Cell Culture and Transfection

10T1/2 cells were grown in MEM Eagles medium in 6-well plates and transiently transfected with 500 ng of empty pcDNA3.1 control or pcDNA3.1-MRTF-A plasmid for 48 hours before RNA isolation. CFs grown in DMEM were treated as noted in the figure legends for 24 to 48 hours, followed by RNA or protein isolation. CF proliferation was determined with the CellTiter 96 Aqueous One Solution Cell Proliferation Assay (Promega) using the protocol of the manufacturer.

For luciferase assays, CFs or COS cells cultured in 24-well plates were transfected with a total of 300 ng of plasmid DNA using FuGENE6 (Roche). pCMV-lacZ (20 ng) was used as an internal control, and total plasmid amount was kept constant using empty pcDNA3.1. Forty-eight hours after transfection luciferase and β-galactosidase assays were carried using a luciferase assay kit (Promega).

Col1a2 Reporter Construction

The mouse Col1a2 promoter (420 bp) was amplified using high-fidelity Taq polymerase (TAKARA), and the following oligonucleotides containing 5′ KpnI and 3′ XhoI linkers: Col1a2 forward, 5′-GGTACCGACAGCTCCTGCCTTTTCATC-3′; Col1a2 reverse, 5′-CTCGAGTAAAATAATAAAGCCCAGACC-3′.

The resulting PCR product was cloned into the KpnI and XhoI sites of the pGL3-basic luciferase vector (Invitrogen). Mutation of the CArG element was accomplished using the QuickChangeII site-directed mutagenesis kit and the protocol of the manufacturer (Stratagene). The oligonucleotides used for PCR amplification were as follows: Col1a2 mutCArG forward, 5′-CCTAAAGTGCTTACACACGTGGCAAGGGCG-3′; and Col1a2 mutCArG reverse, 5′-CGCCCTTGCCACGTGTGTAAGCACTTTAGG-3′.

All constructs were sequence-verified.

Immunocytochemistry

CFs were infected with a flag-tagged MRTF-A adenovirus at a multiplicity of infection of 10 and were treated with TGF-β1 (10 ng/mL) and/or Y-27632 (10 μmol/L) 24 hours before fixation with cold methanol. Indirect immunofluorescence was performed with a mouse monoclonal Flag M2 antibody (Sigma) or Cy3-conjugated anti-SMA antibody (Sigma, clone 1A4, 1:200). Confocal images were captured using a Zeiss LSM-510 microscope.

Western Blot

Antibodies directed against SMA (Sigma), SM22 (Abcam), collagen I (Abcam), and tubulin (Sigma) were used to determine protein levels by Western blot. Nondenaturing PAGE was performed to detect collagen I protein by Western blot.

Histology and Immunohistochemistry

Tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned at 5-μm intervals. Hematoxylin/eosin and Masson’s trichrome staining were performed using standard procedures. SMA staining was performed on paraffin-embedded sections using a Cy3-conjugated anti-SMA antibody (Sigma, clone 1A4, 1:200). Nuclei were visualized using DAPI in Vectashield mounting medium (Vector laboratories). SMA-positive vessels and myofibroblasts were counted in the BZ of 3 WT and 3 MRTF-A^−/− animals and represented as the average±SEM. Proliferation and cell death was detected using a phospho–histone H3 antibody or the In Situ Cell Death Detection Kit and the protocol of the manufacturer (Roche), respectively.

RNA Analyses

Total RNA was isolated using TRIzol reagent (Invitrogen) according to the protocol of the manufacturer. RNA (2 μg) was used to generate cDNA using Superscript III (Invitrogen) according to the protocol of the manufacturer and detected using TaqMan primer and probe sets.

Collagen Synthesis Assay

[³H]Proline incorporation was performed to determine the effects of MRTF-A overexpression on collagen synthesis. CFs at passage 2 or 3 were cultured in 24-well tissue culture dishes for 48 hours, or until confluent, in DMEM supplemented with 10% FBS under standard culture conditions. CFs were then made quiescent by serum starvation for 24 hours, infected with 10 multiplicities of infection of adenovirus mediating the expression of MRTF-A or control β-galactosidase, and cultured in serum-free conditions for an additional 24 hours. CFs were then stimulated with the addition of 2.5% FBS, TGF-β1 (10 ng/mL), Y-27632 (10 μmol/L), or a combination of TGF-β1 and Y-27632 for 48 hours in the presence of [³H]proline (1 μCi/mL, PerkinElmer Life Sciences). CFs were then washed 3 times with Dulbelcco’s PBS, and protein was precipitated with ice cold 5% TCA for 1 hour. The precipitate was then solubilized with 400 μL of 0.2 mol/L NaOH at 37°C for 30 minutes. Radioactivity was determined by liquid scintillation counting. Each condition was performed in quadruplicate and repeated in 3 independent experiments.

Chromatin Immunoprecipitation Assay

Chromatin immunoprecipitation was performed using the EZ-ChIP kit (Millipore) using the instructions of the manufacturer. Briefly, native chromatin from 10T1/2 cells was crosslinked and immunoprecipitated with antibodies directed against SRF (Santa Cruz Biotechnology), RNA PolII (Millipore), or mouse IgG (Millipore). Col1a2 promoter sequences or GAPDH was detected using PCR amplification.

Electrophoretic Mobility-Shift Assay

Electrophoretic mobility-shift assay was performed using double-stranded oligonucleotides corresponding to the Col1a2 CArG sequence. Protein lysate (4 μL) from flag-SRF or empty pcDNA3.1 transfected COS cells was incubated with ³²P-labeled oligonucleotide probes in the presence of 1 μL of poly(dI-dC) (1.0 μg/μL) for 20 minutes at room temperature. Supershift formation was detected by adding 2 μL of anti-Flag M2 antibody (Sigma).

Myocardial Infarction

MIs were generated using male MRTF-A^−/− and WT mice at 12 weeks of age (25 to 30 g) by surgical ligation of the left anterior descending coronary artery (LAD). Sham-operated mice underwent the same procedure without occlusion of the LAD. For determination of infarct size, at least 4 images of trichrome-stained sections per heart were imported to OpenLab 3.1 and the area of trichrome staining was measured and taken as a percentage of the total left ventricular (LV) area in each section. For the studies designed to measure area at risk (AAR) for infarct, 0.3% methylene blue dye was perfused throughout the animal using direct LV administration immediately following confirmation of myocardial ischemia. Perfusion was carried out until significant staining of cardiac tissue had occurred and no further increase in stained area was apparent for 1 minute. The mouse was then further perfused with 4% paraformaldehyde in saline to ensure the proper fixation of tissue and vital dye. The heart was then collected and analyzed for AAR. The heart was photographed in whole mount to document the size of stained (perfused) versus unstained (unperfused) regions. The heart was then histologically dissected into 3 equal-sized transverse sections, starting at the site of ligature and progressing toward the apex of the heart. Stained LV tissue was separated from unstained tissue and weighed. The proportion of unstained versus stained tissue based on dry weight determined the AAR.

Angiotensin II Infusion

Ang II (dissolved in 0.01 mol/L acetic acid) was subcutaneously infused at the rate of 0.6 mg/kg per day for 2 weeks using an osmotic minipump (Alzet model 2002; URECT Corp, Cupertino, Calif) implanted in each mouse. After 2 weeks of Ang II infusion, left ventricles were then fixed in 10% formaldehyde. To determine the extent of collagen fiber accumulation, we randomly selected fields and measured the Masson’s trichrome–stained interstitial fibrosis area in relation to the total LV area using microscopy BIOREVO BZ-9000 (Keyence, Osaka, Japan). Perivascular fibrosis area was excluded in the present study.

Data and Statistical Analysis

Results are presented as means±SEM unless otherwise stated. Statistical analysis of group differences was performed by Student’s 2-tailed t test with unequal variance, and significance between groups of percentage fractional shortening was performed using multiple measures 2-factor ANOVA. Significance was considered as P<0.05.

Mouse Mutants and Animal Care

All experiments using animals were previously approved by the Institutional Animal Care and Use Committee at UT Southwestern Medical Center. The MRTF-A^−/− mouse line used in this study has been previously reported.¹⁹ Mice were genotyped using previously described PCR strategies.

MRTF-A^−/− Mice Display Reduced Cardiac Fibrosis After MI

ROCK signaling has been implicated in myofibroblast activation in diseases associated with excessive fibrosis, including cardiac remodeling following MI.^32,33 In light of the role of MRTF-A as a mediator of ROCK signaling and stress-dependent gene expression in cultured cells,^21,22 we investigated the potential involvement of MRTF-A in the response of the heart to MI, by surgical ligation of the LAD in wild-type (WT) and MRTF-A^−/− mice. Two weeks following ligation, WT mice developed an extensive fibrotic scar that spanned the majority of the LV free wall (Figure 1A, a1 and a2), as visualized grossly and by Masson’s trichrome staining of histological sections. The infarcted region typically displayed significant thinning and dilatation in association with fibrosis. In contrast, we observed a pronounced reduction in infarct size in MRTF-A^−/− hearts, as assessed by the size of the fibrotic scar (Figure 1A, b1 and b2). MRTF-A deletion resulted in an ≈50% reduction in scar size, as quantified by trichrome staining/LV ratio (Figure 1B). Post-MI lethality was nearly identical between WT and MRTF-A^−/− mice (data not shown), implying adequate initial scar formation to allow wound healing and prevent cardiac rupture. Assessment of cardiac function at baseline and at 3, 7, and 14 days after MI revealed that although MRTF-A^−/− mice tended toward improved percentage fractional shortening compared to WT mice, this improvement did not reach statistical significance (Online Figure I, available at http://circres.ahajournals.org).

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Figure 1. MRTF-A deletion results in reduced scar formation following MI. A, Whole-mount images of representative hearts 14 days after MI and Masson’s trichrome staining of sections from 2 representative hearts. Arrow denotes point of ligature, and plane of section is illustrated by horizontal line. Masson’s trichrome staining illustrates reduced and more compact region of scarring in the infarct zone (compare a1 and a2 to b1 and b2). RV indicates right ventricle; LV, left ventricle. Scale bar, 1 mm. B, Quantification of infarct size presented as the percentage of LV area positively stained with Masson’s trichrome (n=8 WT and 8 knockout [KO] animals). C, AAR determined by perfusion with methylene blue. AAR is devoid of staining. Arrow denotes point of ligature on the LAD. D, Quantification of AAR presented as percentage of LV mass that is not stained 10 minutes and 1 day after MI (n=4 WT and 4 MRTF-A^−/− animals at 10 minutes and 3 WT and 3 MRTF-A^−/− animals 1 day after MI). E, Quantification of TUNEL-positive cells 1 day after MI was performed on at least 4 independent fields within the infarct zone of each heart and averaged from 2 WT and 3 MRTF-A^−/− animals. Data are represented as percentages of DAPI-stained nuclei positive for TUNEL. F, Quantification of phospho-histone H3–positive cells 14 days after MI was performed on at least 7 independent fields in the BZ from each heart and averaged from 3 WT and 4 MRTF-A^−/− animals. Error bars indicate the SEM.

The reduction of infarct size in MRTF-A^−/− mice could, in principal, result from a reduced propensity to generate an infarct or an increased capacity for healing and regeneration of healthy cardiac tissue. To address these possibilities, we first determined the size of the AAR for infarction in WT and MRTF-A^−/− mice by perfusing animals with methylene blue. Gross examination of hearts immediately after ligation revealed a similar area of perfusion in WT and MRTF-A^−/− mice (Figure 1C). Quantification of the mass of nonperfused versus perfused myocardial tissue confirmed that deletion of MRTF-A did not result in a reduction of the AAR (Figure 1D). The AAR was also not altered in MRTF-A^−/− mice 24 hours after MI (Figure 1D). These results suggest the collateral vessel architecture is not significantly affected by the absence of MRTF-A.

We next determined whether MRTF-A deletion affected cell death in the infarct zone 24 hours after MI. Staining of histological sections for TUNEL revealed significant cell death throughout the infarcted region, although the percentage of TUNEL-positive nuclei was nearly identical between WT and MRTF-A^−/− animals (Figure 1E and Online Figure II). Finally, to examine the possibility that hearts of MRTF-A^−/− mice might display increased propensity toward regeneration, we determined whether MRTF-A^−/− cardiac myocytes (CMCs) underwent reentry into the cell cycle. Immunostaining of hearts from WT or MRTF-A^−/− mice 14 days after MI for phospho–histone H3 and the CMC marker α-actinin did not reveal a significant alteration in proliferating CMCs (Figure 1F and Online Figure III). There was, however, a trend toward decreased proliferation of non-CMCs in the hearts of MRTF-A^−/− animals (Figure 1F and Online Figure III). We conclude that reduced infarct size observed in MRTF-A^−/− mice is not a consequence of alterations in AAR, cell viability, or regenerative capacity and may reflect a direct role in remodeling and scar formation.

MRTF-A Regulates Collagen Expression After MI

We isolated tissue from the infarct BZ and remote healthy tissue 14 days after MI for quantitative real-time PCR. Sham-operated mice were used as controls. Multiple markers of the ECM and fibrotic remodeling were elevated in the BZ of WT animals, as expected (Figure 2). Upregulation of Col1a1, Col1a2, Col3a1, and elastin (Eln) in the BZ of WT mice was attenuated in the BZ of MRTF-A^−/− mice, as demonstrated by real-time RT-PCR (Figure 2A). The pronounced diminution of these markers in MRTF-A^−/− mice (Figure 2A) is consistent with the reduced injury in these animals (Figure 1A and 1B). The reduction of fibrosis following MI was not accompanied by a significantly diminished expression of TGF-β1, -2, and -3 in the BZ suggesting normal cytokine activation (Figure 2B). Expression of atrial natriuretic factor was reduced in the BZ of MRTF-A^−/− mice, further indicating a decrease of pathological remodeling (Figure 2C) although tenascin C, a marker of cardiac repair after MI, was similarly induced in the BZ of WT and MRTF-A^−/− mice (Figure 2C). The close correlation between the induction of collagen gene expression following MI and MRTF-A genotype implicates MRTF-A in the promotion of scar formation following MI.

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Figure 2. MRTF-A influences ECM gene expression 14 days after MI. A, Real-time PCR for various ECM components in sham hearts (S) and the BZ or remote (R) region of the infarct reveals attenuated expression of collagen genes in the BZ of MRTF-A^−/− hearts. B, The cytokines TGF-β1, -2, and -3 do not display significant alterations in expression in the BZ between WT and MRTF-A^−/− hearts. C, The stress-responsive atrial natriuretic factor (ANF) gene displays reduced expression in MRTF-A^−/− hearts. Tenascin C (TnC) levels are elevated to similar levels in the BZ of both WT and MRTF-A^−/− animals (n=4 WT and 4 MRTF-A^−/− hearts for infarct groups and 2 shams). Error bars represent SEM.

MRTF-A Regulates Myofibroblast Expression of SMC Markers Following MI

Because the myofibroblast is a primary contributor to ECM deposition and scar formation following MI,¹ we examined the expression of smooth muscle markers of myofibroblast activation in 14-day post-MI hearts. Expression of SM22 and SMA was attenuated in the BZ and remote tissue of MRTF-A^−/− animals 14 days after MI, as assessed by quantitative RT-PCR (Figure 3A). We next determined the localization of SMA in WT and MRTF-A^−/− hearts 14 days after MI by immunohistochemistry (Figure 3B). Immunostaining of histological sections for SMA revealed that the BZ of WT mice contained numerous spindle-shaped SMA-positive cells, or aggregates of SMA-positive cells not associated with a vessel, which are indicative of myofibroblast-like cells (Figure 3B, a"). In contrast, the BZ of MRTF-A^−/− mice possessed significantly fewer SMA-positive myofibroblasts than WT animals (Figure 3B, b"). Quantification of SMA-positive myofibroblasts in the BZ demonstrated a higher density (≈3-fold) in WT than in MRTF-A^−/− animals (Figure 3C). SMA is also highly expressed in arterioles that infiltrate the BZ of the infarct. Quantification of the number of SMA-positive arterioles in the BZ of WT and MRTF-A^−/− hearts revealed a slight but insignificant increase in the number of arterioles in MRTF-A^−/− animals (Figure 3D).

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Figure 3. MRTF-A deletion results in altered smooth muscle gene expression 14 days after MI. A, Real-time PCR reveals a significant reduction of SM22 and SMA induction in both the BZ and remote region of MRTF-A^−/− hearts. *P<0.01; †P<0.05 (n=4 WT and 4 MRTF-A^−/− hearts per infarct group and 2 shams). B, Histological sections of hearts from WT and MRTF-A^−/− mice, 14 days after MI are shown. Hematoxylin/eosin-stained (a and b) sections of representative hearts adjacent to those used for immunohistochemistry illustrate ischemic damage and highlight the BZ of the infarct. a′ and b′, Magnification of region that is boxed in a and b. a″ and b″, SMA immunostaining corresponding to the boxed region of representative WT and MRTF-A^−/− infarcted hearts. Arrows mark SMA-positive spindle-shaped myofibroblasts. Scale bars, 1 mm (a and b); 40 μm (a′, a″, and b′, b"). C, Number of SMA-positive myofibroblasts per field of view. D, Number of SMA positive arterioles in the BZ per field of view. Quantification was performed at ×40 magnification on at least 3 fields of view within the BZ of each heart and averaged from 3 WT and 3 MRTF-A^−/− hearts. *P<0.05). Error bars represent SEM.

MRTF-A^−/− Mice Display Reduced Fibrosis in Response to Ang II

Because MRTF-A^−/− mice are protected from excessive scar formation following MI, and the renin–angiotensin system is a major mediator of post-MI fibrosis, we asked whether MRTF-A also played a role in Ang II–mediated fibrosis. Following 14 days of Ang II infusion (0.6 mg/kg per day), WT mice displayed profound interstitial fibrosis, as assessed by Masson’s trichrome staining of histological sections through the LV (Figure 4A). In contrast, MRTF-A^−/− littermates were protected from fibrosis in response to Ang II infusion (Figure 4A). Quantification of the percentage LV area stained for Masson’s trichrome following Ang II treatment revealed a nearly complete protection from fibrosis on MRTF-A deletion (Figure 4B).

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Figure 4. MRTF-A deletion attenuates fibrosis following chronic Ang II administration. A, Masson’s trichrome staining of representative histological sections following 14 days of vehicle (a and c) or Ang II (b and d) administration in WT (a and b) or MRTF-A^−/− (c and d) mice. B, Quantification of trichrome staining of the left ventricle (LV). Error bars represent SEM. *P<0.05. C, Quantitative RT-PCR for collagen genes and smooth muscle differentiation markers reveals attenuated induction of Col1a, Col3a, SMA, and SM22 following Ang II administration of MRTF-A^−/− animals. Error bars represent SEM. *P<0.05.

Quantification of the expression of collagen genes after 14 days of Ang II treatment revealed enrichment of Col1a2 and Col3a1 expression in Ang II–treated WT mice, which was significantly attenuated in MRTF-A^−/− mice (Figure 4C), confirming the reduction of fibrosis observed in these animals. Likewise, the stimulation of SMA and SM22 expression seen in WT animals was not observed in MRTF-A^−/− mice (Figure 4C). These results further suggest a role for MRTF-A in promoting a myofibroblast phenotype and fibrotic remodeling in the heart.

Regulation of a Myofibroblast Phenotype by MRTF-A

To test whether MRTF-A was sufficient to induce myofibroblast-associated genes that were downregulated in MRTF-A^−/− mice, we enforced the expression of MRTF-A in cultured primary ventricular neonatal CFs by adenoviral-mediated expression. In contrast to myocardin, which is specifically expressed in the CMCs of the heart, MRTF-A is expressed in both CMCs and CFs (Figure 5A). As shown in Figure 5, overexpression of MRTF-A in CFs resulted in a dramatic increase in the expression of SMA and SM22 (Figure 5B). Immunocytochemical detection of SMA in cultured CFs confirmed the enrichment of SMA by MRTF-A, compared with β-gal–infected CFs (Figure 5C). MRTF-A overexpression in CFs resulted in the accumulation of SMA into highly organized stress fibers, a hallmark of myofibroblast activation, in contrast to being primarily localized to cortical actin in control cells (Figure 5C).

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Figure 5. MRTF-A regulates the expression of smooth muscle markers in CFs. A, Quantitative real-time PCR reveals myocardin is exclusively expressed in CMCs, whereas MRTF-A is present in both CMCs and CFs at similar levels. B, Quantitative real-time RT-PCR reveals SMA and SM22 are significantly enriched in CFs overexpressing MRTF-A relative to β-gal–infected control CFs. Error bars represent SEM. C, Indirect immunofluorescence of CFs demonstrates MRTF-A overexpression results in the enrichment of SMA into organized stress fibers as compared with Ad-β-gal–infected CFs, which display primarily cortical actin SMA staining. Scale bar, 25 μm.

TGF-β1 Promotes the Nuclear Translocation of MRTF-A in a ROCK-Dependent Manner

TGF-β1 promotes a contractile myofibroblast phenotype in multiple organs including the heart, kidneys, liver, and skin. TGF-β1 induced a robust elevation in SMA immunostaining in CFs, which was localized primarily to stress fibers (Figure 6A). The ROCK inhibitor Y-27632 largely blocked the induction of SMA by TGF-β1 and MRTF-A, while modestly inhibiting SMA staining at baseline (Figure 6A). Because MRTF-A undergoes nuclear translocation in response to TGF-β1 and ROCK signaling in kidney epithelial cells,^36,39 we examined whether TGF-β1 and ROCK could influence the activity of MRTF-A in CFs. MRTF-A displayed predominantly cytoplasmic localization in CFs cultured in serum-free media or in the presence of Y-27632 (≈10% to 20% nuclear; Figure 6B and 6C). Treatment of CFs with TGF-β1 resulted in a shift of MRTF-A to the nucleus; the number of cells displaying cytoplasmic restriction of MRTF-A was significantly reduced with a larger proportion displaying nuclear and cytoplasmic staining or nuclear enrichment (≈50% nuclear; Figure 6B and 6C). Y-27632 partially blunted the nuclear accumulation of MRTF-A in response to TGF-β1, suggesting that TGF-β1–dependent nuclear translocation is also influenced by ROCK activity (≈40% nuclear; Figure 6B and 6C).

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Figure 6. Regulation of MRTF-A nuclear localization and activity in CFs by TGF-β1 and the ROCK inhibitor Y-27632. A, Immunocytochemistry for SMA in CFs grown in serum-free (SF) media or media supplemented with TGF-β1 (10 ng/mL), Y-27632 (10 μmol/L), or both TGF-β1 and Y-27632. All images were captured using identical exposure settings. Scale bar, 25 μm. B, Immunocytochemical detection of Flag-tagged MRTF-A in CFs after 24 hours of growth in serum-free media or media supplemented with TGF-β1, Y-27632, or both TGF-β1 and Y-27632. Scale bar, 100 μm. B, Quantification of the subcellular localization of Flag-MRTF-A under various culture conditions. Subcellular localization of Flag-MRTF-A was scored for ≈20 random fields of view for each condition. Error bars represent SEM. *P<0.01; †P<0.05.

Col1a2 Is a Direct Target of MRTF-A

Because MRTF-A induced the expression of SMC markers indicative of a myofibroblast phenotype, we next assessed the ability of MRTF-A to influence the deposition of collagen by CFs, which largely mediate the fibrotic response following MI. We used [³H]proline incorporation to quantify collagen synthesis by CFs subjected to various stimuli. Importantly, MRTF-A overexpression did not stimulate CF proliferation (Online Figure IV), consistent with reports implicating SRF and myocardin in promotion of differentiation and inhibition of proliferation in CMCs.⁴⁰ Treatment of CFs with serum or TGF-β1 resulted in elevated collagen production (Figure 7A). MRTF-A overexpression in CFs also resulted in a significant elevation in collagen synthesis, and this increase was further stimulated by serum or TGF-β1 (Figure 7A). In contrast, inhibition of Rho signaling with Y-27632 resulted in the diminution of MRTF-A–dependent collagen synthesis (Figure 7A). We also detected increased levels of Col1a2, SMA, and SM22 protein in CFs overexpressing MRTF-A (Online Figure V). Col1a2 mRNA levels were also increased in response to MRTF-A overexpression, as revealed by quantitative Real Time RT-PCR (Figure 7B).

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Figure 7. MRTF-A regulates Col1a2 expression. A, [³H]proline incorporation in CFs demonstrates MRTF-A significantly enriches collagen synthesis. CFs were serum-starved for 48 hours and then infected with 10 multiplicities of infection of Ad-MRTF-A or Ad-β-gal and cultured in serum-free (SF) media or media supplemented with 2.5% FBS, TGF-β1 (10 ng/mL), Y-27632 (10 μmol/L), or both TGF-β1 and Y-27632 for an additional 48 hours. Error bars represent SEM. B, Quantitative real-time RT-PCR demonstrating the expression of Col1a2, Col3a1, SMA, and SM22 in 10T1/2 cells transfected with empty vector control or MRTF-A expression vector. C, Depiction of the region used in transient transfection assays and electrophoretic mobility-shift assays are shown. Blue peaks denote evolutionary conservation, and alignment of mammalian regulatory sequences are at the bottom, highlighting the conserved CArG, SP1, and Smad sites. CArG mutation used in electrophoretic mobility-shift assay and transient transfection assays is aligned with WT CArG. D, Chromatin immunoprecipitation of Col1a2 or GAPDH promoter sequences with an antibody directed against endogenous SRF. Antibodies directed against PolII or IgG serve as positive and negative controls. Input is 1% of total chromatin. E, Electrophoretic mobility-shift assay demonstrates that SRF binds to the conserved CArG box. Flag antibody supershifts the SRF/DNA complex, whereas WT unlabeled competitor oligonucleotide abolishes the shifted complex. Mutant oligonucleotide (m) fails to bind SRF and unlabeled mutant competitor (m) fails to abolish SRF/CArG interaction. F, Transient transfection of COS cells with a WT or CArG mutant Col1a2-luciferase construct reveals dose-dependent responsiveness to MRTF-A. Error bars represent SD. G, MRTF-A induces the expression of Col1a2-luc in transiently transfected CFs and displays increased activity in the presence of 10% FBS. Error bars represent SD. H, MRTF-A dependent Col1a2 promoter activity is stimulated by TGF-β1 treatment. Mutation of the CArG box abrogates responsiveness of the promoter to MRTF-A or TGF-β1 treatment. Error bars represent SD.

An evolutionarily conserved CArG box exists within the previously characterized Smad3- and Sp1-dependent promoter region of the Col1a2 gene (Figure 7C).⁴¹ An antibody directed against the endogenous SRF protein precipitated chromatin containing the Col1a2 CArG box in a TGF-β1–independent manner (Figure 7D). SRF also efficiently bound to the Col1a2 CArG box in gel mobility-shift assays (Figure 7E). A 420bp Col1a2 promoter fragment linked to a luciferase reporter was activated in a dose-dependent manner by MRTF-A, and mutation of the CArG box attenuated MRTF-A responsiveness (Figure 7F). Furthermore, the Col1a2 promoter displayed a dose-dependent induction by MRTF-A in primary CFs (Figure 7G), and mutation of the CArG box completely abolished the stimulation of MRTF-A activity by TGF-β1 (Figure 7H). We conclude that MRTF-A directly regulates Col1a2 gene expression to promote fibrosis and scar formation following MI.

Stress-Responsive Regulation of Collagens by MRTF-A

Upregulation of multiple markers of the ECM, including Col1a1, Col1a2, Col3a1, and elastin, was dramatically attenuated in MRTF-A^−/− mice following MI or Ang II treatment. We identify the Col1a2 promoter as a novel target of MRTF-A/SRF. We have also identified an evolutionarily conserved CArG box upstream of the transcriptional start site of the Col3a1 gene, adjacent to previously characterized regulatory elements.⁴² These findings suggest a potential role for MRTF-A in the direct regulation of a battery of fibrosis-associated genes in addition to previously defined targets associated with the actin cytoskeleton or smooth muscle sarcomeric organization.

Interestingly, the Col1a2 promoter contains a conserved CArG box harboring a G/C substitution in the A/T rich core. This type of CArG degeneracy diminishes binding affinity for free SRF, resulting in a low level of basal expression.⁴³ The results of our study support the hypothesis that CArG degeneracy may be important for stress-responsive activation of gene expression on nuclear accumulation of SRF cofactors such as MRTF-A.

MRTF-A Activation and TGF-β1

TGF-β1–Smad signaling is the best-characterized contributor to myofibroblast activation and fibrosis.^44–47 TGF-β1 and its transcriptional mediators, Smad2, 3, and 4, are activated following MI.^46–48 The TGF-β1–Smad3 pathway is a major mediator of post-MI remodeling, including the induction of SMA and SM22-enriched myofibroblasts and the transition to fibrosis.^44,45 Smad3-null mice display reduced interstitial fibrosis and cardiac remodeling in response to infarction,⁴⁹ and abnormal TGF-β1–Smad–dependent activation of the myofibroblast lineage can lead to excessive fibrosis that results in chronic fibrotic diseases.^45,50 An SRF/myocardin containing complex has been shown to activate the SM22 promoter following TGF-β1 stimulation of 10T1/2 cells to myofibroblasts.³⁷ Recent studies have also documented the modulation of MRTF-A subcellular localization and activity in response to TGF-β1 signaling in kidney epithelial cells.^36,39 The results of our study extend these findings and reveal a novel function of MRTF-A in contributing to myofibroblast activation and ECM deposition in response to TGF-β1 stimulation of CFs.

TGF-β1–induced myofibroblast activation and fibrosis is blocked by ROCK inhibition in certain contexts, suggesting cooperation between these signaling pathways. Rho-ROCK signaling plays a major role in sensing the environment and generating a cellular response to injury or stress. Mechanical force or receptor-mediated stimulation of the Rho signaling cascade has been shown to activate ROCK and MLC-kinase, promoting a smooth muscle–like myofibroblast phenotype. ROCK and MLC-kinase also stimulate actin cytoskeleton rearrangement, and nuclear translocation of MRTF-A, which contributes to SMC-specific gene expression, thus linking cellular stress to SRF/MRTF-A mediated transcriptional activation.^17,20–23 Rho signaling is also involved in pathological fibrosis of multiple tissues.^27–30,35 Therefore, it seems reasonable that MRTF-A may contribute to ROCK-mediated myofibroblast activation and fibrotic remodeling.

Recently, myofibroblast activation following ischemia/reperfusion and kidney injury has been suggested to originate from circulating inflammatory cells.^35,51,52 Monocyte/macrophage deletion significantly attenuated kidney fibrosis following unilateral ureteric obstruction.⁵¹ Furthermore, bone marrow–derived cells from a donor mouse were detected within the myofibroblast population of the fibrotic heart of host mice, whereas ROCK-1–null mice displayed attenuation of this fibrotic response.⁵² Although the most straightforward interpretation of our results is that MRTF-A mediates ROCK-1 signaling in cardiac myofibroblasts during cardiac fibrosis following MI and Ang II treatment, it is formally possible that another MRTF-A–dependent cell population could contribute to this response. It is also possible that MRTF-A activity in CMCs or SMCs may also contribute to the development of fibrosis following MI or Ang II administration. Although MRTF-A is robustly expressed in CFs and overexpression of MRTF-A stimulates ECM deposition by cultured CFs, tissue-specific ablation or bone marrow transplantation would be required to unequivocally pinpoint whether MRTF-A activity primarily occurs in resident CFs or may also function in additional cell types.

ROCK Inhibitors and Therapy for Pathological Fibrosis

Angiotensin-converting enzyme inhibitors and angiotensin receptor blockers are among the most effective therapies aimed at preventing cardiac remodeling and congestive heart failure after MI.⁵³ The Rho-ROCK signaling pathway has begun to attract attention as a potential therapeutic target in the treatment of various pathological conditions, including vasospasm, arteriosclerosis, ischemia/reperfusion injury, and renal disease, among others.^32,33,35,54 In clinical studies, the ROCK inhibitor, fasudil, has shown efficacy for the treatment of vasospasm and hypertension.^55–57 Our findings demonstrate the involvement of MRTF-A as a potential transcriptional mediator of TGF-β1, Ang II and ROCK signaling during fibrotic remodeling following MI. Importantly, MRTF-A^−/− mice do not display cardiac rupture or increased post-MI lethality, implying that they initially form sufficient scar tissue to undergo wound healing. It is possible that MRTF-A plays a specific role in the promotion of interstitial fibrosis and adverse cardiac remodeling. Characterizing the precise mechanism of MRTF-A activation will enhance our understanding of fibrotic pathologies and hasten the development of MRTF-A inhibitors that may prove useful for the treatment of fibrotic or cardiovascular disease, circumventing the obvious limitations of general Rho inhibitors, which greatly alter cytoskeletal dynamics.