Myostatin inhibits proliferation and insulin-stimulated ...

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Myostatin inhibits proliferation and insulin-stimulated glucose uptake in mouse liver cells

Rani Watts1, Mostafa Ghozlan2, Curtis C Hughey2, Virginia L Johnsen1, Jane Shearer2 and Dustin S Hittel2 1

Faculty of Kinesiology, University of Calgary

2

Department of Biochemistry and Molecular Biology, Faculty of Medicine, University of

Calgary

Running Title: Myostatin and liver cell growth and metabolism

Corresponding author:

Dustin Hittel, PhD, Faculty of Kinesiology, University of Calgary, 2500 University Dr.

Calgary, Alberta, Canada, T2N1N4

Tel: +1-403-220-3497

Fax: +1-403-284-3553

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E-mail: [email protected]

Abstract

Although myostatin functions primarily as a negative regulator of skeletal muscle growth and development, accumulating biological and epidemiological evidence indicates an important contributing role in liver disease. In this study we demonstrate that myostatin suppresses the proliferation of Hepa-1C1c7 murine-derived liver cells (50%; P < 0.001) in part by reducing the expression of the cyclins and cyclin-dependent kinases that elicit G1-S phase transition of the cell cycle (P < 0.001). Furthermore, RT-PCR based quantification of the long noncoding RNA Malat1, recently identified as a myostatin-responsive transcript in skeletal muscle, revealed a significant down-regulation (25% and 50% respectively, P < 0.05) in the livers of myostatintreated mice and liver cells. The importance of Malat1 in liver cell proliferation was confirmed via arrested liver cell proliferation (P < 0.05) in response to partial Malat1 siRNA-mediated knockdown. Myostatin also significantly blunted insulin-stimulated glucose uptake and Akt phosphorylation in liver cells while increasing the phosphorylation of Myristoylated AlanineRich Kinase C Substrate (MARCKS), a protein that is essential for cancer cell proliferation and insulin-stimulated glucose transport. Together, these findings reveal a plausible mechanism by which circulating myostatin contributes to the diminished regenerative capacity of the liver and diseases characterized by liver insulin resistance.

Keywords: hepatogenesis; glucose metabolism; myostatin; Malat1 siRNA; cell cycle

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Introduction Myostatin is a secreted member of the transforming growth factor-β (TGF-β) family that acts as a negative regulator of skeletal muscle growth and development (Rios et al. 2004) by suppressing satellite cell activation (McCroskery et al. 2003; McFarland et al. 2006), myoblast proliferation (McFarland et al. 2006; Taylor et al. 2001; Thomas et al. 2000) and myofiber hypertrophy (Allen et al. 2011). Whereas the antagonism or inhibition of myostatin signaling induces striking skeletal muscle hypertrophy (McCroskery et al. 2005; Siriett et al. 2007), systemic over-expression leads to dramatic losses in both skeletal muscle and adipose tissue mass (Zimmers et al. 2002). Myostatin-deficient mice also exhibit reduced adiposity and resistance to dietary-induced obesity (McPherron and Lee 2002) due to increased muscle mass and increased thermogenesis through the activation of brown adipose tissue (Zhang et al. 2012). In addition, myostatin expression has been demonstrated in placenta, heart, aorta, kidney (Guo et al. 2013; Jiao et al. 2011) and pancreatic islets (Bonomi et al. 2012), yet studies investigating its influence over the growth and metabolism of non-muscle tissues are limited. Such observations, as well as the demonstration of increased skeletal muscle and plasma levels of myostatin with obesity and insulin resistance (Hittel et al. 2009) highlight an important, yet poorly understood, role for myostatin in metabolic homeostasis. The ability of the liver to regenerate is critical for survival. A distinctive feature of the liver is its ability to undergo a significant phase of compensatory growth following cell loss through trauma, infection or hepatoxic injury (Chen et al. 2000). Extensive cell proliferation via

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re-entry of hepatocytes into the G1 phase of the cell cycle from quiescence is essential to restore liver mass and necessitates co-operative signaling between a variety of cellular pathways (Chauhan et al. 2011). In contrast to the large number of hepatotropic factors that are known, few negative regulators of liver cell proliferation have been identified or well characterized. Recently, our lab and others have demonstrated that exogenous myostatin administration results in impaired insulin signaling in the livers of mice, and up-regulated lipogenic genes and increased fat accumulation in liver cells (Guo et al. 2013; Hittel et al. 2010; Wilkes et al. 2009). Furthermore, myostatin inhibitor-treated adult mice displayed significantly reduced hepatic expression of lipogenic genes and improvement in liver insulin sensitivity (Guo et al. 2013). Together these studies suggest that the liver is indeed a target for myostatin and that this may have important health implications given that elevated circulating myostatin has been observed in patients with liver disease (Garcia et al. 2010). To determine the influence of myostatin over liver cell proliferation and glucose metabolism we exposed cells of the C57-derived Hepa-1C1c7 mouse hepatoma line and found that myostatin regulates G1- to S-phase transition through a reduction in cyclin dependent kinases (Cdks) 2 and 6, cyclin A2 and cyclin D1 mRNA transcripts. In addition, the long noncoding mRNA Malat1, which plays a role in cancer, myogenesis and type-2 diabetes (Luo et al. 2006; Mirza et al. 2013; Watts et al. 2013), was shown to decrease in the livers and liver cells derived from C57 mice following myostatin exposure. Additionally, Hepa-1C1c7 cells treated with myostatin exhibited significantly reduced insulin-stimulated glucose uptake and AKT phosphorylation, confirming the insulin-desensitizing effects of myostatin on liver. Finally, we identify Myristoylated Alanine-Rich Kinase C substrate (MARCKS) (Chappell et al. 2009) as a novel downstream target of both myostatin and insulin signaling in the liver with a potential role

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in the myostatin-induced suppression of insulin-stimulated AKT phosphorylation and glucose uptake.

Materials and Methods Animal Experimentation Procedures were approved by the University of Calgary Animal Care and Use Committee and abide by the Canadian Association for Laboratory Animal Science guidelines for animal experimentation. Animals were maintained in a humidity-controlled room with a 12 h light:dark cycle. Following weaning at three weeks of age, male C57BL/6J littermates were randomly segregated into two groups (n=10 each) and maintained in microisolator cages for one week. Following this acclimation period, treated animals received 15 µg/kg/day of recombinant (rh) myostatin (R&D Systems, Minneapolis, MN) whilst the control group received 100 µl phosphate-buffered saline (PBS) as described previously (Hittel et al. 2010). Mice were euthanized on the sixth day using pentobarbital and cervical dislocation. The liver was removed, immediately frozen in liquid nitrogen and stored at -80°C for subsequent analysis. Cell Culture Hepa-1C1c7 mouse hepatoma cells (ATCC CRL-2026) were purchased from American Type Culture Collection (Manassas, VA) and grown in α-MEM (Gibco, CA) supplemented with 10% fetal bovine serum (FBS) (Gibco) and 1% antibiotic/antimycotic solution (Sigma, ON). C2C12 mouse myoblasts (ATCC CRL1772) were obtained from the American Type Culture Collection and maintained in high- glucose DMEM supplemented with 10% FBS and 1% antibiotic

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antimycotic solution. Cells were cultured in T-75 culture flasks and maintained at 37°C in a humidified incubator in 5% CO2. Culture medium was changed every 2 or 3 days, and cells were subcultured when 80% confluent. Cell Proliferation Assay Cell proliferation was measured using the Quick Cell proliferation Assay Kit (Biovision Inc, Mountain View, CA). Cells were cultured at a density of 2.5 X 103 cells per well in a flatbottomed 96-well plate and incubated in transfection media. Following 24 and 48 h treatment, WST1/ECS solution was added to each well according to manufacturer’s instructions. Cell viability was determined by measuring the absorbance at 450 nm. All assays were performed in quadruplicate and independently repeated twice. As a blank control, 100 µl α-MEM was used. Flow Cytometry Cell cycle and cell viability analysis was performed according to manufacturer’s instructions. Briefly, 3 x 105 cells were cultured as described above in 100 mm petri dishes with or without myostatin treatment for 48 h. Cells were harvested using 0.05% trypsin/EDTA and centrifuged at 450 g to pellet cells. Cells were resuspended in 1 ml media and cell counting was performed using the Guava ViaCount assay (Millipore, Hayward, CA) on CytoSoft software. Cells were resuspended at a density 5 x 105-1 x 106 cells/well and cell cycle analysis performed using Guava Cell Cycle reagent according to the manufacturers’ instructions (Millipore). Total RNA extraction from cells Cells were washed twice with 1X PBS before total RNA was extracted with TRIzol (Invitrogen, Carlsbad, CA) using the standard phenol/chloroform extraction procedure following

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manufacturer’s instructions. Total RNA was further purified using the PureLink RNA Mini Kit (Ambion, Carlsbad, CA) following manufacturer’s instructions, then quantified via spectrophotometry (Nanodrop, Wilmington, DE). Quantitative RT-PCR Total RNA was synthesized into cDNA using High Capacity RNA-to-cDNA Master Mix according the manufacturer’s instructions (Applied Biosystems, Foster City, CA). Reactions were run in 96-well plates with all cDNA samples from each treatment condition run in duplicate for each gene of interest. Samples were analyzed by real-time PCR using primers designed on Oligo Perfect Designer software (Table 1) on a CFX96 Real-Time System device (Bio-Rad, Hercules, CA) using 12.5 µl IQ SyBR Green Supermix (Bio-Rad) per 20 µl reaction volume. Fluorescent emission data were analyzed for the critical threshold (CT) values, with the expression of the gene of interest normalized to β-actin and expressed as 2-∆CT. 2-[14C]Deoxyglucose Uptake Hepa-1C1c7 were grown in 24-well plates to approximately 90% confluency then serum starved for 4 h and pre-incubated with or without 100 ng/ml rh myostatin for 1 h. Cells were incubated in cell buffer at pH 7.4 (125 mM NaCl, 5 mM KCl, 1.8 mM CaCl2, 2.6 mM MgSO4, 2 mMC3H3NaO3, 25 mM HEPES, 2% BSA) for 10 min with or without 100 nM insulin at 37°C. Glucose uptake was assayed by incubating cells with 2-[14C]deoxy-D-glucose (1 µCi/ml) for 20 min. Intracellular

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C was determined by lysing the cells with 0.5% SDS, followed by liquid

scintillation counting. Total cellular protein was determined using the bicinchoninic acid (BCA)based protein assay kit (Pierce Biotechnology, IL) according to manufacturer’s instructions and results calculated as 2-[14C]deoxy-D-glucose uptake per µg total cellular protein.

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Myostatin Phosphoproteome Characterization Hepa-1C1c7 cells were labeled for SILAC analysis as described previously using a SILAC Phosphoprotein ID and Quantification Kit (Life Technologies) (Hittel et al. 2009). Briefly, cells were cultured in RPMI-1640 with 10% (v/v) FBS, 100 units/mL of penicillin and 100 µg/mL streptomycin. For SILAC labeling, cells were cultured in RPMI lacking lysine or arginine supplemented with 200 mg/L 13C6, 15N4 arginine, 40 mg/L 13C6, 15N2 lysine and 10% dialyzed FBS for eight doublings. For myostatin experiments, confluent cells (Both Labeled and Unlabeled) were serum-starved for 4 hours in SILAC media and then treated with 4 ug/mL rh myostatin or vehicle control in fresh serum-free SILAC media for 1 hour. Cells were then washed and re-suspended with ice-cold PBS containing 1× phosphatase inhibitors (Sigma) and centrifuged at 1500 RPM for 5 min at 4°C. Pellets were then weighed, mixed 1:1 (wet weight Myostatin:Control treated) and re-suspended in 8–10 mL Phosphoprotein Lysis Buffer (Pierce), and enriched for phosphoprotein using a Pierce phosphoprotein enrichment kit according to the manufacturer’s instructions. The phosphoprotein enriched fraction was then quantified, separated by SDS-PAGE, bands visualized and excised using Coomassie G-250 and trypsin digested and identified as previously described (Hittel et al. 2009). Briefly, peptide spectra were obtained from an LTQ-Orbitrap-XL (Thermo Fishe rScientific) Mass Spectrometer and then protein identification and quantification using Integrated Proteomics Pipeline (IP2) version 1.01 software developed by Integrated Proteomics Applications, Inc. A curated list of differentially

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phosphorylated proteins was created from proteins whose phosphorylation state increased at least 1.5 fold in both SILAC labeled and unlabeled cells.

Western Blotting Cells were washed twice with 1X PBS before lysis with Cell Lysis Buffer (Cell Signaling Technology, Danvers, MA) supplemented with 1 mM phenylmethylsulphonyl fluoride (PMSF). Extracts were incubated with continuous rotation at 4°C for 1 h then centrifuged at 13,000 rpm at 4°C for 15 min to pellet cell debris. The supernatant was analyzed for total protein and then equal concentrations of denatured total protein were separated by sodium dodecyl sulphatepolyacrylamide gel electrophoresis and transferred onto nitrocellulose membranes. Membranes were blocked 5% BSA in Tris-buffered saline (50 mM Tris-HCl, 750 mM NaCl) with 0.1% Tween 20 (TBST) for 90 min at room temperature, then incubated overnight at 4°C with phospho-Smad3 (Ser 423/425) or total Smad3 (Cell Signaling, Danvers, MA), pAkt (Thr308) or total Akt (Cell Signaling) as well as B-Actin and the endogenous myostatin receptor ActRIIB (Abcam, Cambridge, UK) to control for loading, followed by incubation with horseradish peroxidase-conjugated secondary antibodies. All signals were detected using enhanced chemiluminescence substrate (ThermoScientific, Rockford, IL). Images were captured using the Chemi Genius2 Bio Imaging System (SynGene, Frederick, MD) and densitometric analysis was performed on bands using GeneTools (SynGene). Malat1 Knockdown A siRNA targeting murine Malat1 was purchased from Life Technologies. Strand sequences (with dTdT overhang) are listed in the 5’ to 3’ direction as follows: sense-

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CUUAUCAAUUCACCAAGGATT;

antisense-

UCCUUGGUGAAUUGAUAAGTA.

A

universal control siRNA was purchased from Sigma (St Louis, MO) as a scrambled sequence with no homology within the mouse or human genomes. siRNA duplexes were transfected using 2 µl/ml Lipofectamine siRNAMAX (Life Technologies) according to the manufacturer’s instructions and as described previously (Watts et al. 2013). Briefly, proliferating Hepa-1C1c7 cells were transfected with 0.5 µM siRNA upon propagation onto plastic ware, where they remained for up to 72 h. Glycogen Quantification Approximately 5 mg liver and muscle tissue collected from mice was homogenized in dH2O on ice. Homogenates were boiled then centrifuged at 13,000 rpm at 4°C for 5 min to remove insoluble material. The glycogen content of the homogenates was determined using a colorimetric glycogen assay kit (BioVision, Milpitas, CA) according to manufacturer’s instructions. Absorbance was measured at 570 nm using a Spectra MAX 190 spectrophotometer (Molecular Devices, Sunnyvale, CA). Statistical Analysis Results are plotted as mean ± SE from at least three independent experiments. Statistical analysis was performed using SPSS, version 20. Significant differences between treatments were analyzed using Student’s unpaired t-test and ANOVA when appropriate with significance set at P < 0.05. Significant differences between groups for dependent variables were tested using either single-factor (group) analysis of variance (ANOVA) or two-way (group and time) ANOVA. For single-factor and two-way ANOVAs, Tukey's multiple comparisons test was used for post hoc analyses for between-group comparisons.

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Results Myostatin inhibits the proliferation of Hepa-1C1c7 cells Myostatin is well known for its ability to suppress the proliferation of cultured myoblasts (Joulia et al. 2003; Thomas et al. 2000; Yang et al. 2007). To determine whether it alters the proliferation of liver cells, we examined the effect of increasing concentrations of myostatin on Hepa-1C1c7 cell proliferation. Significant growth inhibition (35% decrease relative to vehicle treated; P < 0.001) was achieved with 4 µg/ml myostatin following 48 h incubation compared to a 70% decrease in similarly treated C2C12 myoblasts (Figure 1A). Flow cytometry revealed a 50% reduction in viable liver cells (P < 0.001) in response to myostatin with no significant difference in cell death evident (Figure 1B), suggesting that myostatin may affect liver cell growth through altered cell cycle regulation rather than by apoptotic mechanisms. As previous studies have shown myostatin to inhibit skeletal muscle cell proliferation through arrest at the G1 phase of the cell cycle (Thomas et al. 2000; Watts et al. 2013), we performed fluorescenceactivated cell sorting (FACS) analysis on control and myostatin-treated Hepa-1C1c7 cells. However unlike myoblasts, our analysis showed no significant differences in the distribution of cells in the S, G2-M and G1 phases (Table 2) with myostatin treatment. Myostatin has also been shown to reduce myoblast proliferation through the regulation of cell cycle gene expression (Joulia et al. 2003; Thomas et al. 2000; Yang et al. 2007). Given this, we speculated that myostatin may affect one or more of the genes involved in the control of liver cell proliferation. Assessment of the transcript levels of several cell cycle regulators found myostatin to induce

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significant (all P < 0.001) reductions in Cdk2, Cdk6, cyclin A2 and cyclin D1 mRNA of 31%, 46%, 33% and 59%, respectively (Figure 1C). Myostatin down-regulates Malat1 expression in vivo and in vitro We performed quantitative RT-PCR analysis in liver tissue from mice injected with PBS (control) or myostatin as previously described (Hittel et al. 2010). We found a significant decrease in Malat1 (24%; P < 0.05) in the liver of myostatin-treated mice compared to controls (Figure 2A). Malat1 gene expression levels were similarly supressed (48%, P < 0.001) in proliferating Hepa-1C1c7 cells following 48 h treatment with 4 µg/ml rh myostatin (Figure 2A). This is in agreement with previously described effects of myostatin on Malat1 expression in human and murine skeletal muscle cells (Watts et al. 2013). siRNA-mediated knockdown of Malat1 suppresses liver cell proliferation Previously, we determined that partial Malat1 silencing suppresses myoblast proliferation (Watts et al. 2013). As such, we examined the effect of partial Malat1 knockdown (Figure 2B; 34% reduction compared to control; P < 0.001) on Hepa-1C1c7 cell proliferation. A 34% knockdown of Malat1 was sufficient to significantly impair liver cell proliferation compared to cells transfected with an appropriate control siRNA following 24 h and 48 h incubations (Figure 2C; 15% and 45% reductions, respectively). As would be expected, significant time effects were observed for the control group only, with a 52% increase (P < 0.001) in Hepa-1C1c7 cell seen following 48 h incubation. Myostatin inhibits insulin-stimulated glucose uptake and Akt phosphorylation in liver cells

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To validate previous observations that myotatin inhibits insulin-stimulated Akt phosphorylation and glucose uptake in the liver (Hittel et al. 2010) we first performed a glucose uptake assay to analyse the effects of rh myostatin on insulin-stimulated glucose uptake by Hepa1C1c7 cells. As expected, untreated cells exhibited a 68% (P < 0.05) increase in glucose uptake in the presence of 100 nM insulin (Figure 3A). Treatment with myostatin had no effect on basal glucose uptake, yet significantly blunted insulin-stimulated glucose uptake (17%, NS) by Hepa1C1c7s, suggesting reduced insulin sensitivity. Next, we used Western blotting to reveal an 8fold decrease (P < 0.05) in insulin-stimulated Akt (Thr308) phosphorylation in myostatin-treated Hepa-1C1c7 cells (Figure 3B). Finally, using a SILAC-based phosphoproteome experiment (Table 4) we identified MARCKS as a novel phosphorylation target of myostatin (Figure 3B) in liver cells. Furthermore, MARCKS appears to be phosphorylated by both myostatin (3.4-fold) and insulin (3.5-fold) in a convergent manner (5.4-fold, all P < 0.05) (Figure 3C). Finally, since myostatin signaling is conveyed in large part through the canonical Smad signaling pathway, we used Western blot analysis to confirm both the presence of the activin type IIb receptor (ActRIIB) as well as increased Smad3 phosphorylation (Figure 3B) in myostatin-treated liver cells. Myostatin reduces liver mass and glycogen content The effect of myostatin on hepatic glycogen content was examined. As shown in Table 3 5-day administration of 15µg/kg/day myostatin decreased liver glycogen content by 52% (P < 0.05). Intriguingly, chronic myostatin administration also reversed the normal weight gain in mice over the course of a week (Table 3) and resulted in significantly lower gastrocnemius muscle (6.6 %, P < 0.05) and liver (2.8 %, P < 0.05) mass.

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Discussion Although elevated plasma levels of myostatin have been associated with a number of obesity-related conditions such as insulin resistance and fatty liver disease, the underlying mechanisms have not been well characterized (Allen et al. 2011; Pedersen and Febbraio 2012). Therefore, the aims of this study were to determine if myostatin regulates liver cell growth, and to establish whether myostatin impairs glucose homeostasis in the liver. Skeletal muscle and liver cells have very different regenerative mechanisms and capacities. For example, myoblasts must withdraw from the cell cycle in order to begin their complex stage of differentiation into functioning skeletal muscle, whereas liver cells are not required to undergo this process. Myostatin has been shown previously to increase the expression of CK1 and p21which in turn, induces G1 phase arrest and inhibits myoblast proliferation (Joulia et al. 2003; Thomas et al. 2000). Despite its heightened expression and circulation in liver disease, it is not known if myostatin works via a similar mechanism in liver cells (Dasarathy 2012; Dasarathy et al. 2011; Garcia et al. 2010; Lang et al. 2004). We hypothesized that myostatin would suppress liver cell proliferation in a manner similar to that previously demonstrated in myoblasts (Thomas et al. 2000). We revealed reduced liver cell proliferation in response to treatment with myostatin. Cell cycle analysis via FACS did not show a significant reduction in the frequency of liver cells in the S phase, indicative of cell cycle arrest at G1 in response to myostatin. However, the accompanying down-regulation in the transcription of cell cycle regulators, cyclin D1, cyclin A2, Cdk2 and Cdk6 not only supports an inhibitory role for myostatin in the transition from G1 to S, particularly through suppressed Cdk2 expression, but also perturbed progression in early G1 and S phases of the cell cycle (Chauhan et al. 2011). Although increased expression of p21 in myoblasts has been associated with reductions in Cdk2

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in the absence of alterations in Cdk4, cyclin D1 and cyclin E (Thomas et al. 2000), we did not observe this in myostatin-treated liver cells. This may indicate that myostatin inhibits liver cell proliferation by a different mechanism than myoblasts or alternatively, that liver cell p21 expression is more transient than in myoblast. Notwithstanding these observations, we did find that myostatin stimulates Smad3 phosphorylation just like it does in muscle cell precursors (Akita et al. 2013). Furthermore, while proteins of the TGF-β family have been previously shown to induce apoptosis in the liver (Chen et al. 2000) we found no evidence of apoptotic cell death in Hepa-1C1c7 cells in response to myostatin. This is in agreement with previous studies showing that myostatin provides protection against apoptosis in myoblasts (Joulia et al. 2003; Rios et al. 2001). Malat1 is a highly conserved 8.7 kb noncoding transcript that is abundantly expressed in cancer cells and as such, is considered a strong predictor of metastasis (Schmidt et al. 2011). Although it is believed to play a direct role in the processing of pre-mRNA (Lin et al. 2011), growing evidence supports a role for Malat1 in the regulation of cell growth. This role is supported by studies showing suppressed tumor growth in response to partial Malat1 knockdown (Han et al. 2013; Lin et al. 2011), and by our own studies describing its expression during and control over key phases of myogenesis (Watts et al. 2013). Given that myostatin suppresses both liver and muscle cell proliferation, we sought to determine if Malat1 is regulated by myostatin in liver cells and whether it also affects their proliferation. Malat1 was found to be significantly down-regulated in response to treatment with myostatin in both mouse livers and liver cells. We also found siRNA-mediated knockdown of Malat1 to suppress liver cell proliferation. Although unable to completely knockdown Malat1 in Hepa-1C1c7 lines, our findings are consistent with previous efforts to silence Malat1 in cancer cell lines (Schmidt et al. 2011) and is likely due to its

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high expression, evidenced by the high average CT values of Malat1 (data not shown). These results are in agreement with our previous findings in both murine and human myoblasts (Watts et al. 2013) and supports a novel role for this long noncoding RNA in hepatogenesis. Indeed, reduced liver and muscle mass in myostatin-treated animals suggests that Malat1 arrests cell proliferation through a mechanism similar to that seen in myoblasts (Watts et al. 2013). The alarming persistence of insulin resistance in the general population requires a better understanding of secreted proteins from peripheral tissues and their role in energy homeostasis (O'Rahilly 2009; Pedersen and Hojman 2012). Although the effects of exogenous myostatin on glucose metabolism have been explored in skeletal muscle (Chen et al. 2010), adipose tissue (Feldman et al. 2006) and whole animal studies, its effect on the liver is understudied given its central role in whole-body energy homeostasis (Hirota and Fukamizu 2010; Hittel et al. 2010; Wang et al. 2012). Recently, it has been shown that myostatin expression with obesity is a potent inducer of insulin resistance by degrading IRS1 protein via the E3 ubiquitin ligase, Cblb (Bonala et al. 2014). These findings complement our observations using SILAC that myostatin induces the phosphorylation of the ubiquitin E1 activating enzyme Uba1 (Table 4) which has also been implicated in cell cycle arrest (Lee et al. 2008). In this study, we demonstrate that myostatin inhibits insulin-stimulated glucose uptake and Akt phosphorylation in cultured liver cells. This in vitro observation is in agreement with previous in vivo studies where we observed decreased insulin-stimulated Akt phosphorylation in the liver and whole body insulin resistance in mice following chronic myostatin administration (Hittel et al. 2010). In this study we have identified a potentially new player in the myostatin-insulin signaling axis, MARCKS, a protein that has been previously associated with both cell proliferation and glucose transport. We have shown that MARCKS is phosphorylated by both myostatin and insulin stimulation suggesting similar

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upstream regulation. Both glucose and insulin have been shown previously to stimulate PKC to phosphorylate MARCKS, which results in the redistribution of this protein to the cytosol (Calle et al. 1992; Chappell et al. 2009). In its apo form, MARCKS binds to and crosslinks actin filaments which effectively sequesters acidic membrane phospholipids such as PIP2. As such MARCKS represents an essential link between insulin stimulation and glucose uptake by mediating glucose transporter fusion. It is somewhat puzzling therefore, how myostatin can both inhibit insulin-stimulated glucose uptake and stimulate the phosphorylation of MARCKS when the opposite would be predicted. One potential interpretation of our results is that MARCKS may be sequestered away from the insulin signaling pathway by myostatin stimulation (Chen et al. 2013).

Furthermore,

MARCKS

phosphorylation

has

been

shown

to

inhibit

PI3K

(phosphatidylinositol 3'-kinase)/Akt phosphorylation suggesting that these pathways are functionally linked. Future studies examining the pharmacological activation or inhibition of MARCKS in the presence or absence of myostatin are required to experimentally establish the possible functional correlations with insulin stimulated glucose uptake. This is particularly relevant for the potential participation of PKC and of the counterintuitive results obtained regarding MARCKS phosphorylation. There have been conflicting studies about the effect of myostatin treatment on glucose uptake in vivo, in vitro (Mitchell et al. 2006) and ex vivo (Antony et al. 2007). The work of Chen et al. reported the first experimental evidence that myostatin increases both basal glucose disposal and glycogenolysis by an AMPK dependent mechanism in C2C12 muscle cells (Chen et al. 2010). Contradicting these findings however, Zhang et al. (Zhang et al. 2011) found that myostatin inactivation leads to increased AMPK activation resulting in increased glucose uptake and improved insulin sensitivity of skeletal muscles in high-fat-fed, myostatin-deficient mice.

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Although we were unable to find any changes in liver AMPK phosphorylation (data not shown), our studies reveal an additional role for myostatin in the suppression of insulin-stimulated glucose disposal and reduced glycogen content of the liver. In summary, we describe a novel role for myostatin in the regulation of liver cell growth and glucose homeostasis. Although myostatin is a well-known regulator of skeletal muscle myogenesis, there is mounting evidence that it plays an important role in regulating lipoprotein metabolism (Guo et al. 2013) and glucose homeostasis in a number of non-muscle tissues (Allen et al. 2011). The mechanism by which myostatin leads to reduced proliferation of liver cells likely involves the down-regulation of Cdk2, Cdk6, cyclin A2, cyclin D1 as well as the long noncoding RNA, Malat1. From this data it may be postulated that myostatin down-regulates liver cell proliferation, at least in part, through suppression of Malat1. Additionally, mouse liver cells cells treated with myostatin exhibited significantly reduced insulin-stimulated glucose uptake and Akt phosphorylation, confirming the previously described insulin-desensitizing effects of myostatin on liver. Finally, we identify Myristoylated Alanine-Rich C-kinase substrate (MARCKS) (Chappell et al. 2009) as a novel downstream target of both myostatin and insulin signaling in the liver with a potential role in the myostatin-induced suppression of insulinstimulated Akt phosphorylation and glucose uptake. Given the protective role of myostatin inhibition in preventing fatty liver disease in mice, the identification of new targets of myostatin will have important repercussions for diseases characterized by inadequate regenerative growth such as chronic hepatitis, cirrhosis and liver cancer (Leevy 1998). These data, therefore support the development of anti-myostatin strategies for the possible treatment of the diminished regenerative capacity of the liver and diseases characterized by liver insulin resistance.

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Acknowledgements R.W. was supported in part by a donation from Encana Corporation of Canada, to the REACH! Campaign. D.S.H. was supported by a Discovery Grant from the Natural Sciences and Engineering Research Council of Canada (NSERC). J.S. has received support from the Alberta Heritage Foundation for Medical Research, the Canadian Institutes for Health Research, the Heart and Stroke Foundation, and the Canadian Diabetes Association. This work was partially supported by core grants NICHD/NINDS 2R24HD050846-06 (National Center for Medical Rehabilitation Research), NICHD 5P30HD040677-10 (Intellectual and Developmental Disabilities Research Center) and NIH NCATS UL1RR031988 (CTSI-CN). The authors would like to thank Dr. Kristy J. Brown of Children’s National Medical Center for her kind assistance in processing the proteomic mass spectrometry samples.

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References Akita, Y., Sumino, Y., Mori, K., Nomura, T., Sato, F., and Mimata, H. 2013. Myostatin inhibits proliferation of human urethral rhabdosphincter satellite cells. International journal of urology : official journal of the Japanese Urological Association 20(5): 522-529. doi: 10.1111/j.1442-2042.2012.03186.x. Allen, D.L., Hittel, D.S., and McPherron, A.C. 2011. Expression and function of myostatin in obesity, diabetes, and exercise adaptation. Medicine and science in sports and exercise 43(10): 1828-1835. doi: 10.1249/MSS.0b013e3182178bb4. Antony, N., Bass, J.J., McMahon, C.D., and Mitchell, M.D. 2007. Myostatin regulates glucose uptake in BeWo cells. Am J Physiol Endocrinol Metab 293(5): E1296-1302. doi: 10.1152/ajpendo.00331.2007. Bonala, S., Lokireddy, S., McFarlane, C., Patnam, S., Sharma, M., and Kambadur, R. 2014. Myostatin induces insulin resistance via Cblb-mediated degradation of IRS1 in response to high calorie diet intake. The Journal of biological chemistry. doi: 10.1074/jbc.M113.529925. Bonomi, L., Brown, M., Ungerleider, N., Muse, M., Matzuk, M.M., and Schneyer, A. 2012. Activin B regulates islet composition and islet mass but not whole body glucose homeostasis or insulin sensitivity. American journal of physiology. Endocrinology and metabolism 303(5): E587-596. doi: 10.1152/ajpendo.00177.2012. Calle, R., Ganesan, S., Smallwood, J.I., and Rasmussen, H. 1992. Glucose-induced phosphorylation of myristoylated alanine-rich C kinase substrate (MARCKS) in isolated rat pancreatic islets. The Journal of biological chemistry 267(26): 18723-18727. Chappell, D.S., Patel, N.A., Jiang, K., Li, P., Watson, J.E., Byers, D.M., and Cooper, D.R. 2009. Functional involvement of protein kinase C-betaII and its substrate, myristoylated alanine-rich C-kinase substrate (MARCKS), in insulin-stimulated glucose transport in L6 rat skeletal muscle cells. Diabetologia 52(5): 901911. doi: 10.1007/s00125-009-1298-7. Chauhan, A., Lorenzen, S., Herzel, H., and Bernard, S. 2011. Regulation of mammalian cell cycle progression in the regenerating liver. Journal of theoretical biology 283(1): 103-112. doi: 10.1016/j.jtbi.2011.05.026. Chen, C.H., Thai, P., Yoneda, K., Adler, K.B., Yang, P.C., and Wu, R. 2013. A peptide that inhibits function of Myristoylated Alanine-Rich C Kinase Substrate (MARCKS) reduces lung cancer metastasis. Oncogene. doi: 10.1038/onc.2013.336. Chen, W., Woodruff, T.K., and Mayo, K.E. 2000. Activin A-induced HepG2 liver cell apoptosis: involvement of activin receptors and smad proteins. Endocrinology 141(3): 1263-1272. Chen, Y., Ye, J., Cao, L., Zhang, Y., Xia, W., and Zhu, D. 2010. Myostatin regulates glucose metabolism via the AMP-activated protein kinase pathway in skeletal muscle cells. The international journal of biochemistry & cell biology 42(12): 2072-2081. doi: 10.1016/j.biocel.2010.09.017. Dasarathy, S. 2012. Consilience in sarcopenia of cirrhosis. Journal of cachexia, sarcopenia and muscle 3(4): 225-237. doi: 10.1007/s13539-012-0069-3. Dasarathy, S., McCullough, A.J., Muc, S., Schneyer, A., Bennett, C.D., Dodig, M., and Kalhan, S.C. 2011. Sarcopenia associated with portosystemic shunting is reversed by follistatin. Journal of hepatology 54(5): 915-921. doi: 10.1016/j.jhep.2010.08.032.

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21 Feldman, B.J., Streeper, R.S., Farese, R.V., Jr., and Yamamoto, K.R. 2006. Myostatin modulates adipogenesis to generate adipocytes with favorable metabolic effects. Proceedings of the National Academy of Sciences of the United States of America 103(42): 15675-15680. doi: 10.1073/pnas.0607501103. Garcia, P.S., Cabbabe, A., Kambadur, R., Nicholas, G., and Csete, M. 2010. Brief-reports: elevated myostatin levels in patients with liver disease: a potential contributor to skeletal muscle wasting. Anesthesia and analgesia 111(3): 707-709. doi: 10.1213/ANE.0b013e3181eac1c9. Guo, W., Wong, S., and Bhasin, S. 2013. AAV-Mediated Administration of Myostatin Pro-Peptide Mutant in Adult Ldlr Null Mice Reduces Diet-Induced Hepatosteatosis and Arteriosclerosis. PloS one 8(8): e71017. doi: 10.1371/journal.pone.0071017. Han, Y., Liu, Y., Nie, L., Gui, Y., and Cai, Z. 2013. Inducing cell proliferation inhibition, apoptosis, and motility reduction by silencing long noncoding ribonucleic acid metastasis-associated lung adenocarcinoma transcript 1 in urothelial carcinoma of the bladder. Urology 81(1): 209 e201-207. doi: 10.1016/j.urology.2012.08.044. Hirota, K., and Fukamizu, A. 2010. Transcriptional regulation of energy metabolism in the liver. Journal of receptor and signal transduction research 30(6): 403-409. doi: 10.3109/10799893.2010.509730. Hittel, D.S., Axelson, M., Sarna, N., Shearer, J., Huffman, K.M., and Kraus, W.E. 2010. Myostatin decreases with aerobic exercise and associates with insulin resistance. Medicine and science in sports and exercise 42(11): 2023-2029. doi: 10.1249/MSS.0b013e3181e0b9a8. Hittel, D.S., Berggren, J.R., Shearer, J., Boyle, K., and Houmard, J.A. 2009. Increased secretion and expression of myostatin in skeletal muscle from extremely obese women. Diabetes 58(1): 30-38. doi: 10.2337/db08-0943. Jiao, J., Yuan, T., Zhou, Y., Xie, W., Zhao, Y., Zhao, J., Ouyang, H., and Pang, D. 2011. Analysis of myostatin and its related factors in various porcine tissues. Journal of animal science 89(10): 3099-3106. doi: 10.2527/jas.2010-3827. Joulia, D., Bernardi, H., Garandel, V., Rabenoelina, F., Vernus, B., and Cabello, G. 2003. Mechanisms involved in the inhibition of myoblast proliferation and differentiation by myostatin. Experimental cell research 286(2): 263-275. Lang, C.H., Frost, R.A., Svanberg, E., and Vary, T.C. 2004. IGF-I/IGFBP-3 ameliorates alterations in protein synthesis, eIF4E availability, and myostatin in alcohol-fed rats. American journal of physiology. Endocrinology and metabolism 286(6): E916-926. doi: 10.1152/ajpendo.00554.2003. Lee, T.V., Ding, T., Chen, Z., Rajendran, V., Scherr, H., Lackey, M., Bolduc, C., and Bergmann, A. 2008. The E1 ubiquitin-activating enzyme Uba1 in Drosophila controls apoptosis autonomously and tissue growth non-autonomously. Development 135(1): 43-52. doi: 10.1242/dev.011288. Leevy, C.B. 1998. Abnormalities of liver regeneration: a review. Digestive diseases 16(2): 88-98. Lin, R., Roychowdhury-Saha, M., Black, C., Watt, A.T., Marcusson, E.G., Freier, S.M., and Edgington, T.S. 2011. Control of RNA processing by a large non-coding RNA over-expressed in carcinomas. FEBS letters 585(4): 671-676. doi: 10.1016/j.febslet.2011.01.030. Luo, J.H., Ren, B., Keryanov, S., Tseng, G.C., Rao, U.N., Monga, S.P., Strom, S., Demetris, A.J., Nalesnik, M., Yu, Y.P., Ranganathan, S., and Michalopoulos, G.K. 2006. Transcriptomic and genomic analysis of human hepatocellular carcinomas and hepatoblastomas. Hepatology 44(4): 1012-1024. doi: 10.1002/hep.21328. McCroskery, S., Thomas, M., Maxwell, L., Sharma, M., and Kambadur, R. 2003. Myostatin negatively regulates satellite cell activation and self-renewal. The Journal of cell biology 162(6): 1135-1147. doi: 10.1083/jcb.200207056. McCroskery, S., Thomas, M., Platt, L., Hennebry, A., Nishimura, T., McLeay, L., Sharma, M., and Kambadur, R. 2005. Improved muscle healing through enhanced regeneration and reduced fibrosis in myostatin-null mice. Journal of cell science 118(Pt 15): 3531-3541. doi: 10.1242/jcs.02482.

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22 McFarland, D.C., Velleman, S.G., Pesall, J.E., and Liu, C. 2006. Effect of myostatin on turkey myogenic satellite cells and embryonic myoblasts. Comparative biochemistry and physiology. Part A, Molecular & integrative physiology 144(4): 501-508. doi: 10.1016/j.cbpa.2006.04.020. McPherron, A.C., and Lee, S.J. 2002. Suppression of body fat accumulation in myostatin-deficient mice. The Journal of clinical investigation 109(5): 595-601. doi: 10.1172/JCI13562. Mirza, Z., Kamal, M.A., Abuzenadah, A.M., Al-Qahtani, M.H., and Karim, S. 2013. Establishing Genomic/Transcriptomic Links between Alzheimer's Disease and Type II Diabetes Mellitus by MetaAnalysis Approach. CNS & neurological disorders drug targets. Mitchell, M.D., Osepchook, C.C., Leung, K.C., McMahon, C.D., and Bass, J.J. 2006. Myostatin is a human placental product that regulates glucose uptake. J Clin Endocrinol Metab 91(4): 1434-1437. doi: 10.1210/jc.2005-2361. O'Rahilly, S. 2009. Human genetics illuminates the paths to metabolic disease. Nature 462(7271): 307314. doi: 10.1038/nature08532. Pedersen, B.K., and Febbraio, M.A. 2012. Muscles, exercise and obesity: skeletal muscle as a secretory organ. Nature reviews. Endocrinology 8(8): 457-465. doi: 10.1038/nrendo.2012.49. Pedersen, L., and Hojman, P. 2012. Muscle-to-organ cross talk mediated by myokines. Adipocyte 1(3): 164-167. doi: 10.4161/adip.20344. Rios, R., Carneiro, I., Arce, V.M., and Devesa, J. 2001. Myostatin regulates cell survival during C2C12 myogenesis. Biochemical and biophysical research communications 280(2): 561-566. doi: 10.1006/bbrc.2000.4159. Rios, R., Fernandez-Nocelos, S., Carneiro, I., Arce, V.M., and Devesa, J. 2004. Differential response to exogenous and endogenous myostatin in myoblasts suggests that myostatin acts as an autocrine factor in vivo. Endocrinology 145(6): 2795-2803. doi: 10.1210/en.2003-1166. Schmidt, L.H., Spieker, T., Koschmieder, S., Schaffers, S., Humberg, J., Jungen, D., Bulk, E., Hascher, A., Wittmer, D., Marra, A., Hillejan, L., Wiebe, K., Berdel, W.E., Wiewrodt, R., and Muller-Tidow, C. 2011. The long noncoding MALAT-1 RNA indicates a poor prognosis in non-small cell lung cancer and induces migration and tumor growth. Journal of thoracic oncology : official publication of the International Association for the Study of Lung Cancer 6(12): 1984-1992. doi: 10.1097/JTO.0b013e3182307eac. Siriett, V., Salerno, M.S., Berry, C., Nicholas, G., Bower, R., Kambadur, R., and Sharma, M. 2007. Antagonism of myostatin enhances muscle regeneration during sarcopenia. Molecular therapy : the journal of the American Society of Gene Therapy 15(8): 1463-1470. doi: 10.1038/sj.mt.6300182. Taylor, W.E., Bhasin, S., Artaza, J., Byhower, F., Azam, M., Willard, D.H., Jr., Kull, F.C., Jr., and GonzalezCadavid, N. 2001. Myostatin inhibits cell proliferation and protein synthesis in C2C12 muscle cells. American journal of physiology. Endocrinology and metabolism 280(2): E221-228. Thomas, M., Langley, B., Berry, C., Sharma, M., Kirk, S., Bass, J., and Kambadur, R. 2000. Myostatin, a negative regulator of muscle growth, functions by inhibiting myoblast proliferation. The Journal of biological chemistry 275(51): 40235-40243. doi: 10.1074/jbc.M004356200. Wang, F., Liao, Y., Li, X., Ren, C., Cheng, C., and Ren, Y. 2012. Increased circulating myostatin in patients with type 2 diabetes mellitus. Journal of Huazhong University of Science and Technology. Medical sciences = Hua zhong ke ji da xue xue bao. Yi xue Ying De wen ban = Huazhong keji daxue xuebao. Yixue Yingdewen ban 32(4): 534-539. doi: 10.1007/s11596-012-0092-9. Watts, R., Johnsen, V.L., Shearer, J., and Hittel, D.S. 2013. Myostatin-induced inhibition of the long noncoding RNA Malat1 is associated with decreased myogenesis. American journal of physiology. Cell physiology 304(10): C995-1001. doi: 10.1152/ajpcell.00392.2012. Wilkes, J.J., Lloyd, D.J., and Gekakis, N. 2009. Loss-of-function mutation in myostatin reduces tumor necrosis factor alpha production and protects liver against obesity-induced insulin resistance. Diabetes 58(5): 1133-1143. doi: 10.2337/db08-0245.

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23 Yang, W., Zhang, Y., Li, Y., Wu, Z., and Zhu, D. 2007. Myostatin induces cyclin D1 degradation to cause cell cycle arrest through a phosphatidylinositol 3-kinase/AKT/GSK-3 beta pathway and is antagonized by insulin-like growth factor 1. The Journal of biological chemistry 282(6): 3799-3808. doi: 10.1074/jbc.M610185200. Zhang, C., McFarlane, C., Lokireddy, S., Bonala, S., Ge, X., Masuda, S., Gluckman, P.D., Sharma, M., and Kambadur, R. 2011. Myostatin-deficient mice exhibit reduced insulin resistance through activating the AMP-activated protein kinase signalling pathway. Diabetologia 54(6): 1491-1501. doi: 10.1007/s00125011-2079-7. Zhang, C., McFarlane, C., Lokireddy, S., Masuda, S., Ge, X., Gluckman, P.D., Sharma, M., and Kambadur, R. 2012. Inhibition of myostatin protects against diet-induced obesity by enhancing fatty acid oxidation and promoting a brown adipose phenotype in mice. Diabetologia 55(1): 183-193. doi: 10.1007/s00125011-2304-4. Zimmers, T.A., Davies, M.V., Koniaris, L.G., Haynes, P., Esquela, A.F., Tomkinson, K.N., McPherron, A.C., Wolfman, N.M., and Lee, S.J. 2002. Induction of cachexia in mice by systemically administered myostatin. Science 296(5572): 1486-1488. doi: 10.1126/science.1069525.

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Table 1. Primers used for RT-PCR Gene

Accession No.

Primer sequences (5’ → 3’)

β-actin

NM_001101.3

F: TGTTACCAACTGGGACGACA R: GGGGTGTTGAAGGTCTCAAA

Cdk2

NM_183417.2

F: CATTCCTCTTCCCCTCATCA R: GCAGCCCAGAAGAATTTCAG

Cdk4

NM_009870.2

F: CGGCCTGTGTCTATGGTCTG R: TGGTCGGCTTCAGAGTTTCC

Cdk6

NM_009873.2

F: CGCCGATCAGCAGTATGAGT R: GCCGGGCTCTGGAACTTTAT

cyclin A2

NM_009828.2

F: GAACTACAAGACCAGCAGCCG R: ATGGTGAAGGCAGGCTGTTT

cyclin D1

NM_007631

F: GTACCCTGACACCAATCT R: ATCTCCTTCTGCACGCACTT

cyclin E1

NM_007633.2

F: ATTGGCTAATGGAGGTGTGC R: CCACTTAAGGGCCTTCATCA

Malat1

NC_000085.5

F: GAGTTCTAATTCTTTTTACTGCTCAATC R: AGAGCAGAGCAGCGTAGAGC

p21

NM_007669

F: CACAGCTCAGTGGACTGGAA R: ACCCTAGACCCACAATGCAG

F- forward; R- reverse.

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Table 2. Cell cycle distribution of Hepa-1C1c7 cells after myostatin treatment. Percentage of cells from 5000 counts in G1, S or G2-M phases of the cell cycle according to FACS analysis. Data are mean ± SE. Two independent experiments were performed and quantified in triplicate. No statistically significant differences (P < 0.05) between control and myostatin-treated Hepa1C1c7 cells were identified using Student’s t-test.

Treatment

G1 (%)

S (%)

G2-M (%)

control

35.1 ± 2.4

42.5 ± 0.9

22.4 ± 2.9

myostatin

39.4 ± 2.7

41.1 ± 2.1

19.5 ± 2.7

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Table 3. Characteristics of PBS- (CTRL) and MSTN-treated (MSTN) mice (Mean +/- SEM). Statistically significant differences (*P < 0.05) between control and myostatin-treated groups were analyzed by Student’s t-test.

Weight

CTRL (n=10)

MSTN (n=10)

Day 1

22.02±0.39

22.72±0.22

Day 3

22.43±0.45

22.32±0.22

Day 5

22.68±0.68

22.32±0.55

Change

0.66±0.04

-0.40±0.04

Gastrocnemius (mg)

146.65±2.25

136.93±3.19 *

Gastrocnemius Glycogen (mg/g wt)

1.8±0.20

1.5±0.30

Liver Mass (mg)

846.23±4.41

822.21±2.22 *

Liver Glycogen (mg/g wt)

22.3±1.48

10.6±0.99 *

Glucose (mM)

8.43±0.34

8.03±0.20

Mouse Weight (g)

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Table 4. Curated list of differentially phosphorylated proteins isolated from myostatin treated Hepa1C1c7 cells using SILAC. Shown are the protein accession number, average ratio of phosphorylated to total protein, number of peptides identified and a brief description of the protein function.

Accession

Ratio ± SD

Peptides

Description

P26645 P35564

2.57 ± 0.58 2.46 ± 0.49

12 5

Myristoylated alanine-rich C-kinase substrate Calnexin

Q8BP47

1.97 ± 0.46

4

Asparaginyl-tRNA synthetase

Q9Z1Q9

1.86 ± 0.46

4

Valyl-tRNA synthetase

P62984

1.75 ± 0.55

13

Ubiquitin-60S ribosomal protein L40

Q02053 Q3UMU9

1.55 ± 0.52 1.57 ± 0.44

3 5

Ubiquitin-like modifier-activating enzyme 1 Hepatoma-derived growth factor-related protein 2

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Figure 1. Myostatin inhibits liver cell proliferation via the down-regulation of genes that control the G1-S phase transition. (A) Proliferation analysis of C2C12 and Hepa-1C1c7 cells grown for 48 h in the presence of increasing concentrations of rh myostatin (MSTN) (0-4 µg/ml) and monitored by colormetric assay. (B) Cell viability analysis was performed via FACS in liver cells in response to 4 µg/ml myostatin treatment. (C) The relative gene expression levels of p21, Cdk2, Cdk4, Cdk6, cyclin A2, cyclin D1 and cyclin E1 (relative to β-actin) were determined by RT-PCR in proliferating Hepa-1C1c7 cells following 48 h treatment with or without 4 µg/ml rh myostatin. Columns represent the mean ± SE. Two independent experiments were performed and quantified in triplicate. Statistically significant differences (‡P < 0.001) between control and myostatin-treated cells were analyzed by Student’s t-test. Figure 2. Myostatin decreases liver Malat1 expression in vivo and in vitro. (A) The effect of 6 days of 15 µg/kg rh myostatin treatment on mouse liver (Mstn-Liver) and 48 h treatment with 4 µg/ml rh myostatin on mouse liver cells (Mstn-Cells). RT-PCR was performed for indicated genes and expressed relative to levels of β-actin. (B) Next, Hepa-1C1c7 cells were reverse transfected with 0.5 µM universal control (black bars) or Malat1 siRNA (open bars). Following 72 h incubation, Malat1 gene transcript levels (relative to β-actin) were determined by RT-PCR. (C) Finally, proliferation analysis of partial Malat1-knockdown cells was conducted after 24 and 48 h of incubation. Columns represent the mean ± SE. Animal experiments were performed in ten male mice per treatment. Two independent experiments were performed and quantified in triplicate for all cell experiments. Statistically significant differences (*P < 0.05; ‡P < 0.001) between control and Malat1 siRNA-treated Hepa-1C1c7 cells were analyzed by Student’s t-test,

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29

while significant treatment (‡P < 0.001) and time (cP < 0.001) effects were analyzed in Malat1knockdown cells by two-way ANOVA.

Figure 3. Myostatin decreases insulin-stimulated glucose uptake and AKT phosphorylation in liver cells. (A) Basal and insulin stimulated (100 mM) 2-[14C]deoxy-D-glucose (2-DG) uptake was determined after a 1h pretreatment with 4ug/ml myostatin (MSTN). (B) Representative blot and (C) Western blot analysis of AKT, MARCKS and Smad3 phosphorylation status in serum starved cells (S), in the presence of 4ug/ml myostatin (M) or 100 nM insulin (I) and in the presence of both myostatin (4ug/ml) and insulin (100 nM) (M+I). Phosphorylation levels were normalized relative to the intensity of the apo/total forms of their respective protein and the control intensity set to a value of 1. Columns represent the mean ± SE. Two independent experiments were performed and quantified in triplicate. Statistically significant myostatin (cP < 0.001) and insulin-stimulated effects (‡P < 0.001) were analyzed by two-way ANOVA.

Biochem. Cell Biol. Downloaded from www.nrcresearchpress.com by UNIV CALGARY on 04/25/14 ly. This Just-IN manuscript is the accepted manuscript prior to copy editing and page composition. It may differ from the final official Growth inhibition (%) 100

60

‡

40

0 1 2 Myostatin Concentration (ug/ml)

C

‡

20

0

-20 4

C2C12

Hepa-1C1c7

Cell Number (1000s)

A

B 2500

‡

80

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2000

1500

‡

control viable cells

1000 dead cells

500

0 MSTN

Con-Liver

Relative Absorbance

Malat1 mRNA relative to -actin

Biochem. Cell Biol. Downloaded from www.nrcresearchpress.com by UNIV CALGARY on 04/25/14 ly. This Just-IN manuscript is the accepted manuscript prior to copy editing and page composition. It may differ from the final official

1.2

1.0

0.8

‡ ‡

0.6

0.4

0.2

Con-Cells Mstn-Liver

1.8

1.0

24 h

Malat1 mRNA relative to -actin

Page 31 of 32 A

B 1.2

0.0

1.0

0.8

‡

0.8

0.6

0.4

0.2

0.0 48 h

‡

0.6

0.4

0.2

Mstn-Cells 0.0 control

‡ Malat1 siRNA

C 1.6

c

1.4

1.2

control control

0.5 uM siRNA Malat1 siRNA

Relative Expression 2-[3H]deoxy-D-glucose uptake/cell protein content

Biochem. Cell Biol. Downloaded from www.nrcresearchpress.com by UNIV CALGARY on 04/25/14 ly. This Just-IN manuscript is the accepted manuscript prior to copy editing and page composition. It may differ from the final official

A

5.0

80.0

control control

70.0

MSTN 0.1 ug/ml myostatin

basal

4.0

Control

‡

MARCKS

B

60.0

40.0

20.0

6.0

SMAD3

‡

Myostatin Insulin

Page 32 of 32

S

c

pAKT (Thr308)

50.0

pMARCKS (Ser152/156)

30.0

pSmad3 (Ser423/425)

10.0

0.0

B-Actin

100 nM insulin

C ActRIIB

‡

AKT

‡ ‡ ‡

‡

3.0

2.0

1.0

0.0

Myostatin + Insulin

M I M+I