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(The FASEB Journal. 2001;15:785-796.)
© 2001 FASEB

ß1 integrin and organized actin filaments facilitate cardiomyocyte-specific RhoA-dependent activation of the skeletal {alpha}-actin promoter

LEI WEI*, LU WANG*, JAMES A. CARSON{dagger}, JAMES E. AGAN*, KYOKO IMANAKA-YOSHIDA{ddagger} and ROBERT J. SCHWARTZ*1

* Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, Texas, USA;
{dagger} Exercise Science Department, School of Public Health, University of South Carolina, Columbia, South Carolina, USA; and
{ddagger} Department of Pathology, Mie University School of Medicine, Tsu, Mie, Japan

1Correspondence: Department of Cell Biology, Room 145E, Baylor College of Medicine, One Baylor Plaza, Houston, TX 77030, USA. E-mail: schwartz{at}bcm.tmc.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Activation of RhoA GTPase causes actin filament bundling into stress fibers, integrin clustering, and focal adhesion formation through its action on actin cytoskeleton organization. RhoA also regulates transcriptional activity of serum response factor (SRF). Recent studies in NIH 3T3 fibroblasts have shown that SRF activation by RhoA does not require an organized cytoskeleton and may be regulated by G-actin level. In cardiac myocytes, the organization of actin fibers into myofibrils is one of the primary characteristics of cardiac differentiation and hypertrophy. The primary purpose of this study was to examine if RhoA regulates SRF-dependent gene expression in neonatal cardiomyocytes in a manner different from that observed in fibroblasts. Our results show that RhoA-dependent skeletal {alpha}-actin promoter activation requires ß1 integrin and a functional cytoskeleton in cardiomyocytes but not in NIH 3T3 fibroblasts. Activation of the {alpha}-actin promoter by RhoA is greatly potentiated (up to 15-fold) by co-expression of the integrin ß1A or ß1D isoform but is significantly reduced by 70% with a co-expressed dominant negative mutant of ß1 integrin. Furthermore, clustering of ß1 integrin with anti-ß1 integrin antibodies potentiates synergistic RhoA and ß1 integrin activation of the {alpha}-actin promoter. Cytochalasin D and latrunculin B, inhibitors of actin polymerization, significantly reduced RhoA-induced activation of the {alpha}-actin promoter. Jasplakinolide, an actin polymerizing agent, mimics the synergistic effect of RhoA and ß1 integrin on the actin promoter. These observations support the concept that RhoA regulates SRF-dependent cardiac gene expression through cross-talk with ß1 integrin signal pathway via an organized actin cytoskeleton.—Wei, L., Wang, L., Carson, J. A., Agan, J. E., Imanaka-Yoshida, K., and Schwartz, R. J. ß1 integrin and organized actin filaments facilitate cardiomyocyte-specific RhoA-dependent activation of the skeletal {alpha}-actin promoter.


Key Words: serum response factor • cardiac hypertrophy • actin polymerization


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
THE SMALL GTPASE RhoA protein is involved in many actin-based cytoskeletal processes, including formation of stress fibers, cell adhesion, cytokinesis, and contractility (reviewed in refs 1 , 2 ). RhoA also influences gene regulatory activity by stimulating c-fos transcription in fibroblasts (3) and skeletal {alpha}-actin promoter activity in myoblasts (4) via serum response factor (SRF), a homodimeric MADS box-containing transcription factor. SRF is required for mesoderm-dependent muscle formation (5) and expression of skeletal, cardiac, and smooth muscle {alpha}-actin genes, which are the earliest markers for both skeletal and cardiac muscle differentiation (6 7 8) . RhoA’s ability to orchestrate actin filament rearrangement and to regulate SRF transcriptional activity raises the possibility that it might have an important role in facilitating cell morphological change and gene expression during skeletal and cardiac muscle development and hypertrophy. Consistent with this hypothesis, we have shown previously that RhoA signaling through SRF is involved in myoblast terminal differentiation (4) . A recent report also has shown that inhibition of Rho family proteins by the GDP dissociation inhibitor RhoGDI suppresses myoblast terminal differentiation (9) . In addition, RhoA mediates hypertrophic signals induced by {alpha}1-adrenergic agonist phenylephrine (10) , angiotensin II (11) , and mechanical stress (12) in cultured cardiomyocytes.

These observations raise important questions concerning the regulation of SRF-dependent muscle gene expression by RhoA. Specifically, does RhoA-dependent actin cytoskeleton organization facilitate SRF-dependent gene expression? In NIH 3T3 fibroblasts, SRF activation by RhoA does not require an organized cytoskeleton because RhoA activated SRF transcriptional activity in the presence of cytochalasin D, an agent that disrupts actin cytoskeleton (3) . However, cytochalasin D also has been shown to activate RhoA in fibroblasts (13) . In addition, a recent study has implicated the level of G-actin rather than F-actin as a mediator of SRF-dependent gene activity in fibroblasts (14) . Cardiomyocytes are terminally differentiated cells in which actin fiber organization into myofibrils is a primary characteristic of cardiac hypertrophy associated with cardiac hypertrophic gene expression. In addition, skeletal {alpha}-actin may provide a signaling target for RhoA-regulated hypertrophic gene expression in cardiomyocytes. During cardiac hypertrophy, skeletal {alpha}-actin gene expression is induced both in the animal (15) and in culture (16 17 18) . Therefore, it is important to determine if RhoA regulates SRF-dependent gene expression in cardiomyocytes in a manner different from that observed in fibroblasts. Is this mechanism independent or dependent of actin cytoskeleton organization in cardiomyocytes?

Integrins comprise a large family of heterodimeric cell surface receptors that link the extracellular matrix and the intracellular cytoskeleton (reviewed in ref 19 ). Involvement of RhoA in integrin-mediated cell adhesion was characterized in detail in fibroblasts (reviewed in refs 20 21 22 ). RhoA activation causes actin filament bundling into stress fibers and integrin clustering with associated proteins into focal adhesion complexes. Recent studies indicate that this RhoA process is mediated by myosin-based contractility and actin polymerization (reviewed in ref 22 ). RhoA also has direct effects on integrin interactions with extracellular matrix independent of actin cytoskeleton effects (reviewed in ref 20 ). Therefore, it is tempting to speculate that RhoA-induced integrin complex assembly might lead to signaling complex assembly involved in modulating gene expression. ß1 integrin and associated {alpha}-chains are a major subfamily of integrins present in cardiomyocytes. ß1 integrin is up-regulated after induction of hypertrophy in vivo and in vitro (23 , 24) , suggesting an important role in cardiac hypertrophy. ß1D integrin is the major isoform in cardiac and skeletal muscles and replaces the common isoform ß1A during muscle development (25 , 26) . ß1 integrin is also involved in the {alpha}-adrenergic–mediated hypertrophic response in neonatal cardiomyocytes (27) . It is not known if ß1 integrin plays a critical role in coordinating effects of RhoA on the cardiomyocyte actin cytoskeleton with its gene regulation activities.

The main purpose of this study was to ask if RhoA regulates SRF-dependent gene expression in neonatal cardiomyocytes in a manner different from that observed in fibroblasts and if RhoA-dependent skeletal {alpha}-actin promoter activation requires ß1 integrin and a functional cytoskeleton in cardiomyocytes. Our results show that skeletal {alpha}-actin promoter activation by RhoA is greatly potentiated by co-expression of the ß1A or ß1D integrin isoform in cardiac myocytes but is significantly reduced by a dominant negative mutant of ß1 integrin. Furthermore, clustering of ß1 integrin with ß1 integrin-specific antibodies potentiated {alpha}-actin promoter activation by RhoA and ß1 integrin. The inhibition of actin polymerization by cytochalasin D and latrunculin B significantly reduced RhoA-induced skeletal {alpha}-actin promoter activity in cardiac myocytes, whereas jasplakinolide, an actin polymerizing agent, mimicked synergistic effects of RhoA and ß1 integrin on {alpha}-actin promoter activity. Focal adhesion kinase (FAK), an integrator of integrin-dependent signaling, also played a positive role in this pathway. In contradistinction to RhoA-dependent signaling in fibroblasts, our observations support the concept that RhoA regulates SRF-dependent cardiac gene expression through cross-talk with ß1 integrin, and an organized actin cytoskeleton is required for this process.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Plasmid constructs
The reporter plasmid SK-Luc contains the avian skeletal {alpha}-actin promoter from -398 to +25 bp followed by firefly luciferase (16) . The reporter plasmids containing mutations that disrupt respectively the SRF binding site in SRE1 (M-94/-89), the overlapping YY1 site (M-81/-79), transcriptional enhancer factor 1 (TEF-1) binding element (M-70/-65), Sp1 binding site (M-52/-47), and the TATA box (M-28/-23) were described previously (16) .

Expression plasmids pCGN-RhoA, pCGN-V14-RhoA, pCGN-SRF, and pCGN-SRFpm1 were previously described (4) . HA-N19-RhoA inserted into the pCMV5 vector (pCMV5-N19-RhoA) was the gift of Dr. Gary Bokoch (The Scripps Research Institute, San Diego, Calif.) (28) . C3 transferase constructed into the expression plasmid pEF-myc was provided by Dr. Alan Hall (MRC Laboratory for Molecular Cell Biology, London, U.K.) (29) . RhoGDI constructed in the expression plasmid pEF-BOS-Myc was the gift of Dr. Yashimi Takai (Osaka University, Japan) (30) . Expression plasmids pCMVIL2R-ß1 and pCMVIL2R-{alpha}5 encoding chimeric receptors composed of interleukin-2 receptor extracellular and transmembrane domains linked respectively to the cytoplasmic domain of ß1 integrin and {alpha}5 integrin were provided by Drs. Kenneth M. Yamada (NIH, Bethesda, Md.) and Susan E. LaFlamme (Albany Medical College, N.Y.) (31) . Human ß1 integrin isoforms, ß1A and ß1D, constructed into the expression vector pECE were the gift of Dr. Alexey M. Belkin (American Red Cross Holland Laboratory, Md.) (25) . Expressions plasmids pCMV-myctag-FAK and pCMV-myctag-FRNK encoding respectively wild type FAK and the carboxyl-terminal domain of FAK were provided by Dr. J Thomas Parsons (University of Virginia, Charlottesville) (32) . Identity and expression of each expression plasmid used in the study has been confirmed by DNA sequencing and by expression analysis by Western blot or by cellular immunostaining as described below.

Tissue culture, transient transfection, and reporter gene assays
Neonatal rat ventricular myocytes were prepared from hearts of 2- to 3-day-old Sprague-Dawley rat pups as described previously (33) . Briefly, hearts were digested with collagenase, trypsin, and deoxyribonuclease I. Myocytes were then purified over a Percoll gradient. Myocytes were plated at a density of 1 x 106 cells per 35-mm dish (Primaria, Falcon, Becton Dickinson, Franklin Lakes, N.J.) and cultured overnight in Dulbecco’s Modified Eagle’s Medium (DMEM) (Life Technologies/BRL, Gaithersburg, Md.) containing 5% fetal bovine serum and 10% horse serum. Cells were then transfected with up to 1.5 µg of total plasmid DNA containing the indicated reporter plasmid and the indicated expression plasmids balanced with parental expression vector. Transfections were performed using lipofectamine (Life Technologies/BRL) according to the manufacturer’s instructions. After transfection, myocytes were cultured overnight in DMEM with 5% horse serum, then in serum-free medium. After 24 h, cells were harvested and assayed for luciferase activity and protein content as described previously (16) . Luciferase activity was normalized to the total protein, and data were expressed as luciferase activity normalized to baseline reporter gene activity. All experiments were performed in duplicate or triplicate and were repeated 3–5 times. For experiments evaluating cytochalasin D (Biomol, Plymouth Meeting, Penn.), jasplakinolide (kindly provided by the Drug Synthesis & Chemistry Branch, Developmental Therapeutics Program, Division of Cancer Treatment, NCI, NIH, Bethesda, Md.), or latrunculin B (Biomol), initial culture conditions were as described above. Each drug was then added into the serum-free medium ~40 h after transfection, and cultured cells were then harvested after 4 h of treatment.

NIH 3T3 cells were seeded at 5 x 105 cells per 60-mm dish in DMEM supplemented with 10% newborn calf serum (NCS) 24 h before transfection. Cells were transfected as described previously and were maintained in DMEM with 0.5% NCS after transfection for 40–48 h. Cells were then harvested and assayed for luciferase activity and protein content as above. For serum induction, the low serum medium was replaced by DMEM with 10% NCS 40 h after transfection, and cells were harvested after 4 h of incubation.

ß1 integrin clustering
Cardiomyocytes were transfected with SK-Luc reporter plasmid together with expression plasmids as described above. After 40 h, serum-free medium containing monoclonal anti-human ß1 integrin antibody K20 (Dako, Santa Barbara, Calif.) (2.5 µg/ml) was added to cells for 30 min, followed by addition of goat anti-mouse IgG (Sigma, St. Louis, Mo.) (2.5 µg/ml) for 4 h. Control cells received similar treatment except that mouse IgG (Sigma) (2.5 µg/ml) was added instead of K20 antibody. Cells were then harvested and assayed for luciferase activity as described above.

Immunofluorescence analysis
Visualization of transfected ß1A or ß1D integrin expression was implemented in cardiomyocytes attached to coverslips coated with rat tail collagen for 24 h, followed by transfection with expression plasmids encoding human ß1A or ß1D integrin. Cells were incubated with 2.5 µg/ml anti-human ß1 integrin antibody K20 for 30 min at 4°C, washed, incubated with fluorescein-conjugated goat anti-mouse IgG (Molecular Probes, Junction City, Oreg.) for 30 min, and fixed. Cells were then stained with rhodamine conjugate of phalloidin (against F-actin, Molecular Probes). For visualization of chimeric receptor IL2R-ß1 or IL2R-{alpha}5 expression, transfected cardiomyocytes were incubated with mouse monoclonal antibody 7G7B6 (ATCC, Manassas, Va.) against interleukin-2 receptor extracellular domain followed by incubation with fluorescein-conjugated goat anti-mouse IgG as described above.

Statistical analysis
The data are expressed as means ± SE, relative to parallel cultures of control vector-transfected cells. Student’s t test was used for data comparison, using a significance level of P<0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
RhoA regulates skeletal {alpha}-actin promoter activity via SRF in neonatal cardiomyocytes
Cultured neonatal rat ventricular myocytes have served as a model system for studies aimed to understand the mechanisms of cardiac hypertrophy. We asked if RhoA activates skeletal {alpha}-actin promoter in cardiomyocytes. MacLellan et al. (16) have shown that activity of this promoter was 100-fold greater in cardiomyocytes than in cardiac fibroblasts and 30-fold greater than in HeLa cells. RhoA and constitutively active V14-RhoA activated the skeletal {alpha}-actin promoter activity in a dose-dependent manner, and V14-RhoA is a more potent activator than RhoA (Fig. 1 At an optimal transfection dose of 0.5 µg, RhoA and V14-RhoA activated the promoter up to 2.5- or 4-fold, respectively. In addition, dominant negative N19-RhoA and C3 transferase, which inactivates RhoA by ADP-ribosylating RhoA within its effector domain, significantly inhibited the promoter activity (P=0.008 and 0.007, respectively) (Fig. 1A ). In comparison, hypertrophic stimuli such as the {alpha}1-adrenergic agonist, transforming growth factor ß (TGFß), and mechanical stretch elevated the skeletal {alpha}-actin promoter activity by ~2-, 5-, and 2.5-fold, respectively, in cardiomyocytes (12 , 16 , 34) .



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  		Figure 1. RhoA regulates the skeletal {alpha}-actin promoter via SRF in neonatal cardiomyocytes. A) Effect of the wild-type RhoA, V14-RhoA, SRFpm1 (pm1), N19 RhoA (N19), and C3 transferase (C3) on the activity of the skeletal {alpha}-actin promoter. Cultured neonatal cardiomyocytes were transfected with 0.5 µg SK-Luc reporter plasmid together with either the control vector plasmid (1 µg) or expression plasmid encoding RhoA (0.1, 0.25, or 0.5 µg), V14-RhoA (0.1, 0.25, or 0.5 µg), SRFpm1 (0.25 µg), SRFpm1 (0.25 µg) with V14-RhoA (0.5 µg), N19-RhoA (0.5 µg), or C3 transferase (0.25 µg). After 48 h, the cells were harvested and assayed for luciferase activity. Luciferase activity was normalized to the protein content in cell lysates. The values (means ± SE) are expressed relative to control vector transfected cells. The effect of each transfection dose on the activity of the reporter gene was measured in duplicate or triplicate, and the results indicated are representative of three to five independent experiments. B) Activation of the skeletal {alpha}-actin promoter by RhoA requires the proximal SRF binding site and TATA box. Cardiomyocytes were co-transfected with 1 µg of mutated skeletal {alpha}-actin promoter reporter plasmids as indicated and 0.5 µg of control vector or V14-RhoA. For each reporter plasmid, the luciferase activity (mean ± SE) is shown for V14-RhoA–transfected cells relative to control vector-transfected cells. C) Activation of the promoter containing an SRE by V14-RhoA. Cardiomyocytes were transfected with 1 µg of SRE1-Luc, c-fos-SRE-Luc, TEF1-Luc, A20-Luc, or TATA-Luc as indicated and 0.5 µg of control vector or V14-RhoA. *P<0.05 vs. control vector-transfected cells. #P<0.05 vs. skeletal {alpha}-actin promoter activity induced by V14-RhoA.

In cardiomyocytes, the promoter activity in V14-RhoA–transfected cells was inhibited by co-transfected SRFpm1, a dominant negative mutant of SRF (Fig. 1A ). Investigation of the promoter region of the avian {alpha}-actin gene identified several cis-acting elements, located within the first 200 base pairs, which are important in developmental and tissue-specific regulation of this gene and to its response to hypertrophic stimuli. These cis-acting regulatory elements include the SRF binding site in SRE1, a YY1 binding site overlapping SRE1, TEF-1 binding site, two Sp1 binding elements, and a TATA box (Fig. 1B ). Mutations of these regulatory elements efficiently block the binding of cognate transcription factors (16 , 35) . We observed that mutations of SRE1 and TATA box significantly reduced promoter activity induced by V14-RhoA (P=0.003 and 0.018, respectively) (Fig. 1B ). Mutation of the negative-acting YY1 site increased induction of the promoter by V14-RhoA (P=0.042), consistent with the previous findings that YY1 acts as a competitive inhibitor of SRF for regulation of the skeletal {alpha}-actin gene expression in cardiomyocytes (16) . Mutated TEF-1 and Sp1 binding sites had no significant effect (P=0.260 and 0.328, respectively) on the activation of the promoter by RhoA (Fig. 1B ). Effects of these mutations on the basal activity of the promoter (data not shown) were the same as described previously (16) . These results indicate that RhoA stimulates the skeletal {alpha}-actin promoter via an intact SRE1 in cardiomyocytes, consistent with a regulatory role for SRF.

Because the mutation on the SRE1 site disrupted RhoA-dependent activation of the skeletal {alpha}-actin promoter, we then asked if an isolated SRE mediates RhoA-dependent gene activation. The skeletal {alpha}-actin SRE1 (SRE1-Luc) and TEF-1 (TEF1-Luc) sites, c-fos SRE (c-fos-SRE-Luc) site, and high-affinity Nkx2.5 binding site (A20-Luc) were placed upstream of a neutral promoter that contains only the TATA box (TATA-Luc) as described previously (16 , 36) . Only promoters containing an SRE site, such as SRE1 or a c-fos-SRE, were stimulated by V14-RhoA in transfected cardiomyocytes (P=0.033 and 0.008, respectively) (Fig. 1C ), further supporting a positive regulatory role for SRF.

Disruption of integrin ß1 signaling inhibits skeletal {alpha}-actin promoter activity stimulated by RhoA in cardiomyocytes but not in NIH 3T3 fibroblasts
We asked if ß1 integrin mediates RhoA signaling on the skeletal {alpha}-actin promoter activity. We used a chimeric receptor (Fig. 2 composed of the extracellular and transmembrane domains of interleukin-2 receptor and the cytoplasmic domain of ß1A integrin (IL2R-ß1), which has been shown to disrupt integrin signaling in noncardiac cells (37 , 38) . A similar construct containing the cytoplasmic domain of integrin {alpha}5 (IL2R-{alpha}5) and the expression vector served as controls. A recent study has shown that expression of IL2R-ß1 inhibited adrenergic induction of atrial natriuretic factor (ANF) production in cultured cardiomyocytes (27) . Our results show that IL2R-ß1 significantly reduced the activity of the promoter in control vector-transfected cells (P=0.039) and in V14-RhoA-transfected cells (P=0.022) at a transfection dose of 0.25 or 0.5 µg, whereas IL2R-{alpha}5 had no significant effect (P=0.12 and P=0.299, respectively) (Fig. 2B ). Expression of these chimeric proteins at the cell surface was confirmed by immunostaining using an antibody against interleukin-2 receptor (Fig. 2D ). Thus, disruption of ß1 integrin-mediated signaling reduced RhoA-dependent activation of the skeletal {alpha}-actin promoter.



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  		Figure 2. RhoA-dependent activation of the skeletal {alpha}-actin promoter is reduced by alteration of ß1 integrin signaling in cardiomyocytes but not in NIH 3T3 fibroblasts. A) Schematic diagram of IL2Rß1 and IL2R{alpha}5 chimeras. B) Expression of IL2Rß1 inhibits RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes. Cardiomyocytes were transfected with 0.5 µg of SK-Luc together with 1 µg of control vector plasmids or with V14-RhoA (0.5 µg), IL2Rß1 (0.1, 0.25, or 0.5) µg, and IL2R{alpha}5 (0.5 µg) alone or in combination as indicated. C) Expression of IL2Rß1 has no significant effect on serum-induced activation of the skeletal {alpha}-actin promoter in NIH 3T3 fibroblasts. NIH 3T3 cells were transfected with 0.5 µg of SK-Luc together with 0.5 µg of control vector plasmids or with V14-RhoA (0.5 µg), RhoGDI (0.5 µg), IL2Rß1 (0.25 or 0.5 µg), or IL2R{alpha}5 (0.25 or 0.5 µg) as indicated. For serum induction, transfected cells was incubated for 4 h with 10% NCS 40 h after transfection and then harvested. D) Expression of IL2Rß1 in cardiomyocytes. Transfected cells were incubated with monoclonal antibody 7G7B6 against interleukin-2 receptor extracellular domain followed by incubation with fluorescein-conjugated goat anti-mouse IgG. Cells were then fixed and stained with rhodamine-conjugated phalloidin. The same field was visualized for fluorescently labeled IL2Rß1 (left panel) and for phalloidin labeled actin (right panel). Expression of IL2R{alpha}5 was confirmed by the same method. *P < 0.05 vs. control vector-transfected cells. #P < 0.05 vs. promoter activity induced by V14-RhoA alone. **P < 0.05 vs. promoter activity induced by serum.

We also determined if ß1 integrin is involved in RhoA-dependent activation of the skeletal {alpha}-actin promoter in NIH 3T3 fibroblasts. We observed that skeletal {alpha}-actin promoter was stimulated by transfected V14-RhoA (~2.9-fold) and by serum (~3.3-fold) in NIH 3T3 cells (Fig. 2C ). Serum-induced activation was repressed up to 90% by a RhoA inhibitor, RhoGDI (Fig. 2C ), indicating that endogenous RhoA mediates serum-induced activation of SRF as described previously (3) . Actin promoter activity induced by serum was slightly increased by addition of IL2R-ß1, but there was no significant difference between addition of IL2R-ß1 and IL2R-{alpha}5 (P>0.257) (Fig. 2C ). Taken together, these results suggest that mechanisms by which integrins influence RhoA-dependent SRF activation may be cell-type dependent.

Integrin ß1A and ß1D isoforms facilitate robust skeletal {alpha}-actin promoter activation by RhoA in cardiomyocytes but not in NIH 3T3 fibroblasts
We next asked if overexpression of ß1 integrin potentiates RhoA-dependent activation of the skeletal {alpha}-actin promoter. Human ß1D and ß1A expression plasmids were co-transfected with the reporter construct. Human ß1 integrin is similar to rat ß1 integrin but can be distinguished from the endogenous rat ß1 integrin with species-specific antibodies. ß1D and ß1A are very homologous, differing by only 13 amino acids in their cytoplasmic tail. Co-transfecton of ß1A and ß1D expression plasmids with the skeletal {alpha}-actin promoter reporter plasmid in cardiomyocytes increased lucifearse activity up to 1.7- and 2.5-fold, respectively (Fig. 3A ). Co-expression of ß1A or ß1D with V14-RhoA resulted in a strong synergistic augmentation of the skeletal {alpha}-actin promoter activity in a dose-dependent manner up to 10- to 15-fold (Fig. 3A ). Cellular expression of exogenous ß1A and ß1D was confirmed by immunostaining using monoclonal antibody K20, raised against human ß1 integrin (Fig. 3C ). These results indicate that overexpression of ß1 integrin potentiates effects of RhoA on the activation of the skeletal {alpha}-actin promoter. This magnitude of induction of the promoter is higher than all previously reported inductions of the same promoter by hypertrophic stimuli in cardiomyocytes (12 , 16 , 33 , 34) . In contrast to cardiomyocytes, transfection of ß1A and ß1D integrins had no detectable effects (P>0.218) on serum-induced activation of the skeletal {alpha}-actin promoter in NIH 3T3 fibroblasts, which is mediated by RhoA (Fig. 3B ).



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  		Figure 3. ß1A and ß1D integrins potentiate RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes but not in NIH 3T3 fibroblasts. A) Co-activation of the skeletal {alpha}-actin promoter by V14-RhoA with ß1D or ß1A. Cardiomyocytes were transfected with 0.5 µg of SK-Luc together with 1 µg of control vector plasmids or with V14-RhoA (0.5 µg), ß1A, and ß1D (0.25 or 0.5 µg) alone or in combination as indicated. B) Expression of ß1D or ß1A has no significant effect on serum-induced activation of the skeletal {alpha}-actin promoter in NIH 3T3 fibroblasts. NIH 3T3 cells were transfected with 0.5 µg of SK-Luc together with 0.5 µg of control vector, ß1A, or ß1D (0.25 or 0.5 µg) as indicated. For serum induction, transfected cells were incubated for 4 h with 10% NCS 40 h after transfection and then harvested. C) Expression of ß1D integrin in cardiomyocytes. Transfected cells were incubated with monoclonal antibody K20 (2.5 µg/ml) raised against human ß1 integrin followed by incubation with fluorescein-conjugated goat anti-mouse IgG. Cells were then fixed and stained with rhodamine conjugated phalloidin. The same field was visualized for fluorescently labeled ß1D integrin (left panel) and for phalloidin labeled actin (right panel). Expression of ß1A in transfected cells was confirmed by the same method. *P < 0.05 vs. control vector-transfected cells. #P < 0.05 vs. promoter activity induced by V14-RhoA alone.

ß1 integrin clustering enhances co-activation of the skeletal {alpha}-actin promoter by RhoA and ß1 integrin in cardiomyocytes
We then questioned if ß1 integrin clustering, which is a critical event in integrin activation (23 , 24) , potentiates RhoA-dependent skeletal {alpha}-actin promoter activity. We used anti-human ß1 integrin antibody K20 to aggregate exogenous ß1 integrin on the cell surface. The K20 antibody does not affect ligand binding and serves as a nonactivating anti-ß1 antibody. We found that addition of K20 alone actually inhibited the skeletal {alpha}-actin promoter activation by V14-RhoA and ß1 integrin (Fig. 4 ). K20 was previously observed to impart negative signaling in lymphocytes (39) . Increased ß1 integrin clustering by addition of K20 and secondary antibodies nearly doubled the skeletal {alpha}-actin promoter activity induced by V14-RhoA and ß1D compared with the addition of K20 alone (P=0.032) (Fig. 4) . On the other hand, no significant difference was observed when IgG was added alone or in combination with the secondary antibody (P=0.466) (Fig. 4) . These results indicate that ß1 integrin clustering potentiates RhoA-dependent activation of the skeletal {alpha}-actin promoter.



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  		Figure 4. Clustering of ß1 integrin stimulates co-activation of skeletal {alpha}-actin promoter by RhoA and ß1 integrin in cardiomyocytes. Approximately 40 h after transfection, serum-free medium containing monoclonal anti-ß1 integrin antibody K20 (2.5 µg/ml) was added to cells for 30 min, one set of cells was followed by addition of secondary antibody goat anti-mouse IgG (2.5 µg/ml) for 4 h. One set of cells was treated with mouse IgG (2.5 µg/ml) instead of K20 antibody. Cells were then harvested and assayed for luciferase activity as described above. The values (means ± SE) are expressed relative to control vector-transfected cells. *P<0.05 vs. promoter activity induced by V14-RhoA and ß1D integrin. #P<0.05 vs. promoter activity induced by V14-RhoA and ß1D integrin and in the presence of K20 alone.

Focal adhesion kinase is involved in RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes
Our observation that ß1 integrin-mediated signals regulate RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes prompted the examination of whether FAK, an important downstream target of integrins, works as a mediator of RhoA signaling in skeletal {alpha}-actin promoter activation. A truncated mutant of FAK, FAK-related nonkinase (FRNK), which contains the carboxyl-terminal domain of FAK without kinase domain (Fig. 5A ), has previously been shown to act as a dominant negative mutant inhibiting cell spreading and focal adhesion formation. We evaluated the role of FAK and FRNK on the activation of the skeletal {alpha}-actin promoter by RhoA and ß1 integrin using transfection doses of 0.5 µg DNA for FAK and FRNK and 0.25 µg DNA for V14-RhoA and ß1D. It is important to note that co-expression of V14-RhoA and ß1D under this condition resulted in ~6-fold activation of the actin promoter (Fig. 3A ). FRNK reduced the promoter activity in control vector-transfected cells (P=0.011) and in V14-RhoA- (P=0.024), ß1D- (P=0.050), or V14-RhoA and ß1D- (P=0.029) transfected cells (Fig. 5B ), indicating that FAK may be required for efficient activation of the {alpha}-actin promoter. Expression of FAK had no significant effect on the basal activity of the promoter (P=0.238) and slightly increased activation by V14-RhoA (P=0.043), by ß1D (P=0.048) and by co-expression of V14-RhoA and ß1D (P=0.045) (Fig. 5B ). These results suggest that endogenous FAK might not be a limiting factor for the activation of the actin promoter under these conditions. However, the inhibitory effect of FRNK on stimulated {alpha}-actin promoter activity by V14-RhoA and ß1D suggests that FAK cross-talks and contributes to RhoA and ß1 integrin signaling.



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  		Figure 5. Cross-talk between RhoA, ß1 integrin, and FAK co-activates the skeletal {alpha}-actin promoter in cardiomyocytes. A) Schematic diagram of Myc-tagged wild-type FAK and dominant negative FRNK. B) Cardiomyocytes were transfected with 0.5 µg of SK-luc together with 1 µg of control vectors or with V14-RhoA (0.25 µg), ß1D integrin (0.25 µg), FAK (0.5 µg), and FRNK (0.5 µg) alone or in combination as indicated. The values (means ± SE) are expressed relative to control vector-transfected cells. *P < 0.05 vs. control vector-transfected cells. #P < 0.05 vs. promoter activity induced by ß1D integrin. **P < 0.05 vs. promoter activity induced by V14-RhoA. ##P < 0.05 vs. promoter activity induced by V14-RhoA and ß1D.

Treatment with cytochalasin D reduces RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes but not in NIH 3T3 fibroblasts
We asked if treatment with cytochalasin D, a specific inhibitor of actin polymerization (40) , affected activation of the skeletal {alpha}-actin promoter by RhoA. We observed that the treatment of cardiomyocytes with cytochalasin D (0.2 µM) significantly reduced the skeletal {alpha}-actin promoter activation by V14-RhoA (P=0.024), ß1D integrin (P=0.048), or by V14-RhoA and ß1D integrin (P=0.012) (Fig. 6A ). In contrast, cytochalasin D slightly increased activation of the skeletal {alpha}-actin promoter by V12-Ras alone or in combination with ß1D integrin (Fig. 6A ). Previous studies performed with NIH 3T3 fibroblasts have shown that treatment with cytochalasin D allowed activation of SRF-targeted reporter genes and did not interfere with RhoA-mediated induction of c-fos transcription by LPA and serum (3) . Consistent with previous reports, we observed that in NIH 3T3 cells, treatment with cytochalasin D significantly increased basal activity of the actin promoter (P=0.022), and activation of the promoter by transfected V14-RhoA (P=0.003) or by serum treatment (P=0.039) (Fig. 6B ).



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  		Figure 6. Organized actin cytoskeleton is required for RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes. A) Treatment of cardiomyocytes with Cytochalasin D reduces activation of the skeletal {alpha}-actin promoter by RhoA and ß1 integrin but not that induced by Ras and ß1 integrin. Cardiomyocytes were transfected with 1 µg of control vector plasmids or with 0.5 µg of V14-RhoA, V12-Ras, ß1D alone, or in combination as indicated. Approximately 40 h after transfection, cells were treated with or without cytochalasin D (0.2 µM) for 4 h and then harvested. The values (means ± SE) are expressed relative to control vector-transfected cells without drug treatment. B) Treatment of NIH 3T3 fibroblasts with cytochalasin D increases RhoA-dependent activation of the skeletal {alpha}-actin promoter. NIH 3T3 cells were transfected with 0.5 µg of SK-Luc together with 0.5 µg of control vector or with 0.5 µg of V14-RhoA. One set of control vector-transfected cells was incubated for 4 h with 10% NCS 40 h after transfection. Transfected cells were treated with or without cytochalasin D (0.2 µM). C) Activation of the skeletal {alpha}-actin promoter by RhoA is inhibited by latrunculin B and increased by jasplakinolide in cardiomyocytes. Cardiomyocytes were transfected with 1 µg of control vectors or with 0.5 µg of V14-RhoA and ß1D, alone or in combination, as indicated. Approximately 40 h after transfection, cells were treated with or without latrunculin B (0.2 or 1 µM) or jasplakinolide (0.025 or 0.1 µM) for 4 h and then harvested. D) Phalloidin staining of cardiomyocytes treated with cytochalasin D (0.2 µM), latrunculin B (0.2 µM), or Jasplakinolide (0.025 µM) compared with untreated cells. Large arrows indicate nonstriated actin fibers and small arrows indicate striated actin fibers in cardiomyocytes. *P < 0.05 vs. untreated control vector-transfected cells. #P < 0.05 vs. untreated ß1D integrin-transfected cells. **P < 0.05 vs. untreated V14-RhoA-transfected cells. ##P < 0.05 vs. untreated V14-RhoA and ß1D-transfected cells. ***P < 0.05 vs. untreated control vector-transfected cells in the presence of serum.

In cardiomyocytes, we also evaluated another actin polymerization inhibitor, latrunculin B (41) , on the skeletal {alpha}-actin promoter activation induced by V14-RhoA alone or together with ß 1D integrin. Latrunculin B (0.2 µM) significantly reduced basal activity of the skeletal {alpha}-actin promoter (P=0.034), activation by ß 1D integrin (P=0.016), activation by V14-RhoA (P=0.040), and co-activation by V14-RhoA and ß 1D integrin (P=0.014) (Fig. 6C ). The inhibitory effect of latrunculin B on the actin promoter activity was greater in comparison with that of cytochalasin D.

Stabilization of the actin cytoskeleton potentiates the skeletal {alpha}-actin promoter activity induced by RhoA
Jasplakinolide stablizes F-actin and promotes actin polymerization (42) . We observed that jasplakinolide increased activity of the {alpha}-actin promoter in control vector, ß 1D integrin or V14-RhoA-transfected cells up to the level induced by co-expression of V14-RhoA and ß 1D integrin, but had no significant effect (P=0.472 and 0.265 for 25 and 100 nM jasplakinolide, respectively) on promoter activity induced by V14-RhoA and ß 1D integrin (Fig. 6C ). These results indicate that actin polymerization potentiates RhoA-dependent activation of the skeletal {alpha}-actin promoter and jasplakinolide mimics effect of co-expression of V14-RhoA and ß 1D integrin, consistent with the above observation that the actin cytoskeleton is involved in the activation of the skeletal {alpha}-actin promoter by RhoA.

During myofibrillogenesis, cardiomyocytes assemble nonstriated actin fibers, which look similar to stress fibers in fibroblasts by phalloidin staining (43 , 44) . These nonstriated actin fibers develop into striated mature sarcomeric actin fibers (44 , 45) . Phalloidin staining showed that nonstriated actin fibers, which are enriched at the periphery of the untreated myocytes (Fig. 6D , large arrows), were destroyed in cytochalsin D- or latrunculin B-treated myocytes, whereas sarcomeric actin fibers, indicated by punctated actin staining, were still observed (Fig. 6D , small arrows). On the other hand, jasplakinolide caused prominent nonstriated actin fiber formation, especially at the periphery of the myocytes where the sarcomeric actin fibers were masked because of the high levels of nonstriated actin fibers (Fig. 6D , large arrows). On the other hand, both cytochalasin D and latrunculin B abolished stress fiber formation, whereas jasplakinolide enhanced stress fiber organization in cardiac fibroblasts (data not shown). These results suggest that these drugs have similar effects both on nonstriated actin fibers in myocytes and on stress fibers in fibroblasts.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Activation of the skeletal {alpha}-actin promoter by RhoA requires ß1 integrin in cardiomyocytes
Our results demonstrate that in neonatal cardiomyocytes, RhoA regulates the promoter activity of skeletal {alpha}-actin, a representative fetal cardiac gene that is up-regulated in the hypertrophic myocardium (Fig. 1) . A role for SRF in mediating RhoA signaling on skeletal {alpha}-actin promoter is supported by our observation that a dominant negative SRF mutant, SRFpm1, or an SRE1 site-directed promoter mutant inhibited RhoA-mediated promoter activation.

We provided several lines of evidence supporting a role for ß1 integrin in regulating RhoA-dependent activation of the skeletal {alpha}-actin promoter. First, disruption of ß1 integrin-mediated signaling significantly reduced RhoA-dependent activation of the skeletal {alpha}-actin promoter (Fig. 2) . Second, both ß1A and ß1D isoforms greatly increased skeletal {alpha}-actin promoter activity stimulated by RhoA, and the magnitude of induction of the promoter is greater than any of the previous reports evaluating skeletal {alpha}-actin promoter activity induced by hypertrophic stimuli in cardiomyocytes (12 , 16 , 34) (Fig. 3) . Third, clustering of ß1 integrin by antibodies increased co-activation of the skeletal {alpha}-actin promoter by V14-RhoA and ß1 integrin (Fig. 4) . Fourth, an important downstream molecule of integrins, FAK, modulated RhoA signaling in skeletal {alpha}-actin promoter activation (Fig. 5) .

Previous studies strongly support a role for ß1 integrin in cardiac hypertrophy. Stretched cultured cardiomyocytes have a 71% increase in cellular ß1 integrin protein (24) , which is similar to the up-regulation of the protein level of ß1 integrin protein and its associated {alpha}1 and {alpha}5 subunits by pressure overload-induced cardiac hypertrophy (23) . Increasing ß1 integrin augments the {alpha}1-adrenergic–mediated hypertrophic responses that include increased protein synthesis and ANF production in cultured cardiomyocytes (27) . ß1 integrin also regulates myofibrillogenesis in cultured cardiomyocytes (46) . RhoA may mediate stretch and {alpha}1-adrenergic–induced hypertrophic gene expression in cultured cardiomyocytes (10 , 12) . In addition, our results also support a role for ß1 integrin in RhoA-dependent signaling pathways to direct stretch and {alpha}-adrenergic induction of hypertrophic gene expression.

It has recently been shown that the major ß1 integrin isoform in cardiac and skeletal muscles is ß1D, which replaces the common isoform ß1A during development (25 , 26) . Our results show that both isoforms are involved in the regulation of the skeletal {alpha}-actin promoter activity in cardiomyocytes. In addition, both isoforms also potentiate c-fos SRE activity induced by RhoA in cardiomyocytes (data not shown). Together, these results suggest that both isoforms share common signaling pathways in cardiac hypertrophy. A previous study has shown that both integrin isoforms activate FAK in nonmuscle cells (25) . Moreover, a recent study has shown that FAK is also involved in {alpha}1-adrenergic–induced hypertrophic responses in cultured cardiomyocytes (47) . Our results suggest that FAK is also activated by ß1 integrin in cardiomyocytes and involved in mediating ß1 integrin signaling activation of the skeletal {alpha}-actin promoter. FAK has been directly linked through its SH2 and SH3 binding sites to an extensive network of signaling molecules, such as tyrosine kinase Src, adaptor proteins Grb2 and Cas, cytoskeleton proteins paxillin and talin, Rho GTPase activating protein Graf, and lipid kinase PI 3-kinase (reviewed in ref 48 ). The nature of protein–protein interactions mediated by FAK and involved in up-regulation of hypertrophic marker genes induced by RhoA and ß1 integrin appears to be complex and remains to be determined.

An organized actin cytoskeleton facilitates the activation of the skeletal {alpha}-actin promoter by RhoA in cardiomyocytes
Also, several lines of evidence demonstrate that an organized actin cytoskeleton is involved in the activation of the skeletal {alpha}-actin promoter by RhoA and ß1 integrin. First, treatment of cardiomyocytes with cytochalasin D and latrunculin B, two agents that disrupt the actin cytoskeleton, significantly reduced skeletal {alpha}-actin promoter stimulated by RhoA and ß1D integrin (Fig. 6) . In addition, these two drugs disrupted nonstriated actin filaments but not actin-containing sarcomeres in cardiomyocytes (Fig. 6) , suggesting that nonstriated actin fiber formation is required for the activation of the actin promoter by RhoA and ß1 integrin. Second, jasplakinolide, an actin polymerizing agent, increased skeletal {alpha}-actin promoter activity, mimicked the synergy of RhoA and ß1D integrin on promoter activation, and caused prominent nonstriated actin fiber formation in cardiomyocytes (Fig. 6) . This further supports a role for nonstriated actin fiber formation in the activation of the actin promoter by RhoA and ß1 integrin. It is well established that cardiomyocytes assemble nonstriated actin fibers, which develop into striated mature sarcomeric actin fibers, and such organization of actin fibers into myofibrils (morphological hypertrophy) is one of the primary characteristics of cardiac hypertrophy associated with nuclear-directed hypertrophic gene expression (49) . RhoA has been shown to mediate both morphological hypertrophy and hypertrophic gene expression in cardiomyocytes (11 , 50) . Our results suggest that RhoA effects on organization of nonstriated actin fiber may facilitate hypertrophic gene expression in cardiomyocytes.

RhoA is required for the clustering of the integrin in fibroblasts through increasing myosin-based contractility and actin polymerization (reviewed in refs 20 21 22 ). In neonatal cardiomyocytes, it is not certain how RhoA and ß1 integrin signaling pathways interact to regulate skeletal {alpha}-actin promoter activity. Because an organized actin cytoskeleton is required for the activation of the skeletal {alpha}-actin promoter by RhoA and ß1 integrin, RhoA-dependent aggregation of ß1 integrin might be involved in regulating hypertrophic gene expression in cardiomyocytes.

A recent study in fibroblasts reported that the amount of G-actin, rather than F-actin, controls SRF activity and that cytochalasin D activates SRF activity, possibly by preventing interaction of G-actin with the partner molecules involved in SRF regulation (14) . Consistent with this report, we have observed in NIH 3T3 fibroblasts that skeletal {alpha}-actin promoter activation by V14-RhoA or serum, which is mediated by endogenous RhoA, was also increased by cytochalasin D treatment (Fig. 6) . However, we observed significant differences on the effects of cytochalasin D on RhoA-dependent skeletal {alpha}-actin promoter activation in cardiomyocytes versus fibroblasts. Our studies demonstrate that intact actin cytoskeleton organization is required for RhoA-dependent signaling in cardiomyocytes. Although we can not exclude a direct role for G-actin in regulating SRF activity in cardiomyocytes, it does not appear to have a primary role.

RhoA regulates SRF-dependent gene expression in cardiomyocytes through a signal pathway distinct from that in fibroblasts
In addition, we have also found that ß1 integrin is not involved in the induction of the skeletal {alpha}-actin promoter by serum in fibroblasts (Fig. 6) . These observations support the concept that in cardiomyocytes RhoA regulates SRF-dependent gene expression through a signal pathway distinct from that in fibroblasts. RhoA-dependent activation of the skeletal {alpha}-actin promoter in cardiomyocytes may involve recruitment of accessory factors that are absent in fibroblasts. For example, SRF is able to recruit cardiac-enriched transcription factors, including Nkx2.5 (36) and GATA4 (51) , to activate cardiac gene expression. SRF also associates with myogenic factors, such as myogenin and MyoD (52) . Further investigations are under way to evaluate the role of cardiac-specific co-factors in mediating RhoA signaling on SRF-dependent gene expression.

In summary, our studies indicate that ß1 integrin and an organized actin cytoskeleton facilitate skeletal {alpha}-actin promoter activity stimulated by RhoA in neonatal rat cardiomyocytes. Also, RhoA regulates SRF-dependent gene expression in cardiomyocytes through a signal pathway that is different from that observed in fibroblasts. Our observations support the concept that in cardiomyocytes RhoA may mediate hypertrophic gene expression by way of ß1 integrin signaling, an organized actin cytoskeleton, and SRF-dependent activity. Because only a small percentage of cardiomyocytes are transfected in the present study, further studies are necessary to evaluate the role of this important signaling pathway to regulate endogenous hypertrophic gene expression both in cultured neonatal cardiomyocytes as well as in vivo.


   ACKNOWLEDGMENTS
 
We thank Drs. Gary Bokoch (The Scripps Research Institute), Alan Hall (MRC Laboratory for Molecular Cell Biology), Yashimi Takai (Osaka University), Kenneth M. Yamada (NIH), Susan E. LaFlamme (Albany Medical College), Alexey M. Belkin (American Red Cross Holland Laboratory), and J Thomas Parsons (University of Virginia) for providing expression plasmids. We also thank Dr. Michael D. Schneider for helpful discussions. This work was supported by NIH R01 HL50422 and P01 HL49953 (to R. J. S.) and by a Beginning Grant-in-Aid Award from American Heart Association, Texas Affiliate (to L. W.).

Received for publication May 18, 2000. Revision received August 14, 2000.
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 DISCUSSION
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