The FASEB Journal article fj.13-245894. Published online January 3, 2014.

The FASEB Journal • Research Communication

Elevated expression of activins promotes muscle wasting and cachexia Justin L. Chen,*,†,‡ Kelly L. Walton,† Catherine E. Winbanks,* Kate T. Murphy,§ Rachel E. Thomson,* Yogeshwar Makanji,† Hongwei Qian,* Gordon S. Lynch,§ Craig A. Harrison,†,‡,1 and Paul Gregorevic*,‡,§,储,1,2 *Baker IDI Heart and Diabetes Institute, Melbourne, Victoria, Australia; †Prince Henry’s Institute, Clayton, Victoria, Australia; ‡Department of Biochemistry and Molecular Biology, Monash University, Melbourne, Victoria, Australia; §Department of Physiology, University of Melbourne, Melbourne, Victoria, Australia; and 储Department of Neurology, University of Washington School of Medicine, Seattle, Washington, USA In models of cancer cachexia, inhibiting type IIB activin receptors (ActRIIBs) reverse muscle wasting and prolongs survival, even with continued tumor growth. ActRIIB mediates signaling of numerous TGF-␤ proteins; of these, we demonstrate that activins are the most potent negative regulators of muscle mass. To determine whether activin signaling in the absence of tumor-derived factors induces cachexia, we used recombinant serotype 6 adeno-associated virus (rAAV6) vectors to increase circulating activin A levels in C57BL/6 mice. While mice injected with control vector gained ⬃10% of their starting body mass (3.8ⴞ0.4 g) over 10 wk, mice injected with increasing doses of rAAV6:activin A exhibited weight loss in a dose-dependent manner, to a maximum of ⴚ12.4% (ⴚ4.2ⴞ1.1 g). These reductions in body mass in rAAV6:activin-injected mice correlated inversely with elevated serum activin A levels (7- to 24-fold). Mechanistically, we show that activin A reduces muscle mass and function by stimulating the ActRIIB pathway, leading to deleterious consequences, including increased transcription of atrophy-related ubiquitin ligases, decreased Akt/mTOR-mediated protein synthesis, and a profibrotic response. Critically, we demonstrate that the muscle wasting and fibrosis that ensues in response to excessive activin levels is fully

ABSTRACT

Abbreviations: AAV, adeno-associated virus; ActRIIB, type II activin receptor; ALK, activin receptor-like kinase; ANOVA, analysis of variance; BSA, bovine serum albumin; C26, colon26; CMV, cytomegalovirus; DMEM, Dulbecco’s modified Eagle’s medium; ECM, extracellular matrix; Fn14, fibroblast growth factor-inducible receptor 14; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; HEK, human embryonic kidney; HRP, horseradish peroxidase; IGF, insulin-like growth factor; IL, interleukin; mTOR, mammalian target of rapamycin; nNOS, neuronal nitric oxide synthase; PBS, phosphate buffered saline; qRT-PCR, quantitative reverse transcription polymerase chain reaction; rAAV6, recombinant serotype 6 adeno-associated virus; S6K, S6 kinase; S6RP, S6 ribosomal protein; TA, tibialis anterior; Tet, tetracycline; TGF-␤, transforming growth factor-␤, TNF, tumor necrosis factor; TRE, tetracycline responsive element; TWEAK, TNF-like weak inducer of apoptosis; vg, vector genome 0892-6638/14/0028-0001 © FASEB

reversible. These findings highlight the therapeutic potential of targeting activins in cachexia.—Chen, J. L., Walton, K. L., Winbanks, C. E., Murphy, K. T., Thomson, R. E., Makanji, Y., Qian, H., Lynch, G. S., Harrison, C. A., Gregorevic, P. Elevated expression of activins promotes muscle wasting and cachexia. FASEB J. 28, 000 – 000 (2014). www.fasebj.org Key Words: atrophy 䡠 myostatin 䡠 adeno-associated virus Cancer cachexia is a state of severe frailty and fatigue, associated with pronounced loss of skeletal muscle and fat mass (1). Cachexia constitutes a major unmet medical challenge, as up to 80% of patients with advanced cancers exhibit cachectic symptoms, and as many as 25% of cancer-related mortalities may derive from the complications of cachexia rather than direct tumor burden (2, 3). The etiology of cachexia is attributed to abnormal metabolism and catabolism, ostensibly induced by tumor- and host-derived cytokines and factors (4). Recent evidence suggests that signaling via the activin type II receptor (ActRIIB) may play a dominant role in the development and progression of cachexia (5). Significantly, inhibition of circulating ActRIIB ligands via administration of soluble recombinant ActRIIB can reverse muscle wasting and prolong survival in mouse models of cancer cachexia, even in the face of continued tumor growth (5). The findings suggest interventions that inhibit ActRIIB signaling may have considerable therapeutic potential for cachexia. However, as several transforming growth factor-␤ (TGF-␤) family members utilize the ActRIIB pathway to regulate homeostasis in a variety of organ systems, it is critical to identify the specific ActRIIB 1

These authors contributed equally to this work. Correspondence: Baker IDI Heart and Diabetes Institute, P.O. Box 6492, St. Kilda Rd. Central, Melbourne 8008, Australia. E-mail: [email protected] doi: 10.1096/fj.13-245894 This article includes supplemental data. Please visit http:// www.fasebj.org to obtain this information. 2

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ligands that promote muscle wasting and define their mechanisms of action to accelerate the development of effective and safe interventions for cachexia. As increased serum concentrations of activin A have been observed in malignant conditions (6, 7) and tumorbearing mice exhibiting a cachexia-like syndrome (8), we tested the hypothesis that activins signaling via ActRIIB are direct repressors of skeletal muscle mass and functional capacity. Activin A and activin B are synthesized as precursor molecules with the N-terminal prodomain mediating the folding and dimerization of the C-terminal mature domain. Dimeric precursors are cleaved by proprotein convertases, and activins are secreted from the cell noncovalently associated with their prodomains. Extracellularly, the prodomain localizes activin dimers within the vicinity of target cells via interactions with the extracellular matrix (ECM; refs. 9, 10). Activins are displaced from their prodomains by the type II receptors ActRIIA/IIB, which leads to the recruitment, phosphorylation, and activation of the type I receptor activin receptor-like kinase 4 (ALK4). Activated ALK4 phosphorylates intracellular signaling molecules, Smad2/3, which in turn form a complex with the coactivator Smad4. The resulting Smad oligomer localizes within the nucleus to regulate target genes in a cell- and context-dependent manner (11). The ability of activins to access their signaling receptors is regulated by several extracellular proteins, including inhibins and follistatin (12, 13). Targeted deletion of the inhibin ␣-subunit in mice leads to unopposed expression of activin A and activin B within the gonads and the development of ovarian and testicular sex cord-stromal tumors with 100% penetrance, as early as 4 wk of age (14). As these tumors progress, serum levels of activin A and B increase by as much as 500-fold (15) and result in marked cachexia and death (8). Elevated activin levels in these cachectic mice were originally associated with anemia, hepatocellular necrosis in the liver, and atrophy of the stomach (8). Notably, inhibin-deficient mice also exhibit the profound losses of skeletal muscle, heart, and fat mass that are key indicators of mortality in cancer cachexia (5). Similarly, reductions in muscle mass have also been observed in mice bearing activin A-secreting xenografts (5). While these prior studies support the concept that elevated circulating concentrations of activins are associated with a catabolic phenotype, the effects of activin on skeletal muscle have not been examined independent of tumor development and progression. Thus, it remains unclear whether activins are direct promoters of muscle wasting, and if so, how the actions of these ligands alter processes within skeletal muscle to negatively regulate cell size and contractile function. Here, we used recombinant serotype 6 adeno-assoicated virus (rAAV6) vectors to overexpress activins in the skeletal muscles of mice and to undertake the first assessment of local and systemic effects of activins on the morphological, functional, and biochemical attributes of skeletal musculature, independent of tumor 2

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development and progression. Our data demonstrate that activins are potent promoters of skeletal muscle wasting and fibrosis, but that activin-mediated pathology is reversible, even from a considerably advanced state. Thus, activins represent targets for therapeutic manipulation to ameliorate muscle wasting associated with cancer cachexia and conditions where serum activin levels are elevated.

MATERIALS AND METHODS Antibodies All antibodies used were obtained from Cell Signaling Technologies (Beverly, MA, USA), except for antibodies against myostatin, TGF-␤1, and Pi-16 (R&D Systems, Minneapolis, MN, USA); Smad3 and pSmad3 (Epitomics, Burlingame, CA, USA); activin A (E4) and activin B (C5) (Oxford Bioinnovations, Bicester, UK); fibroblast-specific protein-1/S100A4 (Abcam, Cambridge, UK), and glyceraldehyde 3-phosphate dehydrogenase (GAPDH; Santa Cruz Biotechnology, Dallas, TX, USA). Cell culture Human cancer cell lines were grown to confluence, at which point growth medium consisting of Dulbecco’s modified Eagle’s medium (DMEM) and 10%FCS (Life Technologies, Grand Island, NY, USA) was replaced with Opti-MEM (Life Technologies) for 48 h, after which conditioned medium was collected. Production of recombinant adeno-associated virus (AAV) vectors cDNA constructs encoding for activin A, activin B, myostatin, and TGF-␤1 were cloned into an AAV expression plasmid consisting of a cytomegalovirus (CMV) promoter/enhancer and SV40 poly-A region flanked by AAV2 terminal repeats using standard cloning techniques. For inducible vectors, activin A cDNA was subcloned into an AAV expression plasmid consisting of a tetracycline responsive element (TRE) promoter. Transfection of these plasmids with the pDGM6 packaging plasmid into human embryonic kidney (HEK)-293 cells generated type-6 pseudotyped viral vectors, which were harvested and purified as described previously (16). Briefly, HEK-293 cells were plated at a density of 3.2–3.8 ⫻ 106 cells on a 10-cm culture dish, 8 –16 h before transfection with 10 ␮g of a vector genome-containing plasmid and 20 ␮g of the packaging/helper plasmid pDGM6, by means of the calcium phosphate precipitate method. After 72 h, the medium and cells were collected and homogenized through a microfluidizer (Microfluidics, Westwood, MA, USA) before 0.22-␮m clarification (Millipore, Billerica, MA, USA). The vector was purified from the clarified lysate by affinity chromatography over a HiTrap heparin column (GE Healthcare, Little Chalfont, UK) and ultracentrifuged overnight before resuspension in sterile physiological Ringer’s solution. The purified vector preparations were titered with a customized sequencespecific quantitative PCR-based reaction (Life Technologies). Animal experiments All experiments were conducted in accordance with the relevant codes of practice for the care and use of animals for

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scientific purposes (U.S. National Institutes of Health, 1985, and the National Health and Medical Research Council of Australia, 2013). For vector doses of 109–1011 vector genomes (vg), the right tibialis anterior (TA) muscle of 6- to 8-wk-old C57BL/6 male mice were injected with rAAV6 containing the transgene of interest, and control injections of the contralateral limb used a vector lacking a functional gene. In the inducible studies, the TA muscles were injected with a rAAV6 vector encoding the tetracycline (Tet)-On protein under a CMV promoter (rAAV6:Tet-On) and a rAAV6 vector encoding activin A under a TRE promoter. The contralateral TA muscle was injected with rAAV6:Tet-On and a vector lacking a functional gene. Activin A expression was induced by the addition of doxycycline antibiotic to the chow at 600 mg/kg for 4 wk, after which transduction was switched off by substitution of diet to doxycycline-free chow for 1– 8 wk. For high-dose vector delivery (⬎2⫻1011 vg), multiple muscles were injected within a single hindlimb of 20-wk-old mice. All mice were deeply anesthetized with isoflurane, and vectors were injected in Hank’s buffered saline solution (Life Technologies) directly into the hindlimb muscles. For mice designated for testing muscle function, anesthetized 12-wk-old male C57BL/10 mice (via an intraperitoneal injection of a mixture of 100 mg/kg ketamine and 10 mg/kg xylazine) were injected in the right TA muscle only, either with rAAV6: activin A or vector lacking a functional gene. Body composition analysis to assess fat and lean body mass was performed using quantitative magnetic resonance (EchoMRI, Houston, TX, USA). At the experimental end point, mice were killed humanely via cervical dislocation or CO2 asphyxiation, and the muscles and organs were excised rapidly and weighed before subsequent processing. Quantitative reverse transcription polymerase chain reaction (RT-PCR) Total RNA was collected from TA muscle cells using TRIzol (Life Technologies). RNA (1–3 ␮g) was reverse transcribed using the High Capacity RNA-to-cDNA kit (Life Technologies). Expression levels of all genes were analyzed by qRT-PCR, with 18S to standardize cDNA concentrations, using TaqMan assay ondemand kits (Life Technologies) and ABI detection software (Life Technologies). For detection of activin A gene expression, SYBR Green analysis was used (forward: GGAGTGTGATGGCAAGGTCAACA; reverse: GTGGGCACACAGCATGACTTA), with 18S to standardize cDNA concentrations (forward: GGGAGCCTGAGAAACGGC; reverse: GGGTCGGGAGTGGGTAATTT). Data were analyzed using the ⌬⌬CT method of analysis and normalized to a control value of 1. Histology Harvested muscles were placed in optimal cutting temperature (OCT) cryoprotectant (Sakura Finetek, Torrance, CA, USA) and frozen in liquid nitrogen-cooled isopentane. The frozen samples were subsequently cryosectioned at 10 ␮m thickness and stained with hematoxylin and eosin to examine morphology as described previously (17). Sections were mounted using DePeX mounting medium (VWR, Lutterworth, UK), and images of stained sections were captured at room temperature using a U-TV1X-2 camera mounted to an IX71 microscope and an Olympus PlanC ⫻10/0.25 objective lens (Olympus, Tokyo, Japan). DP2-BSW acquisition software (Olympus) was used to acquire images. Immunofluorescence OCT-frozen TA muscles were cryosectioned at 8 ␮m, fixed in methanol, then washed in phosphate buffered saline (PBS) ACTIVINS PROMOTE MUSCLE WASTING AND CACHEXIA

containing 0.2% Tween 20, and subsequently blocked in 5% normal goat serum. The sections were incubated with primary antibodies overnight at 4°C, and appropriate secondary goat antibody Alexa-Fluor-594 (Life Technologies) was used as the fluorescent label, followed by mounting in Vectashield HardSet (Vector Laboratories, Burlingame, CA, USA). Fluorescent images were taken on an Olympus BX61 microscope. The minimum Feret’s diameter of myofibers was determined using ImageJ software (U.S. National Institutes of Health, Bethesda, MD, USA) by measuring ⱖ180 myofibers/mouse TA muscle. Western blotting TA muscles were homogenized in RIPA-based lysis buffer (Millipore) with protease and phosphatase inhibitor cocktails (Sigma-Aldrich, St. Louis, MO, USA). Lysis was followed by centrifugation at 13,000 g for 15 min at 4°C, and samples were denatured for 5 min at 95°C. Protein concentration was determined using a protein assay kit (Thermo Scientific, Rockford, IL, USA). Protein fractions were subsequently separated by SDS-PAGE using precast 4 –12% Bis-Tris gels (Bio-Rad, Hercules, CA, USA) blotted onto nitrocellulose membranes (Bio-Rad) and incubated with the appropriate antibody overnight and detected as described previously (18). Quantification of labeled Western blots was performed using ImageJ pixel analysis. Densitometric analyses of Western blots are presented as band density and normalized to the control value of 1. Activin ELISAs Activin A was measured using a specific ELISA (Oxford Bioinnovations; ref. 19). Briefly, recombinant activin A standard and samples, diluted in 5% bovine serum albumin (BSA)/PBS (pH 7.4), were treated with SDS (final concentration 3%) and boiled for 3 min. Once cooled, samples were treated with H2O2 (final concentration 2%) and incubated for 30 min at room temperature. Samples were added to duplicate wells in E4 antibody-coated plates and incubated for 1 h at room temperature. Plates were then probed with biotinylated-E4 antibody and incubated overnight at room temperature. After being washed, a streptavidin-horseradish peroxidase (HRP) conjugate was added to the wells and incubated at room temperature for 1 h. After further washes, HRP activity was detected with TMB substrate (3,3=,5,5=-tetramethylbenzidine; Life Technologies). Sample preparation was similar for activin B standard and sample, except that after boiling in SDS, 50 ␮l of Triton assay diluent (25 mM Tris, 0.15 M NaCl, 5% Triton X-100, 0.1% NaN3, and 10% BSA) was added to each tube. Samples were added to duplicate wells in 46A/F antibodycoated plates together with biotinylated-46A/F antibody and incubated for 1 h at room temperature. Detection following addition of streptavidin-HRP and TMB was the same as for activin A (20). Assessment of muscle function Mice were anesthetized via an intraperitoneal injection of 0.9% HEPES-buffered sodium pentobarbitone (Nembutal; 120 mg/kg; Sigma-Aldrich, Castle Hill, NSW, Australia) and supplemental injections of unbuffered Nembutal (60 mg/ kg) were administered to maintain unresponsiveness to tactile stimuli. The assessment of the contractile properties of TA muscles in situ has been described previously (21). Briefly, optimal muscle length (Lo) was determined from the micromanipulation of muscle length to produce maximum isometric twitch force (Pt). Maximum isometric tetanic force (Po) was recorded from the plateau of the 3

frequency-force relationship (10 –300 Hz for 350 ms with 2 min rest between stimuli). Muscles were then subjected to a 4 min intermittent stimulation protocol to induce muscle fatigue. Muscles were maximally stimulated for 1 s every 4 s for the duration of the fatigue protocol. Peak tetanic force was assessed at 5 and 10 min following cessation of the fatiguing stimulation protocol. Statistical analysis The 1-way, 2-way analyses of variance (ANOVAs) and repeatedmeasures ANOVAs were used to assess statistical differences across conditions, with the Student-Newman-Keuls post hoc test used for comparisons between the specific group means. Comparisons between 2 conditions only utilized the Student’s t test. Values of P ⬍ 0.05 were considered significant. Data are presented as means ⫾ sem, unless otherwise stated.

RESULTS Elevated circulating activin alone is sufficient to induce cachexia To determine whether activin A and activin B expression is elevated in human malignancy, we measured

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levels in a range of cancer cell lines of human origin. We found that expression of activin A and B was increased in many of the lines we examined, particularly those that displayed a high degree of malignancy (Supplemental Fig. S1A). To determine whether activins, in the absence of other tumor-derived factors, can induce systemic wasting, we used rAAV6 vectors to express activin A from the right hindlimb muscles of 20-wk-old C57BL/6 mice. In this study, the hindlimb muscles act as a local source of activin A expression into the systemic circulation. Mice injected with control vector gained ⬃10% of their starting body mass (3.8⫾0.4 g) over 10 wk (Fig. 1A). In contrast, mice injected with increasing doses of rAAV6:activin A demonstrated weight loss in a dose-dependent manner (Fig. 1A), with the highest dose of rAAV6:activin A (6⫻1011 vg) inducing a 12.4% loss of starting mass (⫺4.2⫾1.1 g; Fig. 1A). These reductions in body mass in rAAV6: activin A-injected mice correlated inversely with elevated serum activin A levels (7- to 24-fold; Fig. 1B), and body composition analysis identified loss of fat mass (⬃50%) and lean mass (⬃9%) (Fig. 1C, D). The loss of

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Figure 1. Elevated circulating activin A induces systemic wasting. A) Right hindlimb muscles of separate cohorts of 20-wk-old C57BL/6 mice were injected with control or activin A-expressing rAAV6 vectors (2, 4, or 6⫻1011 vg). Body weights were measured weekly and plotted as percentage change from week of injection (n⫽4 – 6, 1-way ANOVA at 10 wk postinjection). B) Terminal samples of blood were collected and activin A levels were determined by specific ELISA (n⫽4 – 6, 1-way ANOVA). C, D) Fat mass (C) and lean body mass (D) were measured using quantitative magnetic resonance (n⫽4 – 6, unpaired Student’s t test). E) At the time of death, muscles distant to the injection site, including the left TA, were excised and weighed (n⫽4 – 6, unpaired Student’s t test). *P ⬍ 0.05, **P ⬍ 0.01, ***P ⬍ 0.001. 4

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lean mass was accounted for by small decreases in the mass of skeletal muscle throughout the body (Fig. 1E and Supplemental Fig. S1B) and of internal organs, such as the liver (data not shown). Activins directly induce profound skeletal muscle atrophy To determine the extent and mechanism of activininduced skeletal muscle wasting, we utilized local injection of rAAV6:activin A into the right TA muscles of mice. Elevated activin A mRNA in the TA muscle occurred within 3 d postinjection (Supplemental Fig. S2A), and at 4 wk, protein levels within the TA muscle increased between 4-fold at the lowest vector dose (109 vg) and 52-fold at the highest vector dose (1011 vg; Fig. 2A). All rAAV6:activin A doses tested caused rapid and sustained muscle loss, resulting in a maximal 65% decrease in mass after 8 wk (Fig. 2B). At 4 wk postinjection, histological examination revealed that the decrease in muscle mass was a product of muscle fiber atrophy (Fig. 2C), as demonstrated by decreases in fiber diameter (Fig. 2D). To determine whether other activin isoforms can regulate muscle mass, we also examined

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the effects of administering increasing doses of rAAV6:activin B. Injection of muscles with rAAV6: activin B caused a similar degree of muscle wasting as for activin A, at all vector doses tested after 4 wk (Fig. 2E, F). As myostatin is regarded as one of the dominant repressors of muscle mass, we compared the effects of activin A and B with those of vectormediated expression of myostatin and TGF-␤1 to determine these ligands’ relative potency in muscle. Notably, of the TGF-␤ family proteins we examined, the activin isoforms were, by far, the most potent negative regulators of muscle mass (Fig. 2E), as demonstrated by the significantly greater muscle wasting caused at lower vector doses. These data demonstrate the potential of activins to promote cachexia even when circulating levels are elevated moderately. As follistatin is an endogenous inhibitor of activins, we next tested the hypothesis that activinmediated muscle wasting could be inhibited via increased expression of follistatin. Accordingly, we observed that the effects of activin A on muscle mass and fiber size were prevented 4 wk after the muscles of mice received rAAV6:activin A with rAAV6:follistatin-288 (Supplemental Fig. S2).

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Figure 2. Activins are potent negative regulators of skeletal muscle mass. Right TA muscles of C57BL/6 mice were injected with rAAV6:activin A (109 or 1011 vg), while the left TA muscles were injected with equivalent doses of control vector. A, B) Elevated local activin levels (A), as determined by tissue ELISA 4 wk after rAAV6 injection (n⫽3– 6, unpaired Student’s t test), caused a rapid and sustained decrease in muscle mass (B; n⫽4 – 6, 1-way ANOVA). C, D) Muscle atrophy after 4 wk was a product of decreased muscle fiber size (reported here as representative hematoxylin- and eosin-stained cryosections (C), and a box and whisker plot (D), comprising minimum, lower quartile, median, upper quartile, and maximum values for myofiber diameter). Scale bar ⫽ 100 ␮m (n⫽3, paired Student’s t test). E) Increasing doses of rAAV6:activin A or rAAV6:activin B in the TA muscle resulted in a progressive loss of muscle mass after 4 wk, which exceeded that achieved by related TGF-␤ ligands, myostatin, and TGF-␤1 (n⫽3– 6); lines are fitted curves. F) Western blots show the relative amounts of activin A, activin B, myostatin and TGF-␤1 expressed in TA muscles examined 4 wk after rAAV6 injection. *P ⬍ 0.05, **P ⬍ 0.01, ***P ⬍ 0.001. ACTIVINS PROMOTE MUSCLE WASTING AND CACHEXIA

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Activin-induced muscle wasting reduces muscle function

Activin promotes muscle wasting via the myostatin signaling pathway

Having shown that activin A can induce skeletal muscle atrophy, we assessed the effect of activinmediated signaling on contractile properties by testing muscle function in situ. Here, we expressed activin A at a low vector dose (109 vg) in the right TA muscle of C57BL/10 mice, with a separate cohort of mice treated with rAAV6:control. At 4 wk after vector administration, TA muscles injected with rAAV6: activin A exhibited reduced muscle mass (Fig. 3A), a 14% decrease in peak isometric twitch force, and a 12% decrease in peak isometric tetanic force (Fig. 3B–D), compared with muscles receiving control vector, but no change in specific force (control, 197⫾8; activin A, 188⫾8 kN/m2, P⫽0.96). As patients with cachexia demonstrate reduced fatigue resistance, as well as reduced strength, we utilized a repeat stimulation protocol designed to assess sustained contractile performance. A 4 min intermittent stimulation protocol designed to elicit muscle fatigue demonstrated that activin A-expressing muscles exhibited reduced absolute force-producing capacity (Fig. 3E), increased loss of force (relative to initial maximal capacity) with repeated stimulation, and reduced recovery of force-producing capacity after the completion of the repeated stimulation challenge (Fig. 3F).

Myostatin, an activin-related TGF-␤ protein, is a negative regulator of muscle mass. Activation of the Smad2/3 transcription factors by myostatin has been shown to decrease protein synthesis by inhibiting activation of the Akt/mTOR pathway and increase protein degradation by up-regulating the muscle-specific ubiquitin ligases MuRF-1 and atrogin-1 (2, 22). We therefore assessed whether activin A induced muscle wasting through these same mechanisms. rAAV6:activin A administration (109 vg) to the TA muscles of C57BL/6 mice caused an increase in Smad3 phosphorylation after 7 d (Fig. 4A), which was associated with reduced phosphorylation of AktS473, a mediator of insulin-like growth factor (IGF)-stimulated protein synthesis. Phosphorylation of effector molecules downstream of Akt in the protein synthesis pathway, pS6KT389 and S6RPS235/236, was also reduced (Fig. 4A), thereby demonstrating suppression of this established signaling cascade. As Akt also regulates the expression of key muscle-specific ubiquitin ligases involved in turnover of myofibrillar proteins, we found an up-regulation of atrogin-1 mRNA (Fbxo32) but not MuRF-1 (Trim63) (Fig. 4B). Activin A also increased expression of p21 mRNA (Cdkn1a), a cyclin-dependent kinase inhibitor, which has been implicated in blocking the activation of muscle satellite cells (ref. 23 and Fig. 4B). Together,

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Figure 3. Activin A reduces peak force production and increases fatigability of TA muscles in situ. A, B) Intramuscular injection of rAAV6:activin A (A) at 109vg in C57BL/10 mice reduced muscle mass (n⫽11–12, unpaired Student’s t test) and peak twitch force (B) of TA muscles in situ after 4 wk, compared with the TA muscles from mice injected with control vector (n⫽10 –12, unpaired Student’s t test). C, D) Examination of the frequency-force relationship revealed that intramuscular injection of rAAV6:activin A reduced tetanic force over a range of stimulation frequencies (C; n⫽5–10, group main effect) and decreased peak tetanic force (D; n⫽10 –11, unpaired Student’s t test). E, F) Intramuscular injection of rAAV6:activin A also reduced absolute force (E) and normalized force (F) during and after 4 min of intermittent stimulation (n⫽6 –10, group main effect). *P ⬍ 0.05, **P ⬍ 0.01, ***P ⬍ 0.001. 6

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60

+

-

0.5

1

0

Trim63/ 18S mRNA

Con

Act A

1.0

+

1.0

Con

*

Con

**

0.5

Act A

C 80 2.0

1.0

0

Con

B Fbxo32/ 18S mRNA

*

p-S6RP/S6RP

1.0

-

1.5

1.5

Act A/Con TA mass

**

p-S6K/pS6K

2.0

p-Akt/Akt

p-Smad3/Smad3

p-Smad3 (52 k) -

Con Act A

75

50

25

Act A 0

WT

Mstn-/-

Figure 4. Activin A engages the myostatin signaling pathway, but is not dependent on myostatin to induce muscle atrophy. Right TA muscles of C57BL/6 mice were injected with rAAV6:activin A at 109 vg, while the left TA muscles were injected with an equivalent dose of control vector. A) Western blot analysis of TA muscles 7 d after vector administration demonstrated that activin A promoted Smad3 phosphorylation, which was accompanied by decreased phosphorylation of AktS473, S6KT389 and S6RPS235/236 (n⫽4 –7, paired Student’s t test). B) qRT-PCR indicated that activin A significantly increases transcription of Fbxo32 (atrogin-1) and Cdkn1a (p21) during muscle wasting (n⫽5– 6, paired Student’s t test) after 7 d. C, D) Myostatin-deficient mice are more susceptible to activin-induced atrophy (C; n⫽4 –7, paired Student’s t test), demonstrating exacerbated muscle fiber atrophy (D; n⫽3). *P ⬍ 0.05, **P ⬍ 0.01, ***P ⬍ 0.001.

these results suggest that activin utilizes the myostatin signaling pathway within skeletal muscle to promote muscle atrophy. In light of these findings, we next considered whether activin-induced wasting was dependent on the presence of myostatin. Myostatin-deficient Mstn⫺/⫺ mice have nearly twice the muscle mass of wild-type mice (24), but following rAAV6:activin A administration, the masses of TA muscles in Mstn⫺/⫺ and wild-type mice were not significantly different (Fig. 4C). In terms of relative mass change, the TA muscles of wild-type mice decreased 23% from control values by 4 wk after rAAV6:activin A administration, whereas the treated TA muscles of Mstn⫺/⫺ mice were reduced in mass by 47% (Fig. 4C). Histological examination revealed that activin exacerbated muscle fiber atrophy in Mstn⫺/⫺ mice (Fig. 4D, E), concomitant with higher ACTIVINS PROMOTE MUSCLE WASTING AND CACHEXIA

expression of Cdkn1a (p21), but no difference in Fbxo32 (atrogin-1) or phosphorylation of S6 ribosomal protein (S6RP; Supplemental Fig. S4B, C). These data demonstrate that skeletal muscles from Mstn⫺/⫺ mice are more sensitive to activin-induced atrophy. Transcription profile of activin-treated skeletal muscle To further understand the mechanisms underlying activin-induced muscle atrophy, we subjected RNA extracted from skeletal muscles harvested 7 d after administration of rAAV6:activin A (109 vg) or rAAV6:control to transcriptional profiling via next-generation sequencing. In muscles injected with rAAV6:activin A, we found that 131 genes were differentially expressed, with 7

fold change ⬎ 1.5 and nominal P ⬍ 0.05 as cutoffs. Notably, the majority of these genes (94%) were upregulated following activin treatment. To further delineate activin-responsive genes that may be involved in regulating muscle growth, we compared our data set to gene changes reported previously in response to muscle hypertrophy caused by administration of soluble activin type II receptor (sActRIIB; 25). By this comparison, we identified 39 genes that were up-regulated by activin A, but significantly down-regulated by sActRIIB (Table 1). We verified these findings for a subset of 6 genes (Cyr61, Igfn1, Scx, Csrp3, Tbc1d1, and Pi16) using qRT-PCR or Western blot (Supplemental Fig. S4A–F). Of the genes significantly up-regulated by activin A alone, Tnfrsf12a [also known as fibroblast growth factor-inducible receptor 14 (Fn14)], the receptor for tumor necrosis factor (TNF)-like weak inducer of apoptosis (TWEAK) was identified as a candidate of particular interest, as TWEAK-Fn14 signaling can promote skeletal muscle atrophy (26). Consistent with increased

transcription of Tnfrsf12a in muscles undergoing activin-induced atrophy, Western blot analysis demonstrated a significant increase in the abundance of Fn14 protein in muscles examined 56 d after administration of rAAV6:activin A (Supplemental Fig. S4G). Activin induces fibrosis in skeletal muscle Activation of TGF-␤ signaling pathways has been associated with a number of diseases where fibrosis is prominent (27). We therefore examined whether overexpression of activin A in skeletal muscle caused fibrosis. Histological examination of TA muscles harvested 56 d after rAAV6:activin A injection (109 or 1011 vg) demonstrated significant endomysial cellular infiltration (Fig. 5A). Immunofluorescent labeling identified cells positive for fibroblast-specific protein 1, indicating transformation of fibroblasts into collagen-secreting myofibroblasts (Fig. 5B). Accordingly, the presence of myofibroblasts within skeletal muscle was associated

TABLE 1. Genes up-regulated by activin A treatment of tibialis anterior muscle, which have previously been shown to be significantly down-regulated by sActRIIB treatment (25) Gene

Angptl7 Cyr61 Igfn1 Pmepa1 Zmynd17 Cdkn1a Prima1 Csrp3 Fxyd6 Eln Serpine1 Myh2 Ddah1 Hspb7 Klhl34 Lmod2 Pi16 Tmem9 Dusp18 Smtnl1 Skil Fos Itgb5 Ctgf Ddit4 Tmem100 Meg3 Nudt18 Art5 Nos1 Rhou Tbc1d1 Scx Igfbp6 Rhbdf1 Mgp Phlda3 Clec3b Gpx3

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Angiopoietin-like 7 Cysteine-rich protein 61 Immunoglobulin-like and fibronectin type III domain-containing 1 Prostate transmembrane protein, androgen induced 1 Zinc finger, MYND domain-containing 17 Cyclin-dependent kinase inhibitor 1A (P21) Proline-rich membrane anchor 1 Cysteine- and glycine-rich protein 3 FXYD domain-containing ion transport regulator 6 Elastin Serpin peptidase inhibitor, clade E, member 1 Myosin, heavy polypeptide 2, skeletal muscle, adult Dimethylarginine dimethylaminohydrolase 1 Heat-shock protein family, member 7 Kelch-like 34 (Drosophila) Leiomodin 2 (cardiac) Peptidase inhibitor 16 Transmembrane protein 9 Dual-specificity phosphatase 18 Smoothelin-like 1 SKI-like oncogene FBJ osteosarcoma oncogene Integrin ␤ 5 Connective tissue growth factor DNA-damage-inducible transcript 4 Transmembrane protein 100 Maternally expressed 3 (non-protein coding) Nudix (nucleoside diphosphate linked moiety X)-type motif 18 ADP-ribosyltransferase 5 Nitric oxide synthase 1, neuronal Ras homolog family member U TBC1 domain family, member 1 Scleraxis Insulin-like growth factor binding protein 6 Rhomboid family 1 (Drosophila) Matrix Gla protein Pleckstrin homology-like domain, family A, member 3 C-type lectin domain family 3, member B Glutathione peroxidase 3

3.32E-07 2.86E-22 3.24E-21 4.03E-09 2.43E-15 2.39E-18 2.87E-03 0.007 7.78E-19 1.02E-07 0.019 2.81E-04 5.32E-05 7.77E-05 1.26E-04 2.05E-05 2.72E-03 1.51E-04 0.035 2.57E-04 5.48E-04 0.025 2.23E-04 4.00E-04 6.27E-04 0.016 2.41E-03 1.59E-03 4.00E-04 1.66E-03 0.015 0.010 8.20E-04 3.48E-04 0.021 8.20E-04 0.030 0.018 0.050

5.5 3.8 3.3 2.9 2.7 2.7 2.4 2.4 2.3 2.2 2.2 2.1 2.1 2.1 2.0 2.0 2.0 2.0 1.9 1.9 1.9 1.8 1.8 1.8 1.8 1.8 1.8 1.8 1.8 1.7 1.7 1.7 1.7 1.7 1.7 1.7 1.6 1.6 1.5

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3

p=0.053 2

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109 vg

Con 1011

Act A 1011

CTGF/18S mRNA

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5

p=0.054

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Con 109

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109 vg

1011 vg

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109 vg

1011 vg

**

**

109 vg

1011 vg

Act A 109

Act A 1011

Comp/18S mRNA

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1011 vg

** 8

0

1011 vg

*

109 vg

C

p=0.052

Eln/18S mRNA

Act A 1011

D

Adamts8/18S mRNA

Con 1011

Act A 109

*

20

10

**

0

109 vg

10

Mfap4/18S mRNA

Con 109

Col1a1/18S mRNA

A

1011 vg

5

0

Con Act A

Figure 5. Activin A induces muscle fibrosis. A–C) Sections of TA muscle collected from C57BL/6 mice 56 d after injection of rAAV6:control into the left TA or rAAV6:activin A into the right TA (109 or 1011 vg), were stained with hematoxylin and eosin (A), fibroblast-specific protein-1 (B), or Masson’s trichrome (C). Scale bars ⫽ 100 ␮m. D) qRT-PCR analysis of TA muscles demonstrating increased transcription of extracellular matrix genes in muscles administered rAAV6:activin A (n⫽3– 4, paired Student’s t test). *P ⬍ 0.05, **P ⬍ 0.01.

with substantial collagen deposition (Fig. 5C). Analysis of gene expression via qRT-PCR indicated that, in addition to collagen, a number of other extracellular matrix genes were up-regulated in response to rAAV6: activin A administration, including Eln, CTGF, AdamTS8, Comp, and MFAP4, the transcription of which increased in a vector dose-dependent manner (Fig. 5D). These data demonstrate that increased expression of activin in muscles promotes fibrosis that is broadly consistent with pathology observed in some chronic myopathies. Activin-induced muscle pathology is reversible Because it may not always be possible to administer interventions to patients with cancer before the onset of muscle wasting and cachexia, we sought to test the potential for rehabilitation from activin-induced wasting. We generated an inducible rAAV6:activin A vector, controlled by the presence or absence of doxycycline. Injection of this vector into the TA muscles of C57BL/6 mice, subsequently fed doxycycline-supplemented chow for 4 ACTIVINS PROMOTE MUSCLE WASTING AND CACHEXIA

wk, caused a 50% reduction in muscle mass (Fig. 6A). Termination of activin overexpression via removal of doxycycline from the diet led to a progressive increase in TA muscle mass, such that by 8 wk after doxycycline withdrawal, the mass of treated muscles was indistinguishable from control muscles (Fig. 6A). Histological examination indicated that changes in TA muscle mass following activin overexpression and withdrawal were paralleled by changes in muscle fiber size (Fig. 6B, C). Notably, at 2 wk after activin withdrawal, the endomysial cellular infiltration was reduced substantially (Fig. 6B). Accompanying the observed reductions in myofibroblast numbers was a resolution of collagen deposition (Fig. 6D), and suppression of the extracellular matrix gene program that had previously been initiated by activin (Fig. 6E). Mechanistically, the abrogation of activin expression led to attenuation of Smad3 phosphorylation and a corresponding increase in activation of molecules downstream of Akt in the protein synthesis pathway, including S6RP (Fig. 6F). Together, these results indicate that the deleterious effects of increased 9

D

TA mass (mg)

80

Con Act A

E

Con Act A

Con

60

10

ns

*

** 40

Eln/18S mRNA

A

***

*** 20

4+

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4+

4+,1-

4+,2-

4+,4-

4+,8-

6

CTGF/18S mRNA

4+, 24+, 8-

4+

4+,1-

4+,8-

p=0.068

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2

p=0.065

ns

0

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4+, 4-

F

4+ TRE:Act A

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ns

ns 0

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-

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+

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+

Act A (13 k) p-Smad3 (52 k) p-S6RP (37 k) S6RP (37 k)

C p-S6RP/S6RP

TA fiber size (µm)

3

Con Act A

100

75

50

25

0

4+

**

*

*** *** ** *

* *

Con Act A

* 2

ns

1

** 0

4+,1-

4+,2-

4+,4-

4+,8-

Weeks with induction (+) and withdrawal (-)

4+

4+,1-

4+,8-

Weeks with induction (+) and withdrawal (-)

Figure 6. Effects of activin A on skeletal muscle are reversible. Right TA muscles of C57BL/6 mice were injected with a rAAV6 encoding the Tet-On protein under a CMV promoter, and a rAAV6 encoding activin A under a TRE promoter. Left TA muscles of these mice were injected with a rAAV6 encoding the Tet-On protein under a CMV promoter, and a control vector. A) Activin A expression was induced for 4 wk, after which transduction was switched off by substitution of diet to doxycycline-free chow for up to 8 wk. TA muscle mass was determined following death of mice at the indicated time-points (n⫽4 – 6, paired Student’s t test). B, C) Initial muscle atrophy, and subsequent recovery from atrophy, correlated with changes to muscle fiber size, (n⫽3– 4, paired Student’s t test). Scale bar ⫽ 100 ␮m. D, E) Activin A-induced collagen deposition, identified by Masson’s trichrome stain, was completely resolved 8 wk after removal of doxycycline from the diet (D) and was accompanied by the down-regulation (E) of ECM genes elastin (Eln) and connective tissue growth factor (CTGF; n⫽3– 4, paired Student’s t test). F) Western blot analysis and densitometry of TA muscles correlated with expected changes in phosphorylation of Smad3 and S6RP (n⫽3– 4, paired Student’s t test). ns, not significant. *P ⬍ 0.05, **P ⬍ 0.01, ***P ⬍ 0.001.

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activin signaling on the attributes of skeletal muscle are reversible with adequate activin inhibition.

DISCUSSION Activin A and B isoforms promote signaling via engagement with ActRIIB in many cell types and organs systems. In mouse models of cancer cachexia where activin levels became elevated, treatment with soluble ActRIIB restored muscle mass, and consequently prolonged survival, despite continued tumor growth (5). These data have implicated activins as promoters of cachectic muscle wasting, but the effect of activins on musculature has remained unclear, as those studies examined tumorigenic models where other factors associated with tumor development and progression constitute confounding influences. To determine whether activin signaling in the absence of tumor-derived factors was sufficient to induce cachexia, we used rAAV6 vectors to express activins from the hindlimb muscles of wild-type mice. Increased activin expression in treated limbs induced severe local muscle wasting and led to increases in circulating activin A levels above those noted in cachectic mice bearing colon-26 (C26) tumors (unpublished results) but not as high as levels reported in cachectic inhibin-deficient mice (15). We found that increased expression of activin A alone caused systemic wasting, with graded increases in circulating levels corresponding to a more profound loss of body mass. Activin-induced weight loss was primarily due to a substantial decrease in skeletal muscle and adipose mass, which represent the most debilitating aspects of cachexia. Interestingly, the extent of muscle atrophy induced by increasing circulating activin A levels 25-fold for 10 wk was similar to that observed in mice bearing C26 tumors for only 3 wk, despite the fact that activin levels are only 20-fold elevated in the latter model (unpublished results). It is probable that other tumor-derived factors operating in concert with activin likely exacerbate muscle wasting. The inflammatory cytokines interleukin (IL)-1␤, IL-6, TNF-␣, and TWEAK have been implicated as procachectic factors, and thus represent the leading candidate molecules for this synergistic role in the etiology of cancer cachexia (4, 26, 28). In support, activin A increased expression of the TWEAK receptor Fn14 in skeletal muscle to a level previously shown to regulate denervation-induced atrophy (26, 29). As endogenous expression of Fn14 in skeletal muscle is negligible, this activinmediated expression of Fn14 could represent a potential mechanism by which increased concentrations of activins and TWEAK, as a consequence of tumor progression, accumulate in the circulation and cooperatively promote muscle wasting in cachexia. Myostatin is regarded as one of the most potent negative regulators of muscle mass; however, we found that the activin isoforms were 100-fold more potent in causing muscle wasting when expressed via local administration of rAAV6 vectors. This disparity in activity likely arises from differences in how these TGF-␤ family members interact with their respective prodomains during synthesis and ACTIVINS PROMOTE MUSCLE WASTING AND CACHEXIA

secretion. For activins, the affinity of prodomain binding is insufficient to interfere with receptor binding or suppress biological activity (9), whereas myostatin binds its prodomain with high affinity and is secreted from the cell in a latent form (30). Activation of myostatin requires cleavage of its prodomain by tolloid-like metalloproteases (30), which are conceivably rate limiting within skeletal muscle. The magnitude of muscle loss induced by activin A and activin B exceeded those commonly reported for factors acting on other atrophy-related pathways. These findings may explain the propensity of activins to promote cachexia when elevated relatively moderately in circulation. Mechanistically, our data support a role for activin A to induce muscle wasting by activating the Smad2/3 transcription factors, which increase protein degradation by up-regulating the expression of the muscle-specific ubiquitin ligase atrogin-1 and depress protein synthesis via inhibition of the Akt/mTOR pathway. Interestingly, the magnitude of muscle wasting induced by activin was enhanced in mice lacking myostatin. These data suggest that, in the absence of myostatin, muscles are primed to maximally respond to a Smad2/3 signal. To better understand the molecular mechanisms of activin-mediated muscle atrophy, we used next generation sequencing to identify activin target genes in mouse skeletal muscle. Comparing our data with those from transcriptional profiling of muscles undergoing hypertrophy after treatment with soluble ActRIIB (25), we identified 39 genes that were differentially regulated in response to modifying ActRIIB signaling. Among the differentially regulated genes, we identified 3 up-regulated genes (Phlda3, Ddit4, and DUSP18) that are predicted to disrupt Akt signaling. PHLDA3 is a PH domainonly protein, which competes with Akt for binding to membrane lipids, thereby inhibiting Akt translocation to the membrane and activation (31). This, in turn, would lead to derepression of the TSC1/TSC2 complex and a decrease in mammalian target of rapamycin (mTOR)mediated protein synthesis (32); an effect potentially enhanced by activin up-regulation of Ddit4 (REDD1), an inhibitor of mTOR signaling (33). Reduced Akt activation would also promote nuclear translocation of Foxo3a, where it controls protein degradation pathways, via upregulation of the ubiquitin ligases MuRF-1 and atrogin-1. Normally, stress-activated protein kinase (SAPK) promotes nuclear export of Foxo3a in muscle cells (34). However, activin-mediated up-regulation of the SAPK inhibitor dual specificity phosphatase 18 (DUSP18; ref. 35) would help prolong nuclear localization of Foxo3a and thus support continued transcription of the atrogenes that promote muscle catabolism. Other genes previously associated with muscle atrophy that were up-regulated by activin A, but decreased following soluble ActRIIB treatment, include Nos1 and Igfn1 (25). Nos1 [neuronal nitric oxide synthase (nNOS)] is a component of the dystrophin-glycoprotein complex, which rapidly translocates from the sarcolemma to the cytoplasm during unloading-induced muscle atrophy (36). Nitric oxide production by nNOS in atrophying muscles promotes nuclear accumulation of Foxo3a and 11

the subsequent up-regulation of atrogin-1 and MuRF-1. Activin induction of dimethylarginine dimethylaminohydrolase 1 (Ddah1), which metabolizes asymmetric dimethylarginine, an inhibitor of NOS, could potentiate nNOS-induced wasting (37). Increases in Igfn1 expression following activin treatment may also contribute to wasting, given that IGFN1 is a sarcomeric protein postulated to down-regulate protein synthesis during muscle denervation via interactions with eukaryotic translation elongation factor 1A (eEF1A; ref. 38). IGFN1 also interacts with kyphoscoliosis peptidase (KY), the knockout of which exhibits a primary degenerative myopathy preceding chronic thoraco-lumbar kyphoscoliosis (39). A role for activin in regulating the important muscle functions of KY, via up-regulation of IGFN1, is supported by the observation that inhibin-deficient mice, where activin levels are dramatically elevated, develop severe thoracic kyphoscoliosis as cachexia progresses (8). The most prominent set of genes differentially regulated by altering Smad2/3 signaling in muscle were those involved in extracellular matrix production/fibrosis (Angptl7, Cyr61, Eln, Serpine1, Ctgf, Mgp, Scx, and Clec3b). This was consistent with increased fibroblast/myofibroblast numbers and extensive fibrosis observed in activin-treated muscles. To determine the potential for rehabilitation from activin-induced fibrosis and muscle atrophy, we generated an inducible rAAV6:activin A system. Strikingly, when activin A overexpression was switched off after 4 wk, the observed pathological phenotype proved to be reversible, indicating that muscles can fully recover from a sustained activin A insult. These data have significant therapeutic implications because they demonstrate that the severe adverse effects of activin A on muscle appear amenable to treatment after diagnosis. Increasing evidence indicates that TGF-␤ family members that utilize the ActRIIB receptor, particularly myostatin, activin A, and activin B are highly catabolic factors when elevated systemically (5, 40). The physiological relevance of these findings is highlighted by reports documenting elevated serum activin A in patients with cancer (6, 7) and our discovery that nearly half of all human cancer cell lines we examined expressed high levels of activin A and/or activin B. Given the propensity of these growth factors to be associated with cachexia in murine cancer models (5), future studies need to confirm whether elevated serum activins play a role in the etiology of human cachexia. Our results showing that severe muscle wasting and fibrotic accumulation caused by activin overexpression were fully reversible indicate that rehabilitation from activin-induced wasting is possible. Strategies being developed to ameliorate muscle wasting via inhibition of ActRIIB signaling include the use of ligand traps (soluble receptors, modified propeptides), binding proteins (follistatin), inhibitors of Smad signaling, or monoclonal antibodies (5, 15, 41, 42). While these interventions are capable of diminishing the effects of activins, many also perturb the effects of other TGF-␤ family members, with consequences for generating off-target effects. With our demonstration that activins are produced by malignant cell lines and that activins alone can cause muscle wasting, we 12

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contend that these specific TGF-␤ family members are critical regulators of muscle mass. Interventions that specifically target these molecules may offer the potential to effectively prevent muscle wasting associated with life-threatening cancer cachexia, and other conditions where excessive ActRIIB signaling is implicated. The authors thank Drs. Jonathan Davey and Kevin Watt (Division of Cell Signaling and Metabolism, Baker IDI Heart and Diabetes Institute) and Emily Kelly, Karen Chan, and Enid Prusyers (Prince Henry’s Institute) for technical assistance; David de Kretser and Susan Hayward (Monash Institute of Medical Research, Melbourne, VIC, Australia) for assistance with the activin ELISAs; Drs. Mark Ziemann, Ross Lazarus, and Assam El-Osta (Division of Diabetic Complications, Baker IDI Heart and Diabetes Institute) for support with bioinformatics analyses; and Prof. Mark A. Febbraio (Division of Cell Signaling and Metabolism, Baker IDI Heart and Diabetes Institute) for technical advice and assistance concerning measurement of animals’ body composition. The human cancer cell lines were a gift from Assoc. Prof. Guiying Nie and Dr. Colin Clyne (Prince Henry’s Institute). Myostatin-null mice were a gift of Prof. Se-Jin Lee (Johns Hopkins University, Baltimore, MD, USA). This work was supported by grant funding (526648, 566820, 1006488) from the National Health and Medical Research Council (NH&MRC) of Australia. P.G., C.A.H., and K.T.M. are supported by Career Development Fellowships (1046782, 1013533, and 1023178) from the NH&MRC. P.G. was previously supported by a senior research fellowship, sponsored by Pfizer Australia. The Baker IDI Heart and Diabetes Institute and Prince Henry’s Institute are supported in part by the Operational Infrastructure Support Program of the Victorian government. Author contributions: J.L.C., K.L.W., C.A.H. and P.G. designed the research; J.L.C., K.L., K.T.M., R.E.T., Y.M., and H.Q. performed the experimental work; C.E.W., G.S.L., C.A.H., and P.G. contributed reagents and analytical tools and technical advice; J.L.C., C.E.W., K.T.M., and P.G. analyzed the data; J.L.C., G.S.L., C.A.H., and P.G. wrote the manuscript. The authors declare no conflicts of interest.

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11. 12.

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14. 15.

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