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. 2011 Feb 21;589(Pt 8):2027–2039. doi: 10.1113/jphysiol.2011.205468

Striated muscle activator of Rho signalling (STARS) is a PGC-1α/oestrogen-related receptor-α target gene and is upregulated in human skeletal muscle after endurance exercise

Marita A Wallace 1, M Benjamin Hock 2, Bethany C Hazen 2, Anastasia Kralli 2, Rod J Snow 1, Aaron P Russell 1
PMCID: PMC3090601  PMID: 21486805

Non-technical summary

Exercise improves the ability of skeletal muscle to metabolise fats and sugars. For these improvements to occur the muscle detects a signal caused by exercise, resulting in changes in genes and proteins that control metabolism. We show that endurance exercise increases the amount of a protein called striated muscle activator of Rho signalling (STARS) as well as several other proteins influenced by STARS. We also show that the amount of STARS can be increased by signals directed from proteins called peroxisome proliferator-activated receptor gamma co-activator 1-α (PGC-1α) and oestrogen-related receptor-α (ERRα). We also observed that when we reduce the amount of STARS in muscle cells, we block the ability of PGC-1α/ERRα to increase a gene called carnitine palmitoyltransferase-1β (CPT-1β), which is important for fat metabolism. Our study has shown that the STARS pathway is regulated by endurance exercise. STARS may also play a role in fat metabolism in muscle.

Abstract

Abstract

The striated muscle activator of Rho signalling (STARS) is an actin-binding protein specifically expressed in cardiac, skeletal and smooth muscle. STARS has been suggested to provide an important link between the transduction of external stress signals to intracellular signalling pathways controlling genes involved in the maintenance of muscle function. The aims of this study were firstly, to establish if STARS, as well as members of its downstream signalling pathway, are upregulated following acute endurance cycling exercise; and secondly, to determine if STARS is a transcriptional target of peroxisome proliferator-activated receptor gamma co-activator 1-α (PGC-1α) and oestrogen-related receptor-α (ERRα). When measured 3 h post-exercise, STARS mRNA and protein levels as well as MRTF-A and serum response factor (SRF) nuclear protein content, were significantly increased by 140, 40, 40 and 40%, respectively. Known SRF target genes, carnitine palmitoyltransferase-1β (CPT-1β) and jun B proto-oncogene (JUNB), as well as the exercise-responsive genes PGC-1α mRNA and ERRα were increased by 2.3-, 1.8-, 4.5- and 2.7-fold, 3 h post-exercise. Infection of C2C12 myotubes with an adenovirus-expressing human PGC-1α resulted in a 3-fold increase in Stars mRNA, a response that was abolished following the suppression of endogenous ERRα. Over-expression of PGC-1α also increased Cpt-1β, Cox4 and Vegf mRNA by 6.2-, 2.0- and 2.0-fold, respectively. Suppression of endogenous STARS reduced basal Cpt-1β levels by 8.2-fold and inhibited the PGC-1α-induced increase in Cpt-1β mRNA. Our results show for the first time that the STARS signalling pathway is upregulated in response to acute endurance exercise. Additionally, we show in C2C12 myotubes that the STARS gene is a PGC-1α/ERRα transcriptional target. Furthermore, our results suggest a novel role of STARS in the co-ordination of PGC-1α-induced upregulation of the fat oxidative gene, CPT-1β.

Introduction

Skeletal muscle contraction is a catalyst for numerous physiological adaptations including enhanced substrate metabolism, growth and regeneration (Hawke, 2005; Hawley et al. 2006; Leger et al. 2006a; Russell, 2010). For these skeletal muscle adaptations to occur the extracellular signals must be detected, then transmitted to the cell (Schiaffino et al. 2007). These signals result in the intracellular activation of transcriptional co-activators and transcription factors (McGee et al. 2006) with a consequent increase in their target genes that encode proteins mediating metabolic, structural and contractile function of skeletal muscle (Hawley et al. 2006; Hood, 2009). Understanding the molecular targets and intracellular signals activated by exercise is important for identifying potential therapeutic targets to combat diseases characterized by perturbed skeletal muscle metabolism, growth and regeneration.

The striated muscle activator of Rho signalling (STARS), also known as myocyte stress-1 (ms1) (Mahadeva et al. 2002) and actin-binding rho activating protein (Abra) (Troidl et al. 2009), is an actin-binding protein specifically expressed in cardiac, skeletal and smooth muscle (Arai et al. 2002; Mahadeva et al. 2002; Troidl et al. 2009). STARS, in part in collaboration with RhoA, enhances the binding of free G-actin to F-actin filaments, resulting in enhanced or stabilized actin polymerization (Arai et al. 2002). As a consequence there is a reduction in free G-actin, which permits the nuclear translocation of the transcriptional co-activator myocardin-related transcription factor-A (MRTF-A) (Sotiropoulos et al. 1999). MRTF-A is a transcriptional co-activator of SRF (Miralles et al. 2003), a transcription factor known to regulate genes involved in muscle metabolism and growth (Charvet et al. 2006; Miano et al. 2007). STARS gene expression is increased in rat heart during pressure overload (Mahadeva et al. 2002) and its mRNA and proteins levels are increased in rabbit smooth muscle following fluid shear stress-induced arteriogenesis (Troidl et al. 2009). Forced over-expression of STARS causes cardiac hypertrophy in mice (Kuwahara et al. 2007) and increases the proliferation of porcine smooth muscle cells (Troidl et al. 2009). STARS mRNA, as well as several members of the STARS signalling pathway, are upregulated in hypertrophied human skeletal muscle following increased muscle loading caused by resistance exercise training and downregulated following reduced training (Lamon et al. 2009). STARS has therefore been suggested to provide an important link between the transduction of external stress to intracellular signalling pathways and the control of genes involved in the maintenance of muscle function (Arai et al. 2002; Troidl et al. 2009; Russell, 2010).

Moderate-intensity skeletal muscle contraction induced by endurance exercise has positive health implications for diseases such as diabetes, obesity and cardiovascular disease as well as maintaining skeletal function in the elderly (Booth et al. 2002; Ventura-Clapier et al. 2007; Koutroumpi et al. 2008). It has recently been observed that endurance exercise increases the phosphorylation of SRF (Rose et al. 2007), a key downstream target of the STARS signalling pathway (Arai et al. 2002; Kuwahara et al. 2005). Whether endurance exercise increases the levels of STARS, members of the STARS signalling pathway and/or downstream transcriptional targets involved in muscle metabolism and growth has not been investigated.

The STARS gene has recently been identified as a transcriptional target of MEF2 in cardiac muscle (Kuwahara et al. 2007) as well as MyoD in C2C12 myotubes (Ounzain et al. 2008). Analysis of the human STARS promoter reveals also an oestrogen-related receptor-α (ERRα) binding site, located 150 bp upstream of the transcriptional start site. ERRα, in combination with its transcriptional co-activator PGC-1α, is known to regulate genes involved in skeletal muscle function (Hock & Kralli, 2009). As both ERRα and PGC-1α are upregulated in human skeletal muscle following endurance exercise (Cartoni et al. 2005; Russell et al. 2005), it is possible that STARS is a target of the PGC-1α/ERRα transcriptional program.

Therefore the primary aim of the present study was to determine if STARS and members of its downstream signalling pathway, including MRTF-A and SRF, as well as several SRF target genes including CPT-1β, JUNB, α-actin and insulin-like growth factor-1 (IGF-1), are upregulated following acute endurance exercise. A secondary aim was to determine if the STARS gene is regulated via PGC-1α/ERRα signalling.

Methods

Subjects

Nine healthy male subjects participated in the study: age 24 ± 5 years, weight 84 ± 10 kg, single leg Inline graphic 36.6 ± 3.8 ml kg−1 min−1. The study was approved by the Deakin University Human Research Ethics Committee in accordance with the Declaration of Helsinki (2000). All participants gave their informed consent and agreed to muscle biopsies and physiological testing.

Preliminary testing

Subjects were familiarised with the electromagnetically braked cycle ergometer (Lode B.V, Groningen, The Netherlands) and single leg cycling on two separate occasions prior to a single leg cycling Inline graphic test. The Inline graphic test involved single leg cycling at three incremental work rates of 65, 80 and 95 W for 3 min followed by an increase of 15 W min−1 until exhaustion. Expired air was collected and analysed using zirconia cell O2 and infrared CO2 analysers (Applied Electrochemical Instrument, Pittsburgh, PA, USA) that were calibrated against 0.01% alpha-rated gases (Medgraphics, St Paul, MN, USA).

Experimental procedure

Subjects commenced the experimental trial 1 week after the Inline graphic test and abstained from strenuous exercise during this time. For each testing session the subjects were required to report to the laboratory having fasted overnight, and abstained from caffeine and alcohol consumption for 24 h. On the first day of the trial a resting muscle sample (pre) was obtained from the non-exercising (NEx) leg to avoid the potential influence of a stress response from the biopsy needle. With the exercising (Ex) leg subjects then performed single leg cycling at 65% single leg Inline graphic (95.9 ± 19.5 W) until exhaustion (1.2 ± 0.2 h), which was determined by the subjects’ choice to cease cycling or by the subjects’ inability to maintain the required cadence. Muscle samples were taken from the Ex leg at 3 and 24 h post-exercise.

Muscle biopsies

Skeletal muscle samples were obtained under local anaesthesia (1% Xylocaine) from the belly of the vastus lateralis muscle using a percutaneous needle biopsy technique (Bergstrom, 1962) modified to include suction (Evans et al. 1982). Following a small incision through the skin, muscle biopsies were taken using a Bergstrom needle, snap frozen and stored in liquid nitrogen until required for analysis.

RNA extraction

Total RNA was extracted from 5–20 mg skeletal muscle samples using Tri-Reagent Solution (Ambion Inc., Austin, TX, USA) according to the manufacturer's protocol. First-strand cDNA was generated from 1 μg RNA in 20 μl reaction buffer using the AMV reverse transcriptase (RT) kit (Promega, Madison, WI, USA) and Random Hexomer primers (500 ng) (Promega) according to manufacturer's protocol.

Real-time quantitative PCR

Real-time PCR was carried out using either a 7500 Real-Time PCR system (Applied Biosystems, Forster City, CA, USA) or a Stratagene MX3000 PCR system (Agilent Technologies, Santa Clara, CA, USA) to measure mRNA levels of STARS, α-actin, IGF-1, PGC-1α, ERRα, CPT-1β, cytochrome c oxidase subunit IV (COX4) and JUNB. To compensate for variations in input RNA amounts and efficiency of the reverse transcription, data were normalized to cyclophilin mRNA levels when assaying muscle biopsies, and ribosomal protein 36B4 (also known as RPLPO) RNA levels when assaying C2C12 cells. PCR primers for the mouse genes are as follows; Stars, forward TCA AAC GCC CCT TGC TCT C, reverse CGT GTT CAT CGG CCC ACT; Cpt-1β, forward ATT CTG TGC GGC CCT TAT TGG AT, reverse TTT GCC TGG GAT GCC TGT AGT GT; Cox4, forward ACT ACC CCT TGC CTG ATGTG, reverse GCC CAC AAC TGT CTT CCA TT; Vegf, forward AAG CCA GCA CAT AGG AGA GAT GA, reverse TCT TTC TTT GGT CTG CAT TCA CA. Other primer sequences and PCR conditions for the amplified genes have been published previously (Puntschart et al. 1998; Russell et al. 2003a; Cartoni et al. 2005; Leger et al. 2006b; Lamon et al. 2009).

Protein extraction

Nuclear and cytosolic proteins were extracted from skeletal muscle using the NE-PER Nuclear and Cytoplasmic extraction kit (Pierce Biotechnology, Rockford, IL, USA) as performed previously by our group (Lamon et al. 2009). Exactly 2 μl of inhibitor cocktail, containing protease and phosphatase inhibitors (Sigma, St Louis, MO, USA) at a ratio of 1:1 per 20 mg of muscle sample was added before the extraction of both cytosolic and nuclear proteins. In brief, the muscle samples were homogenised in the CER I solution provided in the NE-PER kit using a Polytron PT 1200C (Kinematica AG, Luzern, Switzerland). The following steps were performed in accordance with the manufacturer's protocol. For myotubes, total protein was extracted using 1× RIPA buffer (Millipore, North Ryde, NSW, Australia) with 1 μl ml−1 protease inhibitor cocktail (Sigma, Castle Hill, NSW, Australia) and 10 μl ml−1 Halt Phosphatase Inhibitor Single-Use Cocktail (Thermo Scientific, Rockford, IL, USA). Total protein content was determined using the BCA protein assay kit (Pierce Biotechnology, Rockford, IL, USA) according to the manufacturer's instructions.

Western blotting

Electrophoresis and protein transfer were performed using the XCell Surelock Novex Mini-Cell (Invitrogen, Carlsbad, CA, USA) system. Protein lysates were separated by SDS-PAGE using pre-cast NuPAGE Novex 4 to 12% Bis-Tris gels (Invitrogen) and transferred to a PVDF membrane (Millipore, Billerica, MA, USA). Membranes were then incubated at room temperature for 1 h in blocking buffer (0.1% Tween 20–Tris-buffered saline (TBST)) supplemented with 5% gelatin. After blocking, membranes were incubated overnight at 4°C with the following primary antibodies diluted 1:1000 in 5% gelatin–TBST: STARS (Institute of Medical and Veterinary Science, Adelaide, SA, Australia); SRF (Santa Cruz, sc-335; CA, USA); MRTF-A (Santa Cruz, sc-32909). Following washing the membranes were incubated for 1 h with horseradish peroxidase-conjugated secondary antibody diluted 1:2000 in 5% gelatin–TBST. After washing, the specific proteins were revealed using ECL-Plus (GE Healthcare, Buckinghamshire, UK) and exposed on a Kodak 2000 mm image station (Eastman Kodak, Rochester, NY, USA). Individual protein band optical densities were determined using ImageJ Software (National Institutes of Health, Bethesda, MA, USA). Tubulin (cytoplasmic fractions) and Lamin A (nuclear fractions) as well as Ponceau staining (0.5% Ponceau-S (w/v), 1% acetic acid (v/v)) were used to control for protein loading as published previously by our group (Lamon et al. 2009). A representative blot is shown in Supplementary Fig. S1. Validation of our STARS antibody using siRNA Knockdown of STARS and adenoviral over-expression of STARS is shown in Supplementary Fig. S2.

Immunofluorescence

Immunfluorescence experiments for identifying fibre-specific protein levels of STARS were performed as reported previously by our group (Russell et al. 2003b, 2004). The antibodies raised against STARS, MHC type I and MHC type IIa were diluted 1:200, 1:10 and 1:10, and visualized using a goat anti-rabbit (IgG) fluorescein isothiocyanate (FITC), donkey anti-mouse (IgM)-TRITC and a goat anti-mouse (IgGγ1)-Alexaflour 320, respectively. Negative controls, using only the secondary antibody, were also performed in parallel and on the same glass slide as the experimental samples. Sections were viewed and photographed using an Olympus IX70 fluorescent microsope and a Magna Fire camera with dedicated software. Due to freezing artifacts only 6 of the 9 subjects were analysed. Between 80 and 120 fibres were visualized per subject.

C2C12 myotubes

C2C12 myoblasts (ATCC) were plated in 12-well or 6-well tissue culture plates in complete DMEM (10% fetal bovine serum). As the cultures approached confluence (∼90% confluent), media was changed to differentiation medium (DMEM) supplemented with 2% horse serum. Differentiation media was replaced on days 2, 4 and 6.

Adenoviral infections

For siERRα experiments, myotubes were infected with multiplicity of infection of 200 (MOI 200) siERRα or vector control adenovirus on day 4 of differentiation. On day 6 of differentiation, myotubes were infected with an additional dose of MOI 200 siERRα (or vector control) and MOI 50 of PGC-1α (tagged with a FLAG epitope) or LacZ (control) expressing adenovirus. Twenty-four hours later, RNA or DNA was harvested for RT-qPCR gene expression analysis or ChIP analyses, respectively. For expression of a constitutively active ERRα (VP16-ERRα), day 6 myotubes were infected with MOI 50 of adenoviruses expressing LacZ (control) or VP16-ERRα (Schreiber et al. 2004). RNA was harvested 24 h later.

siRNA knockdown of STARS combined with PGC1α over-expression

STARS knockdown in C2C12 myotubes was achieved using 100 pmol of Stealth RNAi siRNA (target sequence: 5′-ACCAGAACGGTTGTCAGCAAGGCTT-3′; Invitrogen) over a 72 h period. Stealth RNAi™ siRNA Negative Control Hi GC (Invitrogen) was used as the control siRNA, which contains the same GC content as the target sequence. siRNA transfection of C2C12 myotubes was performed using 5 μl Lipofectamine 2000 (Invitrogen). Twenty-four hours after transfection, the C2C12 myotubes were infected with either a green fluorescent protein (GFP; control) or PGC-1α adenovirus at a MOI of 25 for a further 48 h.

Chromatin immunoprecipitation (ChIP)

ChIP was performed on adenovirus-transduced C2C12 myotubes as described previously (Cartoni et al. 2005). Briefly, C2C12 myotubes were crosslinked for 10 min at 37°C in 1% formaldehyde in PBS. After quenching, sonication to ∼500 bp fragments and pre-clearing by treatment with protein A/G sepharose, soluble chromatin was immnuoprecipitated with the following antibodies: anti-GFP (control), anti-FLAG (clone M2) for the detection of FLAG-tagged PGC-1α, or anti-ERRα (a generous gift of Dr V. Giguere (Sladek et al. 1997)). Genomic DNA (gDNA) from each immunoprecipitation was quantified by real-time PCR, using primers (TCTCCTCCAAGTGCCAGAGT and AGCTGTATTGAGGGCTACCG) to amplify an ERR response element (ERRE) 272 bp upstream of Stars, as well as primers specific to the ERR response element-containing region of the Esrra gene (positive control) and a region that lacks ERREs, distal to the Esrra promoter (negative control) (primers described in Villena et al. 2007). Data were normalized first against total genomic input and then expressed relative to levels of immunoprecipitated DNA from the ERRα negative control region (Villena et al. 2007). Values shown are the mean and standard deviation of three independent experimental replicates.

Statistics

Differences between time were determined using a one-way ANOVA. Experiments combining over-expression and knockdown of targets were analysed using a two-way ANOVA. The level of significance was set at P < 0.05. Where required, a post hoc analysis was performed using either paired or unpaired t tests with a Bonferroni adjustment.

Results

STARS mRNA and protein levels were increased by 140% and 40%, respectively (P < 0.05), when measured 3 h post-exercise, with both returning to near-basal levels 24 h post-exercise (Fig. 1). MRTF-A nuclear protein content increased by 34% (P < 0.05) 3 h post-exercise and returned to basal lines levels 24 h post-exercise. SRF nuclear protein content also increased by 39% when measured 3 h post-exercise (P < 0.05); however, it remained elevated 24 h post-exercise (P < 0.01) (Fig. 1).

Figure 1. Effect of acute endurance exercise on STARS mRNA, STARS protein, MRTF-A and SRF nuclear protein.

Figure 1

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Effect of acute endurance exercise on the levels of: A, STARS mRNA; B, STARS protein; C, MRTF-A; and D, SRF nuclear protein. Pre, before exercise; 3 h, 3 h post-exercise; 24 h, 24 h post-exercise. *P < 0.05, significantly different from the other groups, **P < 0.01, significantly different from pre.

Levels of the SRF targets genes, CPT-1β and JUNB mRNA increased by 2.3- and 1.8-fold (P < 0.05), respectively, 3 h post-exercise and returned to baseline levels 24 h post-exercise (Fig. 2). No change in other SRF target genes, α-actin or IGF-1 mRNA was observed. PGC-1α mRNA and ERRα mRNA were also increased 4.5- and 2.7-fold, respectively 3 h post-exercise and returned to baseline levels 24 h post-exercise (Fig. 2), a response mirroring that of STARS, CPT-1β and JUNB mRNA. As PGC-1α is known to transcriptionally regulate CPT-1β (Moore et al. 2003), it was of interest to determine if STARS could also be regulated via a PGC-1α/ERRα transcriptional program. Infection of C2C12 myotubes with an adenovirus-expressing human PGC-1α resulted in a 3-fold increase in Stars mRNA (P < 0.01). This was abolished when the myotubes were infected with PGC-1α as well as an adenovirus containing an siRNA sequence against ERRα. Additionally, expression of a constitutively active form of ERRα, VP16-ERRα, resulted in a 7-fold increase in Stars mRNA (P < 0.01) (Fig. 3).

Figure 2. Effect of acute endurance exercise on CPT-1β, JUNB, α-actin, IGF-1, PGC-1α and ERRα mRNA levels.

Figure 2

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Effect of acute endurance exercise on: A, CPT-1β; B, JUNB; C, α-actin; D, IGF-1; E, PGC-1α; and F, ERRα mRNA levels. Pre, before exercise; 3 h, 3 h post-exercise; 24 h, 24 h post-exercise.. Significantly different from the other groups; *P < 0.05; **P < 0.01.

Figure 3. Stars is regulated via the PGC-1α/ERRα transcriptional program in C2C12 myotubes.

Figure 3

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A, Stars mRNA levels 24 h after expression of LacZ (control) or PGC-1α, in the absence or presence of siRNA against ERRα (siERRα). N = 6 per group (duplicate samples from 3 separate experiments). B, Stars mRNA levels 24 h after expression of LacZ (control) or the constitutively active VP16-ERRα. N = 4 per group. **P < 0.01, significantly different from the other groups.

To test if ERRα and PGC-1α act directly at the Stars gene to induce its expression, we asked whether ERRα and PGC-1α interact physically with the Stars promoter. Analysis of the Stars genomic sequence indicated a putative ERRα binding site matching the known 9 bp ERRE consensus 5′-TNAAGGTCA-3′ (Sladek et al. 1997) and was conserved across species (Fig. 4A). To determine binding of endogenous ERRα and exogenous adenovirally expressed PGC-1α at this region, we immunoprecipitated crosslinked chromatin with antibodies against either GFP (control antibody measuring background), FLAG-tagged PGC-1α or ERRα, and measured the enrichment of the Stars locus DNA in the immunoprecipitated material. Both PGC-1α and ERRα were detected at the Stars ERRE (Fig. 4B), as well as at the previously defined ERRE of the Esrra gene that was used as positive control (Fig. 4C). ERRα was detected at the Stars locus only when PGC-1α was expressed, i.e. when endogenous ERRα levels were induced (Cartoni et al. 2005). Knockdown of endogenous ERRα expression by siERRα ablated binding of PGC-1α to the Stars ERRE (Fig. 4B), demonstrating a critical role for ERRα in recruitment of the PGC-1α co-activator to the Stars promoter.

Figure 4. PGC-1α and ERRα bind an ERRE upstream of Stars in C2C12 myotubes.

Figure 4

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A, a University of California, Santa Cruz (UCSC) genome browser screenshot depicting the presence of an ERRE (at 272 bp 5′ of the Stars transcription site and its conservation across species. Occupancy of PGC-1α and ERRα at the Stars (B) and Esrra (positive control) (C) ERREs, as determined by ChIP assays in C2C12 myotubes expressing LacZ (control) or PGC-1α, in the absence (Super) or presence of siRNA for ERRα (siErrα). Data are expressed relative to a region devoid of ERREs and are the mean ± SD of triplicates. **P < 0.01, significantly different from all treatments within the group.

In human skeletal muscle we have previously shown that PGC-1α protein levels are greater in oxidative type I and oxidative-glycolytic type IIa fibres than in glycolytic type IIx fibres (Russell et al. 2003a). Using immunofluorescence staining we observed that STARS protein levels are also predominately expressed in oxidative type I and oxidative-glycolytic type IIa fibres when compared with glycolytic type IIx fibres (Fig. 5A and B). STARS was also expressed in the myonuclei as shown via its co-localization with the 4,6-diamidino-2-phenylindole (DAPI) stain (Fig. 5C–E). Due to limited nuclear protein extracts STARS nuclear protein content could only be measured in 6 of the 9 subjects. Following exercise, 5 of 6 subjects showed an increase in STARS nuclear protein levels. Although the increase was by 60% this did not reach statistical significance (Fig. 5F).

Figure 5. The STARS protein is more abundantly expressed in oxidative type I and oxidative-glycolytic type IIa fibres, than in glycolytic type IIx fibres and is also found in the nucleus.

Figure 5

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Immunofluorescence staining of: A, the STARS protein; and B, MHCI (red; oxidative type I), MHCIIa (blue; oxidative-glycolytic type IIa) and MHCIIx (not stained; glycolytic type IIx). The same fibres to the marks of I, IIa or IIx in figures A and B. C, DAPI stain; D, STARS stain; and E, their co-localization, respectively, is shown in the section. Arrows represent co-localization of the nuclei and STARS. F, Western blot showing the STARS protein levels in nuclear protein fraction before, 3 h after and 24 h post-exercise.

As PGC-1α is known to regulate genes involved in mitochondrial biogenesis, angiogenesis and oxidative phosphorylation, it was of interest to see if knockdown of STARS could ablate this response. In C2C12 myotubes PGC-1α over-expression resulted in a 2.0-fold increase in Cox4 and Vegf mRNA, an effect not influenced by the knockdown of STARS. PGC-1α over-expression also increased Cpt-1β mRNA by 6.6-fold. However, the endogenous knockdown of STARS significantly reduced the basal levels of Cpt-1β mRNA levels by 8.2-fold and ablated the PGC-1α-induced increase in Cpt-1β mRNA (Fig. 6).

Figure 6. Regulation of: A, Cox4; B, Cpt-1β; and C, Vegf mRNA by PGC-1α with and without STARS knockdown in C2C12 myotubes.

Figure 6

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N = 6 per group. **P < 0.01, significantly different from the other groups.

Discussion

Identifying the molecular targets and intracellular signals activated by endurance exercise will significantly enhance our understanding of how skeletal muscle adapts to increased contractile load and may identify potential therapeutic targets to combat diseases characterized by perturbed skeletal muscle metabolism, growth and regeneration. The present study demonstrates several new and important findings, which are summarized in Fig. 7. Firstly, we show for the first time that an acute bout of single-leg endurance exercise is sufficient to induce an increase in STARS mRNA and protein levels; secondly, the response of STARS is paralleled by an increase in the nuclear protein content of several key members of the STARS signalling pathway, MRTF-A and SRF; thirdly, in C2C12 myotubes the STARS gene is a target of the PGC-1α/ERRα transcriptional complex; and finally, the presence STARS appears to be required for the PGC-1α regulation of the CPT-1β gene in C2C12 myotubes.

Figure 7. A diagram summarizing the main findings from the study.

Figure 7

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STARS is transcriptional target of PGC-1α/ERRα signalling. Acute endurance exercise increases STARS mRNA and protein levels as well as the nuclear translocation of the transcriptional co-activator MRTF-A and its target transcription factor SRF. Increased nuclear abundance of MRTF-A and SRF should increase the potential for SRF target transcription.

STARS, an actin-binding protein, is considered to communicate mechanical stress signals to downstream transcriptional regulators, such as MRTF-A and SRF, via its influence on actin dynamics (Arai et al. 2002; Kuwahara et al. 2005). SRF is known to regulate genes involved in muscle structure, function and growth such as α-actin, MHC, IGF and JUNB (Charvet et al. 2006; Lindecke et al. 2006; Miano et al. 2007) as well as fat metabolism, such as CPT-1β (Moore et al. 2001). Therefore, STARS may be an upstream regulator of signals controlling the expression of genes which encode proteins involved in muscle function, growth and metabolism. In support of this notion, forced over-expression of STARS in vivo (Kuwahara et al. 2007) and in vitro (Koekemoer et al. 2009) results in hypertrophy of cardiac tissue and protects against apoptosis. Additionally, we have recently shown that the STARS signalling pathway is increased in hypertrophied human skeletal muscle following resistance exercise (Lamon et al. 2009). In contrast, STARS and several members of its signalling pathway are reduced in skeletal muscle of sarcopenic mice (Sakuma et al. 2008) and after 1 day of limb immobilization in rats (Giger et al. 2009). However, after 20 days of limb immobilization in humans there were no observed changes in STARS (Sakuma et al. 2009). The results from the present study show that acute moderate-intensity exercise results not only in increased levels of STARS mRNA but also the STARS protein. Combined, these observations suggest that the regulation of STARS is rapid and transient which is a logical regulatory response for a protein believed to be responsible for the sensing and transmission of extracellular stress signals. It should be noted that this is the first measurement of endogenous STARS protein levels and shows, at least in the present experimental model, that the regulation of STARS mRNA is closely mirrored by the regulation of its protein.

In parallel with the increase in STARS, we show for the first time that both MRTF-A and its transcriptional target SRF have enhanced nuclear abundance following endurance exercise, a response consistent with enhanced STARS activity (Kuwahara et al. 2005). These results support our previous observation following high-intensity resistance training (Lamon et al. 2009). Along similar lines, endurance exercise training has also been shown to increase SRF activity, as shown by enhanced phosphorylation levels, in human skeletal muscle (Rose et al. 2007). In the present study, we also observed an increase in the SRF target genes CPT-1β, a regulator of long-chain fatty acid β-oxidation, and JUNB, a cell-cycle and growth regulator, but not the expression levels of α-actin and IGF-1, genes involved in structure and growth. While these results suggest that both moderate and intense exercise elicits stress signals that can be detected by STARS and transmitted downstream to its effector targets, such as MRTF-A and SRF, their transcriptional control might be dependent on the exercise intensity. For example, SRF is known to regulate over 200 genes, many of which are involved in muscle structure, function, growth and regeneration (Charvet et al. 2006; Miano et al. 2007) but not all are regulated at the same time. The selective regulation of certain SRF target genes might be attributed to their sensitivity to the stress signals initiated by the acute nature of the endurance exercise protocol.

While we show here that endurance exercise, and previously that resistance exercise (Lamon et al. 2009), regulates STARS transcription and translation, the exact stress signals initiating this are unknown. During exercise, numerous stress signals are initiated by changes in motor nerve activation and calcium concentrations mechanical and contractile stress, increased muscle blood flow and shear stress, hormonal and metabolic stress which are known to activate intracellular signalling pathways controlling skeletal muscle gene transcription and translation (Bassel-Duby & Olson, 2006; Koulmann & Bigard, 2006; Park et al. 2007; Favier et al. 2008; Russell, 2010). Identifying the primary stressor/s activated by endurance exercise which regulate STARS, and consequently its downstream signalling pathway, is beyond the scope of this study; however, this should be an area investigated in the future.

Our observation that STARS is increased in response to moderate-intensity endurance exercise suggests that the STARS signalling pathway may also play a role in skeletal muscle adaptations associated with endurance exercise such as substrate oxidation, capillary function and mitochondrial biogenesis, factors often perturbed in metabolic diseases (Schrauwen & Hesselink, 2004). Indeed SRF, a downstream target of STARS signalling, has also been shown to control the transcription of CPT-1β (Moore et al. 2001), a regulator of fat metabolism, as well as vascular endothelial growth factor (VEGF), a regulator of angiogenesis (Franco et al. 2008). Another commonalty between CPT-1β and VEGF is that they can also be regulated by the transcriptional co-activator PGC-1α (Moore et al. 2003; Zhang et al. 2011). PGC-1α is upregulated in human skeletal muscle in response to both moderate- and high-intensity endurance exercise (Russell et al. 2003a, 2005; Cartoni et al. 2005). PGC-1α co-activates several transcription factors that regulate genes involved in skeletal muscle substrate oxidation and mitochondrial biogenesis (Knutti & Kralli, 2001; Hock & Kralli, 2009). We identified that one of these transcription factors, ERRα, has a binding site on the STARS promoter. Therefore, we performed in vitro experiments using C2C12 myotubes whereby we manipulated PGC-1α and ERRα levels using adenoviral infection. Our results demonstrate that STARS is a transcriptional target of the PGC-1α/ERRα transcriptional program.

Previous work by our group has shown that in human skeletal muscle, PGC-1α protein levels are greater in oxidative type I and oxidative-glycolytic type IIa fibres than in glycolytic type IIx fibres (Russell et al. 2003a). In the present study, we also observed a similar fibre type expression for the STARS protein. In addition to this, we observed that the STARS protein was also localized to the nucleus, which supports observations made in transiently transfected COS cells (Arai et al. 2002) and in rat endothelial cells (Troidl et al. 2009). The PGC-1α regulation of STARS, their similar fibre type expression profile, as well as the localization of STARS in the nucleus, suggested to us that a PGC-1α/STARS transcriptional relationship may exist. Our observation that the knockdown of endogenous STARS reduces CPT-1β mRNA and attenuates the PGC-1α upregulation of CPT-1β mRNA supports this possibility. However, it remains to be determined whether STARS has the capacity to directly bind DNA or whether its downregulation impacts indirectly on transcriptional signalling pathways, potentially through SRF. These in vitro results, combined with our observation of increases in PGC-1α and ERRα mRNA following acute endurance exercise, suggest that a possible feed-forward loop involving SRF, PGC-1α, ERRα and STARS may exist during moderate-intensity muscle contraction which controls oxidative metabolism and muscle function. In contrast to the increase in these genes following exercise, skeletal muscle SRF, PGC-1α and STARS are all reduced in conditions such as ageing (Ling et al. 2004; Lahoute et al. 2008; Sakuma et al. 2008) and limb immobilization (Oishi et al. 2008; Giger et al. 2009), situations that present reduced skeletal muscle mass, function and attenuated metabolism. The possible existence of this novel transcriptional pathway requires further investigation.

In conclusion, we show for the first time that STARS mRNA and protein as well as MRTF-A and SRF nuclear content are increased following endurance exercise. Additionally, we have identified the STARS gene as a PGC-1α/ERRα transcriptional target and a gene that may be involved in the regulation of oxidative metabolism. Future investigations should focus on STARS and its downstream signalling pathway on the regulation of skeletal muscle oxidative function, potentially via its influence on actin polymerization.

Acknowledgments

This work was funded in part by a Faculty Research Development Grant to A.P.R. M.A.W. is supported by an Australia Postgraduate Award; A.P.R. is supported by an NH and MRC Biomedical Career Development Award (479536); M.B.H., B.C.H. and A.K. were supported by an NIH/NIDDK grant (DK064951). We wish to thank Kenny Watt for his role in the recruitment and testing of the subjects and Marin Gantner for assistance with the ChIP experiments.

Glossary

Abbreviations

Abra

actin-binding rho activating protein

CPT-1β

carnitine palmitoyltransferase 1β

COX4

cytochrome c oxidase subunit IV

DAPI

4,6-diamidino-2-phenylindole

ERRα

oestrogen-related receptor-α

IGF-1

insulin-like growth factor-1

JUNB

jun B proto-oncogene

MRTF-A

myocardin-related transcription factor-A

ms1

myocyte stress-1

MHCI

myosin heavy chain I

MHCIIa

myosin heavy chain IIa

PGC-1α

peroxisome proliferator-activated receptor gamma co-activator-1α

SRF

serum response factor

STARS

striated muscle activator of rho signalling

VEGF

vascular endothelial growth factor

Author contributions

All in vivo experiments and in vitro experiments required for the antibody validation were performed at Deakin University while the in vitro experiments completed in Figs 5 and 6 were performed at the Scripps Research Institute. M.A.W. completed the analyses using of human muscle samples, validated the STARS antibody, performed the PGC-1/siSTARS experiments and assisted with the manuscript preparation; M.B.H. and B.C.H. completed the adenoviral over expression, siRNA and ChIP experiments and participated with the manuscript preparation; A.K. was responsible for designing the in vitro experiments and manuscript preparation; R.J.S. was responsible for designing the in vivo human exercise study and manuscript preparation; A.P.R. was responsible for management of the project, supervision of M.A.W., funding the project and manuscript preparation. All authors approved the final version of the manuscript.

Author's present address

M. Benjamin Hock: Department of Clinical Immunology, Amgen Inc., Thousand Oaks, CA, USA.

Supplementary material

Suppl. Fig. 1

Suppl. Fig. 2

Preparation of human primary myotubes

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References

  1. Arai A, Spencer JA, Olson EN. STARS, a striated muscle activator of Rho signaling and serum response factor-dependent transcription. J Biol Chem. 2002;277:24453–24459. doi: 10.1074/jbc.M202216200. [DOI] [PubMed] [Google Scholar]
  2. Bassel-Duby R, Olson EN. Signaling pathways in skeletal muscle remodeling. Annu Rev Biochem. 2006;75:19–37. doi: 10.1146/annurev.biochem.75.103004.142622. [DOI] [PubMed] [Google Scholar]
  3. Bergstrom J. Muscle electrolytes in man. Scand J Clin Invest Suppl. 1962;14:1–110. [Google Scholar]
  4. Booth FW, Chakravarthy MV, Spangenburg EE. Exercise and gene expression: physiological regulation of the human genome through physical activity. J Physiol. 2002;543:399–411. doi: 10.1113/jphysiol.2002.019265. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Cartoni R, Leger B, Hock MB, Praz M, Crettenand A, Pich S, et al. Mitofusins 1/2 and ERRα expression are increased in human skeletal muscle after physical exercise. J Physiol. 2005;567:349–358. doi: 10.1113/jphysiol.2005.092031. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Charvet C, Houbron C, Parlakian A, Giordani J, Lahoute C, Bertrand A, et al. New role for serum response factor in postnatal skeletal muscle growth and regeneration via the interleukin 4 and insulin-like growth factor 1 pathways. Mol Cell Biol. 2006;26:6664–6674. doi: 10.1128/MCB.00138-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Evans WJ, Phinney SD, Young VR. Suction applied to a muscle biopsy maximizes sample size. Med Sci Sports Exerc. 1982;14:101–102. [PubMed] [Google Scholar]
  8. Favier FB, Benoit H, Freyssenet D. Cellular and molecular events controlling skeletal muscle mass in response to altered use. Pflugers Arch. 2008;456:587–600. doi: 10.1007/s00424-007-0423-z. [DOI] [PubMed] [Google Scholar]
  9. Franco CA, Mericskay M, Parlakian A, Gary-Bobo G, Gao-Li J, Paulin D, Gustafsson E, Li Z. Serum response factor is required for sprouting angiogenesis and vascular integrity. Dev Cell. 2008;15:448–461. doi: 10.1016/j.devcel.2008.07.019. [DOI] [PubMed] [Google Scholar]
  10. Giger JM, Bodell PW, Zeng M, Baldwin KM, Haddad F. Rapid muscle atrophy response to unloading: pretranslational processes involving MHC and actin. J Appl Physiol. 2009;107:1204–1212. doi: 10.1152/japplphysiol.00344.2009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Hawke TJ. Muscle stem cells and exercise training. Exerc Sport Sci Rev. 2005;33:63–68. doi: 10.1097/00003677-200504000-00002. [DOI] [PubMed] [Google Scholar]
  12. Hawley JA, Hargreaves M, Zierath JR. Signalling mechanisms in skeletal muscle: role in substrate selection and muscle adaptation. Essays Biochem. 2006;42:1–12. doi: 10.1042/bse0420001. [DOI] [PubMed] [Google Scholar]
  13. Hock MB, Kralli A. Transcriptional control of mitochondrial biogenesis and function. Annu Rev Physiol. 2009;71:177–203. doi: 10.1146/annurev.physiol.010908.163119. [DOI] [PubMed] [Google Scholar]
  14. Hood DA. Mechanisms of exercise-induced mitochondrial biogenesis in skeletal muscle. Appl Physiol Nutr Metab. 2009;34:465–472. doi: 10.1139/H09-045. [DOI] [PubMed] [Google Scholar]
  15. Knutti D, Kralli A. PGC-1, a versatile coactivator. Trends Endocrinol Metab. 2001;12:360–365. doi: 10.1016/s1043-2760(01)00457-x. [DOI] [PubMed] [Google Scholar]
  16. Koekemoer AL. Myocyte stress 1 plays an important role in cellular hypertrophy and protection against apoptosis. FEBS Lett. 2009;583:2964–2967. doi: 10.1016/j.febslet.2009.08.011. [DOI] [PubMed] [Google Scholar]
  17. Koulmann N, Bigard AX. Interaction between signalling pathways involved in skeletal muscle responses to endurance exercise. Pflugers Arch. 2006;452:125–139. doi: 10.1007/s00424-005-0030-9. [DOI] [PubMed] [Google Scholar]
  18. Koutroumpi M, Pitsavos C, Stefanadis C. The role of exercise in cardiovascular rehabilitation: a review. Acta Cardiol. 2008;63:73–79. doi: 10.2143/AC.63.1.2025335. [DOI] [PubMed] [Google Scholar]
  19. Kuwahara K, Barrientos T, Pipes GC, Li S, Olson EN. Muscle-specific signaling mechanism that links actin dynamics to serum response factor. Mol Cell Biol. 2005;25:3173–3181. doi: 10.1128/MCB.25.8.3173-3181.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Kuwahara K, Teg Pipes GC, McAnally J, Richardson JA, Hill JA, Bassel-Duby R, Olson EN. Modulation of adverse cardiac remodeling by STARS, a mediator of MEF2 signaling and SRF activity. J Clin Invest. 2007;117:1324–1334. doi: 10.1172/JCI31240. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Lahoute C, Sotiropoulos A, Favier M, Guillet-Deniau I, Charvet C, Ferry A, et al. Premature aging in skeletal muscle lacking serum response factor. PLoS One. 2008;3:e3910. doi: 10.1371/journal.pone.0003910. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Lamon S, Wallace MA, Leger B, Russell AP. Regulation of STARS and its downstream targets suggest a novel pathway involved in human skeletal muscle hypertrophy and atrophy. J Physiol. 2009;587:1795–1803. doi: 10.1113/jphysiol.2009.168674. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Leger B, Cartoni R, Praz M, Lamon S, Deriaz O, Crettenand A, et al. Akt signalling through GSK-3β, mTOR and Foxo1 is involved in human skeletal muscle hypertrophy and atrophy. J Physiol. 2006a;576:923–933. doi: 10.1113/jphysiol.2006.116715. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Leger B, Vergani L, Soraru G, Hespel P, Derave W, Gobelet C, et al. Human skeletal muscle atrophy in amyotrophic lateral sclerosis reveals a reduction in Akt and an increase in atrogin-1. FASEB J. 2006b;20:583–585. doi: 10.1096/fj.05-5249fje. [DOI] [PubMed] [Google Scholar]
  25. Lindecke A, Korte M, Zagrebelsky M, Horejschi V, Elvers M, Widera D, et al. Long-term depression activates transcription of immediate early transcription factor genes: involvement of serum response factor/Elk-1. Eur J Neurosci. 2006;24:555–563. doi: 10.1111/j.1460-9568.2006.04909.x. [DOI] [PubMed] [Google Scholar]
  26. Ling C, Poulsen P, Carlsson E, Ridderstrale M, Almgren P, Wojtaszewski J, et al. Multiple environmental and genetic factors influence skeletal muscle PGC-1α and PGC-1β gene expression in twins. J Clin Invest. 2004;114:1518–1526. doi: 10.1172/JCI21889. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. McGee SL, Sparling D, Olson AL, Hargreaves M. Exercise increases MEF2- and GEF DNA-binding activity in human skeletal muscle. FASEB J. 2006;20:348–349. doi: 10.1096/fj.05-4671fje. [DOI] [PubMed] [Google Scholar]
  28. Mahadeva H, Brooks G, Lodwick D, Chong NW, Samani NJ. ms1, a novel stress-responsive, muscle-specific gene that is up-regulated in the early stages of pressure overload-induced left ventricular hypertrophy. FEBS Lett. 2002;521:100–104. doi: 10.1016/s0014-5793(02)02833-8. [DOI] [PubMed] [Google Scholar]
  29. Miano JM, Long X, Fujiwara K. Serum response factor: master regulator of the actin cytoskeleton and contractile apparatus. Am J Physiol Cell Physiol. 2007;292:C70–C81. doi: 10.1152/ajpcell.00386.2006. [DOI] [PubMed] [Google Scholar]
  30. Miralles F, Posern G, Zaromytidou AI, Treisman R. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell. 2003;113:329–342. doi: 10.1016/s0092-8674(03)00278-2. [DOI] [PubMed] [Google Scholar]
  31. Moore ML, Park EA, McMillin JB. Upstream stimulatory factor represses the induction of carnitine palmitoyltransferase-Ibeta expression by PGC-1. J Biol Chem. 2003;278:17263–17268. doi: 10.1074/jbc.M210486200. [DOI] [PubMed] [Google Scholar]
  32. Moore ML, Wang GL, Belaguli NS, Schwartz RJ, McMillin JB. GATA-4 and serum response factor regulate transcription of the muscle-specific carnitine palmitoyltransferase Iβ in rat heart. J Biol Chem. 2001;276:1026–1033. doi: 10.1074/jbc.M009352200. [DOI] [PubMed] [Google Scholar]
  33. Oishi Y, Ogata T, Yamamoto KI, Terada M, Ohira T, Ohira Y, et al. Cellular adaptations in soleus muscle during recovery after hindlimb unloading. Acta Physiol (Oxf) 2008;192:381–395. doi: 10.1111/j.1748-1716.2007.01747.x. [DOI] [PubMed] [Google Scholar]
  34. Ounzain S, Dacwag CS, Samani NJ, Imbalzano AN, Chong NW. Comparative in silico analysis identifies bona fide MyoD binding sites within the myocyte stress 1 gene promoter. BMC Mol Biol. 2008;9:50. doi: 10.1186/1471-2199-9-50. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Park JY, Farrance IK, Fenty NM, Hagberg JM, Roth SM, Mosser DM, et al. NFKB1 promoter variation implicates shear-induced NOS3 gene expression and endothelial function in prehypertensives and stage I hypertensives. Am J Physiol Heart Circ Physiol. 2007;293:H2320–H2327. doi: 10.1152/ajpheart.00186.2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Puntschart A, Wey E, Jostarndt K, Vogt M, Wittwer M, Widmer HR, et al. Expression of fos and jun genes in human skeletal muscle after exercise. Am J Physiol Cell Physiol. 1998;274:C129–C137. doi: 10.1152/ajpcell.1998.274.1.C129. [DOI] [PubMed] [Google Scholar]
  37. Rose AJ, Frosig C, Kiens B, Wojtaszewski JF, Richter EA. Effect of endurance exercise training on Ca2+ calmodulin-dependent protein kinase II expression and signalling in skeletal muscle of humans. J Physiol. 2007;583:785–795. doi: 10.1113/jphysiol.2007.138529. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Russell AP. The molecular regulation of skeletal muscle mass. Clin Exp Pharmacol Physiol. 2010;37:378–384. doi: 10.1111/j.1440-1681.2009.05265.x. [DOI] [PubMed] [Google Scholar]
  39. Russell AP, Feilchenfeldt J, Schreiber S, Praz M, Crettenand A, Gobelet C, et al. Endurance training in humans leads to fiber type-specific increases in levels of peroxisome proliferator-activated receptor-γ coactivator-1 and peroxisome proliferator-activated receptor-α in skeletal muscle. Diabetes. 2003a;52:2874–2881. doi: 10.2337/diabetes.52.12.2874. [DOI] [PubMed] [Google Scholar]
  40. Russell AP, Hesselink MK, Lo SK, Schrauwen P. Regulation of metabolic transcriptional co-activators and transcription factors with acute exercise. FASEB J. 2005;19:986–988. doi: 10.1096/fj.04-3168fje. [DOI] [PubMed] [Google Scholar]
  41. Russell AP, Somm E, Debigare R, Hartley O, Richard D, Gastaldi G, et al. COPD results in a reduction in UCP3 long mRNA and UCP3 protein content in types I and IIa skeletal muscle fibers. J Cardiopulm Rehabil. 2004;24:332–339. doi: 10.1097/00008483-200409000-00009. [DOI] [PubMed] [Google Scholar]
  42. Russell AP, Wadley G, Hesselink MK, Schaart G, Lo S, Leger B, et al. UCP3 protein expression is lower in type I, IIa and IIx muscle fiber types of endurance-trained compared to untrained subjects. Pflugers Arch. 2003b;445:563–569. doi: 10.1007/s00424-002-0943-5. [DOI] [PubMed] [Google Scholar]
  43. Sakuma K, Akiho M, Nakashima H, Akima H, Yasuhara M. Age-related reductions in expression of serum response factor and myocardin-related transcription factor A in mouse skeletal muscles. Biochim Biophys Acta. 2008;1782:453–461. doi: 10.1016/j.bbadis.2008.03.008. [DOI] [PubMed] [Google Scholar]
  44. Sakuma K, Watanabe K, Hotta N, Koike T, Ishida K, Katayama K, Akima H. The adaptive responses in several mediators linked with hypertrophy and atrophy of skeletal muscle after lower limb unloading in humans. Acta Physiol (Oxf) 2009;197:151–159. doi: 10.1111/j.1748-1716.2009.01995.x. [DOI] [PubMed] [Google Scholar]
  45. Schiaffino S, Sandri M, Murgia M. Activity-dependent signaling pathways controlling muscle diversity and plasticity. Physiology (Bethesda) 2007;22:269–278. doi: 10.1152/physiol.00009.2007. [DOI] [PubMed] [Google Scholar]
  46. Schrauwen P, Hesselink MK. Oxidative capacity, lipotoxicity, and mitochondrial damage in type 2 diabetes. Diabetes. 2004;53:1412–1417. doi: 10.2337/diabetes.53.6.1412. [DOI] [PubMed] [Google Scholar]
  47. Schreiber SN, Emter R, Hock MB, Knutti D, Cardenas J, Podvinec M, et al. The estrogen-related receptor alpha (ERRα) functions in PPARgamma coactivator 1alpha (PGC-1α)-induced mitochondrial biogenesis. Proc Natl Acad Sci U S A. 2004;101:6472–6477. doi: 10.1073/pnas.0308686101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Sladek R, Beatty B, Squire J, Copeland NG, Gilbert DJ, Jenkins NA, Giguere V. Chromosomal mapping of the human and murine orphan receptors ERRα (ESRRA) and ERRβ (ESRRB) and identification of a novel human ERRα-related pseudogene. Genomics. 1997;45:320–326. doi: 10.1006/geno.1997.4939. [DOI] [PubMed] [Google Scholar]
  49. Sotiropoulos A, Gineitis D, Copeland J, Treisman R. Signal-regulated activation of serum response factor is mediated by changes in actin dynamics. Cell. 1999;98:159–169. doi: 10.1016/s0092-8674(00)81011-9. [DOI] [PubMed] [Google Scholar]
  50. Troidl K, Ruding I, Cai WJ, Mucke Y, Grossekettler L, Piotrowska I, et al. Actin-binding rho activating protein (Abra) is essential for fluid shear stress-induced arteriogenesis. Arterioscler Thromb Vasc Biol. 2009;29:2093–2101. doi: 10.1161/ATVBAHA.109.195305. [DOI] [PubMed] [Google Scholar]
  51. Ventura-Clapier R, Mettauer B, Bigard X. Beneficial effects of endurance training on cardiac and skeletal muscle energy metabolism in heart failure. Cardiovasc Res. 2007;73:10–18. doi: 10.1016/j.cardiores.2006.09.003. [DOI] [PubMed] [Google Scholar]
  52. Villena JA, Hock MB, Chang WY, Barcas JE, Giguere V, Kralli A. Orphan nuclear receptor estrogen-related receptor alpha is essential for adaptive thermogenesis. Proc Natl Acad Sci U S A. 2007;104:1418–1423. doi: 10.1073/pnas.0607696104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Zhang K, Lu J, Mori T, Smith-Powell L, Synold TW, Chen S, Wen W. Baicalin increases VEGF expression and angiogenesis by activating the ERRα/PGC-1α pathway. Cardiovasc Res. 2011;89:426–435. doi: 10.1093/cvr/cvq296. [DOI] [PMC free article] [PubMed] [Google Scholar]

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