAPK via specific siRNA treatments compared with hypertrophied fibroblasts. JNK-specific siRNA treatment in AngII-treated fibroblasts showed no significant regression of collagen gene expression compared with AngII-treated cells. Control and AngII-treated cells were also treated with equivalent amounts of NS siRNA. Rpl32 was used as internal reference control. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 5 for each group. **, p < 0.01 with respect to control cells; ***, p < 0.001 with respect to control cells; #, p < 0.05 with respect to AngII-treated cells; ##, p < 0.01 with respect to AngII-treated cells. C, dual-luciferase assay showing significant increase in the Col1a1 promoter activity in AngII-treated fibroblasts compared with control fibroblasts. AA treatment in hypertrophied fibroblasts showed significant reduction in Col1a1 promoter activity compared with AngII-treated fibroblasts that again showed significant restoration in AA-treated hypertrophied fibroblasts pretreated with PPARα siRNA. Reduced Col1a1 promoter activity shown in NF-κBp65 siRNA pretreated hypertrophied cells compared with AngII-treated cells was used as negative control. Control and AngII-treated cells were also treated with NS siRNA with or without DMSO. AA-treated hypertrophy samples were also treated with equivalent amounts of NS siRNA. Results were normalized by Renilla luciferase activity in all the treatment groups. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 5 for each group. **, p < 0.01 compared with DMSO and NS siRNA-treated control cells; ρρ, p < 0.01 compared with NS siRNA-treated control cells; ##, p < 0.01 compared with DMSO and NS siRNA-infused AngII-treated cells; ↑↑, p < 0.01 with respect to AA-treated hypertrophied cells pretreated with NS siRNA; йй, p < 0.01 with respect to AngII-treated cells pretreated with NS siRNA.

Figure 7.

Analyses of interaction of PPARα with TAK1. A, overall schematic representation of the docking simulation between predicted structures of full-length rat TAK1 (silver) and different domains of rat-PPARα (cyan, AF-1; orange, DBD; green, H + LBD) based on the best fit HADDOCK score. B, FRAP analysis showed a positive FRET efficiency between endogenous PPARα and TAK1 in cardiac fibroblasts. Cells were probed for endogenous PPARα and TAK1 expressions with respective primary antibodies and stained with PPARα-FITC (green) and TAK1-TRITC (red). TRITC was subjected to 50% photobleaching. n = 5. Panel i, prebleach donor; panel ii, postbleach donor; panel iii, delta donor; panel iv, prebleach acceptor; panel v, postbleach acceptor; panel vi, FRET efficiency. C, co-IP experiments were done by immunoprecipitating proteins with anti-PPARα antibody followed by immunoblotting with anti-TAK1 antibody in vitro. PPARα-overexpressed AngII-treated fibroblasts were used as a positive control. Normalization was done by Western blot with anti-PPARα antibody in the same samples. Control and AngII-treated cells were also treated with either DMSO or empty pCDNA6/V5-HisB vector yielding similar results. n = 5 for each group. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. Graph showing relative changes in the level of interaction between PPARα and TAK1 between different experimental groups. C, control fibroblasts; A, AngII-treated fibroblasts; A + AA, AA co-treated AngII infused fibroblasts; A + pOV, PPARα overexpressed AngII-treated fibroblasts. *, p < 0.01 with respect to control fibroblasts; ##, p < 0.01 with respect to AngII-treated fibroblasts.

Figure 8.

Study of the effect of different PPARα domains upon PPARα-TAK1 interaction and their roles in modulation of non-canonical TGF-β pathway-induced collagen synthesis in hypertrophied fibroblasts. A, co-IP experiments were done by immunoprecipitating proteins with anti-His antibody followed by immunoblotting with anti-TAK1 antibody in vitro. Normalization was done by immunoblotting with anti-His antibody. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. n = 5 for each group. E, empty pCDNA6/V5 HisB vector; F, full-length PPARα plasmid; AF-1, PPARα N-terminal transactivation domain plasmid; DBD, PPARα DNA-binding domain; H + LBD, PPARα combined Hinge region+ C-terminal ligand-binding domain plasmid. All the plasmids were transfected into AngII-treated fibroblasts. n = 5 for each group. B, Western blot (WB) analyses showing alterations in expression levels of phospho/total TAK1, NF-κBp65, and p38 MAPK in AngII-treated cells transfected with full-length or individual domains of PPARα plasmids compared with AngII-treated fibroblasts. GAPDH was used as internal loading control. C, control fibroblasts; A, AngII-treated fibroblast; A + F, full-length PPARα transfected AngII-treated fibroblasts; A + AF-1, PPARα N-terminal transactivation domain transfected AngII-treated cells; A + DBD, PPARα DNA-binding domain transfected AngII-treated cells; A + H + LBD, PPARα combined Hinge region+ C-terminal Ligand-binding domain transfected AngII-treated cells. C and A cells were also treated with empty pCDNA6/V5 HisB vector. n = 5 for each group. C, graphical representation of qRT-PCR analyses showing changes in levels of col-1 and col-3 gene expressions in AngII-treated cells transfected with full-length or individual domains of PPARα plasmids compared with AngII-treated fibroblasts. Rpl-32 was used as internal loading control. Results were analyzed by ANOVA followed by Tukey's post hoc test and expressed as ±S.E. of three independent experiments. C, control fibroblasts; A, AngII-treated fibroblast; A + F, full-length PPARα transfected AngII-treated cells; A + AF-1, PPARα N-terminal transactivation domain transfected AngII-treated cells; A + DBD, PPARα DNA-binding domain transfected AngII-treated cells; A + H + LBD, PPARα combined Hinge region+ C-terminal ligand-binding domain transfected AngII-treated cells. C and A cells were also treated with empty pCDNA6/V5 HisB vector. n = 5 for each group. **, p < 0.01 with respect to control cells; ***, p < 0.001 with respect to C; ##, p < 0.01 with respect to A; ###, p < 0.001 with respect to A; ϕϕ, p < 0.01 with respect to A + F; ϕϕϕ, p < 0.001 with respect to A + F.

Figure 9.

Schematic representation of the molecular mechanism of AA action upon cardiac hypertrophy-associated fibrosis. A, during cardiac hypertrophy TGF-β signaling pathway action is promoted leading to excess collagen synthesis with down-regulated PPARα expression. Treatment with AA in hypertrophy samples increases PPARα expression in an autoregulatory loop leading to increased binding of PPARα to TAK1 thereby ameliorating TAK1-driven non-canonical TGF-β axes with subsequent regression of collagen synthesis. B, schematic representation of different PPARα domains interacting with TAK1 and the role of PPARα-TAK1 interaction in prevention of phosphorylation-dependent activation of TAK1 for subsequent regression of collagen synthesis in AngII-treated adult cardiac fibroblasts.