1. Introduction
The liver and skeletal muscle systems have a well-established interdependent endocrinological connection in both physiological and pathological states. One example of this inter-dependency is observed through their mutual contribution to the maintenance of glucose homeostasis through storage and metabolic mechanisms. In contrast, patients with liver cirrhosis exhibit abnormal capacity to store glucose as glycogen in skeletal muscle [1]. Moreover, skeletal muscle wasting, or sarcopenia, is recognized as a major complication in patients with cirrhosis, non-alcoholic fatty liver disease (NAFLD), and the more severe form known as nonalcoholic steatohepatitis (NASH) [2,3]. Recent clinical studies demonstrate a linear association of sarcopenia with severity of liver fibrosis in NASH patients [4,5,6]. In addition, the degree of muscle wasting is a strong correlate to adverse clinical outcomes and follow-up hospital costs in these patients [4,7,8,9,10,11,12]. Patients with cirrhotic livers and concomitant muscle atrophy tend to have lower survival rates and even more post-liver transplant complications. Paradoxically, most complications in liver cirrhosis patients resolve following a successful transplant, except for the underlying muscle wasting [13]. Thus, the ability to treat muscle mass loss is paramount for the clinical management of these patients [14,15,16]. Unfortunately, effective therapies are lacking because the signaling mediators of the liver–muscle axis remain unclear [17]. Therefore, it is a priority to identify and delineate the systemic cues that initiate and promote this sequela of liver diseases.
Carbon tetrachloride (CCl4) is the most commonly used toxin to induce experimental liver fibrosis [18]. CCl4 is metabolized mainly by Cyp2E1 in hepatocytes. This process produces toxic free radicals causing massive centrilobular necrosis in the liver [19]. This in turn triggers inflammatory responses manifested by the activation of immune cells and the production of proinflammatory factors by these cells [20]. Subsequently, hepatocyte death and inflammation induce robust repair responses, indicated by hepatocyte proliferation and the activation of hepatic stellate cells (HSCs). Activated HSCs produce excessive collagens, initiating liver fibrosis [21]. The CCl4 model recapitulates human liver fibrosis in many aspects, including hepatocyte necrosis, inflammation, HSC activation, and liver fibrogenesis [18].
The TGFβ superfamily of secreted proteins include activins, growth differentiation factors (Gdfs), and bone morphogenetic proteins (BMPs). They engage with activin type II and type I receptor complexes to initiate Smad signaling to modulate organ development, growth, homeostasis, and repair [17,22,23]. Notably, it has been proposed that members of the TGFβ superfamily play integral roles in homeostasis and disease states of both the liver and skeletal muscle, and certain members may even serve as a nexus between the two organs [24,25,26]. In the present study, we set out to determine whether TGFβ ligands are causal mediators responsible for the deleterious communication between the injured liver with normal or injured skeletal muscle. We also evaluated the potential to prevent or reverse liver injury-induced muscle atrophy by inhibiting TGFβ superfamily signaling. We utilized both a fusion protein (ActRIIB-Fc) and a neutralizing antibody to myostatin (Gdf8) to demonstrate the importance of this signaling axis in this pre-clinical setting. ActRIIB-Fc is a fusion protein consisting of the extracellular ligand-binding domains of activin type IIB receptor with the Fc portion of mouse immunoglobulin G (IgG). ActRIIB-Fc primarily binds to and inhibits activins, Gdf8, and other TGFβ superfamily members [27]. We found that Gdf8 largely mediates the negative crosstalk of injured liver with skeletal muscle and simultaneously promotes liver injury.
2. Materials and Methods
2.1. Approval of Animal Studies
All mouse experiments were performed with the approval of the Institutional Animal Care and Use Committee of Indiana University—Purdue University Indianapolis (SC337R) and are in accordance with the National Institute of Health Guide for the Care and Use of Laboratory Animals. For all studies described here, ten-week-old C57BL/6 female or male mice were used (Harlan, Indianapolis, IN, USA). Animals were housed in a room with controlled temperature (22 ± 2 °C) and a 12:12 h light–dark cycle (lights on at 6:00 a.m.) with ad lib access to food (TD 5001 with 0.95% calcium and 0.67% phosphorus, Teklad, Madison, WI, USA) and water.
2.2. Carbon Tetrachloride Liver Fibrosis Model
Hepatic fibrosis was induced by 10 mL/kg intraperitoneal injections of 1:10 diluted carbon tetrachloride (CCl4) (Sigma Aldrich, St. Louis, MO, USA) in corn oil twice weekly for a total of 6 to 9 weeks for chronic studies [28]. For acute studies, tissues were harvested 6, 24, or 48 h following a single injection of CCl4.
2.3. Skeletal Muscle Injury Model
Muscle injury was induced as previously described [29], with slight modifications. Briefly, muscle injury was induced by a 100 µL injection of a 10 µM cardiotoxin (CTX) (Sigma-Aldrich, St. Louis, MO, USA; part #C3987) solution into the gastrocnemius muscle with a three-point injection technique to fully cover the lateral and medial gastrocnemius.
2.4. Gene Expression Analysis
Total RNA was extracted from isolated tissues using TRIzol reagent (Life Technologies, Grand Island, NY, USA) and treated with RNase-free DNase I (ThermoFisher, Waltham, MA, USA). RNA concentrations and A260/A280 ratios were determined by a NanoDrop Microvolume Spectrophotometer (ThermoFisher). RNA integrity was confirmed using RNA gel electrophoresis. Total RNAs (1 µg/sample) were reverse transcribed using a High-Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA, USA). The cDNAs were assayed for genes of interest using TaqMan Gene Expression Analysis (Applied Biosystems) and quantified by the 2−ΔΔCt method. The initial hold step (50 °C for 2 min followed by 95 °C for 10 min) and 40 cycles of a two-step PCR (92 °C for 15 s and 60 °C for 1 min) were performed.
2.5. Protein Quantification
Tissue lysates were generated using 1 mL per 100 mg tissue in lysis buffer (Cell Signaling Technologies, Danvers, MA, USA). Total protein contents were quantified using a Pierce BCA Protein Assay Kit (Thermo Scientific). Gdf8 and Smad2/3 proteins were quantified by ELISA methods (GDF8/Myostatin Quantikine ELISA kit, Cat #DGDF80, R&D Systems, Minneapolis, MN, USA; PathScan phospho-Smad2 Ser465/467/Smad3 ser423/425 and total for Smad 2 and 3, kits #12001, 12002, 7244, Cell Signaling, Danvers, MA, USA). Procedures were followed according to the manufacturer’s guidelines.
2.6. Histology and Immunohistochemistry
Muscle or liver tissue was evaluated using hematoxylin and eosin (H&E) or Masson’s trichrome staining (Abcam, Cambridge, MA, USA). For each muscle, distribution of the fiber diameter was calculated by analyzing ~200 myofibers using digital slide scanning (ScanScope XT, Aperio, Vista, CA, USA). A collagen proportionate area (CPA) was quantified using the HALO Image Analysis Platform (Albuquerque, NM, USA) of liver sections stained with Masson’s trichrome. FFPE liver sections were assayed for Gdf8 content via a commercial antibody (LSBio, LS-C37420/170820, Washington, DC, USA).
2.7. Human Liver Sample Collection
Liver tissue was collected from individuals with established cirrhosis awaiting liver transplantation at the time of their liver transplantation procedure in the operating room. Demographic data, etiology of cirrhosis, and other relevant information such as medication, alcohol use, and smoking history were captured at the time of enrollment. Liver tissue samples were snap frozen using liquid nitrogen and stored at −80 °C until use. All samples were collected and handled equally except for the duration of storage. This study was reviewed and approved by the Institutional Review Board (IUPUI IRB: EX0904-11).
2.8. ActRIIA-Fc and ActRIIB-Fc Proteins and Antibodies
The ActRIIA-Fc and ActRIIB-Fc proteins described in this report were expressed in stably transfected Chinese hamster ovary cells and were generated at Eli Lilly (Indianapolis, IN, USA). Isolation of the chimeric proteins from concentrated cell culture supernatants was performed using a two-step purification method. In the first step, crude conditioned cell culture media containing the specific variant was captured onto Mab select sepharose (GE Healthcare, Buckinghamshire, UK) under high salt conditions (1 M sodium chloride) and eluted using a step-gradient of 10 mM sodium citrate, pH 3.0. Pooled protein was concentrated using an Amicon Ultra-15 concentrator (Millipore, Boston, MA, USA) and further purified using a Superdex G200 preparative gel-filtration step (GE Healthcare, Buckinghamshire, UK). These steps generally resulted in protein purity of > 95%, as assessed by SimplyBlue staining (Invitrogen, Carlsbad, CA, USA) SDS-PAGE and analytical gel filtration on a TSKG3000SWXL column (Tosoh Bioscience, Tokyo, Japan). GDF8 antibody was generated by Eli Lilly and Company and its properties were described previously [30,31]. Activin A antibody was purchased from R&D System (Cat #MAB3381, Minneapolis, MN, USA).
2.9. Body Composition
Lean body mass and fat mass were measured in live conscious animals using nuclear magnetic resonance (NMR; Echo Medical Systems, Houston, TX, USA).
2.10. Cell Culture and In Vitro Assays
Experiments were performed using primary hepatocytes isolated from mice. Briefly, under anesthesia, the peritoneal cavity was opened, and the liver was perfused in situ via the portal vein for 4 min at 37 °C with calcium–magnesium (CM)-free HEPES buffer and for 7 min with CM-free HEPES buffer containing type IV collagenase (35 mg/100 mL) and CaCl2 (10 mM). Cells were used only if the cell viability was above 90% as assessed by trypan blue exclusion. After three centrifugations (44 g for 2 min) in Leibovitz’s L-15 washing media supplemented with 0.2% bovine albumin, cells were plated onto 24-well or 96-well plates (26,000 cells/cm2). Cells were cultured in high-glucose (25 mM) DMEM supplemented with 10% FBS. All culture media contained penicillin (100 units/mL) and streptomycin (100 μg/mL). After cell attachment for 2 h, the medium was replaced with fresh medium supplemented with 10% fetal bovine serum (FBS). PMH cultures were maintained under 5% CO2 atmosphere at 37 °C. CCl4 or corn oil was administered at 0.5% volume directly onto plated hepatocytes for 24 h prior to media collection. C2C12 myoblasts were obtained from ATCC (Manassas, VA, USA) and were thawed and plated per ATCC protocols in 10% FBS-supplemented media. Prior to cells reaching confluence, they were differentiated in growth media supplemented with 2% horse serum for 5 days with daily media changes. Collected hepatocyte media were diluted 1:10 into myoblast differentiation media and supplemented with IgG or ActRIIB-Fc at 100 ng/mL prior to addition to C2C12 cells, whereas corn oil media received no treatment. After 5 days of daily media change, muscle cells were imaged and myotube length quantified using Aperio ImageScope 12.3 software. Hepatocyte media aspartate aminotransferase (AST) and alanine aminotransferase (ALT) were assayed with a Hitachi instrument. LX-2 cells, a human hepatic stellate cell line, was a gift from Dr. Scott L. Friedman from the Mount Sinai School of Medicine (New York, NY, USA). They were cultured in DMEM supplemented with 2% FBS (Gibco, Invitrogen, Carlsbad, CA, USA).
2.11. Data Analysis and Presentation
All results were expressed as mean ± standard error of the mean (SEM). All datasets were assessed for normality via Shapiro–Wilk test. Statistical analysis was performed by ordinary ANOVA if normality assessments were passed or by Kruskal–Wallis if normality assessments were not passed. Significant differences were defined when p-value ≤ 0.05.
3. Results
3.1. Acute Liver Injury Induces Rapid and Negative Responses in Skeletal Muscle
To understand how liver injury affects skeletal muscle, we evaluated the acute muscle response to a model of liver injury induced by carbon tetrachloride (CCl4) [32]. This model consistently demonstrates muscle wasting reminiscent of clinical observations of liver disease patients [33]. As early as 6 h following a single CCl4 administration, the expression of ubiquitin ligase Trim63 and Fbxo32, critical regulators of early muscle turnover [34], was markedly increased and persisted for 3 days (Figure 1A). Additionally, expression of three transcription factors (Lif, Myod1, and Pax7) essentially required for myogenesis were rapidly downregulated, whereas ankyrin repeat domain 2 (Ankrd2), a powerful regulator of myogenesis and stress responses, was upregulated (Figure 1A). Smad proteins are intracellular signal-transducing proteins known to be regulated by TGFβ family members to modulate myogenesis. We observed alterations in total Smad 2 or 3 protein content 6–24 h post CCl4 injection and increases in phosphorylated Smad2 2 days after CCl4 exposure (Figure 1B–D). These data indicate that skeletal muscle sensitively and negatively responds to liver injury at least via a mechanism that suppresses satellite cell markers, potentially limiting the muscle’s regenerative capacity and potential long-term preservation of lean mass. Moreover, the observed increases in muscle Smad2 activity following CCl4 dosing strongly support a role for TGFβ signaling in the muscle’s response to acute liver injury.
3.2. Neutralization of ActRIIB Ligands but Not Activin A May Prevent the Initiation of Liver Injury-Induced Muscle Atrophy
To evaluate whether ActRIIB ligands mediate the initiation of injured liver–muscle communication in vivo, female mice were treated with IgG, activin A antibody (activin A-Ab), or ActRIIB-Fc 16 h prior to a single dose of CCl4. Activin A was included in this study because it has been suggested by others to be the most likely ActRIIB receptor ligand responsible for muscle mass loss [35]. Three days following CCl4 injection, acute liver injury reduced muscle mass, whereas ActRIIB-Fc but not activin A antibody prevented this event (Figure 1E). However, these changes did not reach statistical significance, which was most likely due to the short period of exposure of the muscle to the injured liver. Notably, the expression of ubiquitin ligase Fbxo32, a potent muscle atrophy promotor, was increased in acutely injured liver, which was attenuated by ActRIIB-Fc but not activin A-Ab (Figure 1F). These data suggest that ActRIIB-binding ligands modulate the initiation of muscle atrophy after liver injury and appear to be independent of activin A.
3.3. Neutralization of ActRIIB Ligands Prevents Chronic Liver Injury-Induced Muscle Atrophy Independent of Gender
To elucidate whether ActRIIB ligands mediate progressive muscle mass loss caused by chronic liver injury, male mice were administered CCl4 twice weekly for 6 weeks. Sixteen hours before the first CCl4 injection each week, mice received weekly IgG, ActRIIA-Fc, ActRIIB-Fc, or a combination of these two fusion proteins, which function as pharmacological ligand traps for their respective receptors (Figure 2A). As a result, CCl4-induced chronic liver injury caused significant muscle mass loss, which was prevented by ActRIIB-Fc but not ActRIIA-Fc. Treatment with ActRIIB-Fc and ActRIIA-Fc combination did not show additive effects in muscle mass relative to ActRIIB-Fc alone, demonstrating that an activin receptor IIB-specific ligand is primarily responsible for the induction of muscle atrophy (Figure 2B). These data suggest that inactivation of ActRIIB ligands prevents muscle atrophy during chronic liver injury.
We performed a similar study to the one described above in female mice. Female mice received weekly IgG, activin A-Ab, or ActRIIB-Fc along with twice weekly CCl4 administrations (Figure 2A). Chronic liver injury induced by CCl4 resulted in a decrease in muscle mass, which was prohibited with ActRIIB-Fc but not activin A-Ab (Figure 2C). Further analysis showed that, in response to chronic liver damage, the muscle exhibited reduced myofiber diameter, whereas ActRIIB-Fc treatment fully prevented this event (Figure 2D–F). Collectively, our results indicate that chronically injured liver negatively communicate with skeletal muscle independent of gender, and TGFβ family ligands neutralized by ActRIIB-Fc appear to mediate this negative crosstalk.
3.4. Neutralization of ActRIIB Ligands Reverses Chronic Liver Injury-Induced Muscle Atrophy
To evaluate whether ActRIIB ligand inhibition can reverse atrophied muscle after chronic liver injury, we first dosed female mice twice weekly with CCl4 for six weeks. Following that, mice treated with either ActRIIB-Fc or IgG were dosed weekly for 3 additional weeks with continued CCl4 injections (Figure 3A). Consequently, chronic liver injury caused muscle mass loss, which was reversed by ActRIIB-Fc (Figure 3B,C). The results further demonstrate that ActRIIB ligands mediate the negative crosstalk of injured liver to muscle.
3.5. Injured Liver Produces Gdf8 in Both Humans and Mice
Gdf8 is a myokine expressed in muscle that potently suppresses muscle mass and myogenesis in an autocrine/paracrine manner. It has high affinity to ActRIIB, and this receptor–ligand interaction inhibits proliferation and activation of satellite cells and myoblasts via downregulation of Pax7 and Myod1 expression in muscle [36,37,38,39]. Recent studies have hypothesized a role of Gdf8 in liver disease with concomitant sarcopenia, and it is associated with poor survival outcomes in cirrhosis patients [40,41]. Moreover, Gdf8 has been shown to regulate the fibrogenic phenotype of hepatic stellate cells in vitro, implying a role in hepatic injury response [42,43]. Thus, we hypothesized that Gdf8 may transduce negative signaling from injured liver to muscle. To test this, we performed the following studies.
We first assayed human cirrhotic liver samples and observed abundant Gdf8 protein expression, whereas it was virtually undetectable in healthy liver (Figure 4A). In male mice, within 48 h after a single dose of CCl4, acutely damaged liver produced increased Gdf8 protein along with elevated circulating Gdf8 (Figure 4B,C). No significant changes in muscle Gdf8 protein were observed (Figure 4D). Immunohistochemistry detected abundant Gdf8 protein in pericentral (Zone 3) hepatocytes in injured liver (Figure 4E). This finding is consistent with the well-characterized function for zone 3 hepatocytes in processing xenobiotics and explains why fibrosis manifests pericentrally in the CCl4 model. This indicates that hepatocytes metabolizing CCl4 produce Gdf8 protein, representing a novel source of this TGFβ ligand in a pathological condition.
To ascertain how muscle directly responds to liver injury, primary mouse hepatocytes were damaged via exposure to 0.5% CCl4 in culture medium. The concentration was determined as the minimal concentration to cause maximal release of liver enzymes (Figure 5A,B). ActRIIB-Fc soluble decoy receptor or Gdf8 antibody was added to differentiating C2C12 myotubes along with diluted injured hepatocyte or control medium with an equivalent CCl4 concentration. Culture medium from insulted hepatocytes but not medium from healthy hepatocytes inhibited myotube formation (myogenesis), as assessed by the reduced myotube diameter. The addition of an ActRIIB-Fc or a Gdf8 antibody fully rescued myotube formation potential (Figure 5C,D). This observation suggests that injured hepatocytes release ActRIIB-binding ligand(s), mainly Gdf8, which may exert direct adverse effects on muscle.
3.6. Neutralization of Gdf8 Prevents Chronic Liver Injury and Concomitant Muscle Mass Loss
Gdf8 appeared to mediate injured liver–muscle crosstalk, and thus Gdf8 neutralization may lead to the maintenance of muscle mass in the context of chronic liver injury. To test this, male mice were treated weekly with either IgG, Gdf8-Ab, or ActRIIB-Fc and administered CCl4 twice weekly for 6 weeks (Figure 6A). Consequently, we found that Gdf8 antibody and ActRIIB-Fc completely and equivalently protected against CCl4-induced skeletal muscle mass loss (Figure 6B). Surprisingly, we observed that Gdf8 neutralization also reduced hepatic collagen deposition (Figure 6C,D) and circulating bilirubin levels (Figure 6E) to a similar extent as ActRIIB-Fc treatment. Furthermore, hepatic Gdf8 protein content was increased with chronic CCl4 injury and reduced with Gdf8-Ab and ActRIIB-Fc treatment (Figure 6F). These results together demonstrate that Gdf8 neutralization inhibits liver fibrogenesis, improves liver function, and simultaneously prevents concomitant muscle atrophy.
3.7. Neutralization of Gdf8 Reverses Chronic Liver Injury and Concomitant Muscle Mass Loss
To assess the therapeutic potential of Gdf8 neutralization in an existing state of liver disease, we injured male mice twice weekly with CCl4 for a period of 11 weeks. Starting from the eighth week, mice received either anti-Gdf8 Ab, ActRIIB-Fc, or IgG once per week (Figure 7A). Compared to IgG controls, anti-Gdf8 therapy largely recovered the lost lean mass (Figure 7B), gastrocnemius mass (Figure 7C), and myofiber cross-sectional area (Figure 7D) observed in the CCl4-injured animals, similarly to ActRIIB-Fc treatment. Simultaneously, both ActRIIB-Fc and anti-Gdf8 antibody treatments significantly improved liver injury and function, manifested by reduced circulating liver enzymes ALT (Figure 7E) and AST (Figure 7F) and total bilirubin (Figure 6G). Concordantly, Gdf8 neutralization reduced hepatic collagen deposition as measured by picrosirius red staining, again, similarly to ActRIIB-Fc (Figure 7H,I). Collectively, these results demonstrate that hepatic Gdf8 is largely responsible for transducing adverse signaling effects of injured liver to skeletal muscle, and that Gdf8 promotes hepatic fibrotic response to chronic liver injury.
To determine whether Gdf8 exerts a direct effect on hepatic stellate cells, the major contributor to liver fibrogenesis, we exposed human hepatic stellate (LX-2) cells to exogenous Gdf8 at a concentration of 100 ng/mL [42]. We found that Gdf8 induced a gene expression signature indicative of hepatic stellate cell activation manifested by decreased expression of Hgf and increased expression of Fn14, Ctgf, and Tgfβ1 (Figure 8A). Remarkably, Gdf8 protein stimulated morphological changes in LX-2 cells, redolent of a septa-like structure commonly observed in liver fibrosis (Figure 8B). These results suggest that increased Gdf8 following liver injury may directly act on hepatic stellate cells to promote liver fibrogenesis.
3.8. Liver Injury Negatively Affects Muscle Regeneration, Which Is Prohibited by Neutralization of ActRIIB Ligands
Due to the finding that liver injury causes myogenic satellite cell marker suppression (Figure 1A), we queried whether liver injury would adversely affect muscle repair (myogenesis) and, if so, whether TGFβ family members also mediate this effect. To answer this question, we treated female mice with CCl4 or corn oil every three days for a period of 10 days. A single dose of cardiotoxin (CTX) was injected into the gastrocnemius muscle 6 h after the first CCl4 administration to induce focal muscle injury and repair [29]. IgG or ActRIIB-Fc was dosed prior to CTX treatment. As a result, 10 days after CTX injection, simultaneous liver and muscle injury led to skeletal muscle calcification (Figure 9A) and fibrosis (Figure 9B). We observed blunted regeneration in the removal of the necrotic sarcoplasm, followed by concurrent regeneration within the myotube membrane around the necrotic cytoplasm (Figure 9A). In addition, prominent collagen deposition and replacement of skeletal muscle tissue with non-muscle cells (Figure 9B) and significant reduction in size for nascent fibers (Figure 9C,D) were found with concomitant liver and muscle injury. ActRIIB-Fc treatment prevented these defects in muscle repair (Figure 9A–D). Taken together, this demonstrate that chronic liver injury results in detrimental effects on muscle repair after focal injury, strongly suggesting a role for hepatic dysfunction to inhibit muscle satellite cell activation and myogenesis. Furthermore, these effects were blunted by ActRIIB-Fc-mediated inhibition of TGFβ members, suggesting that extracellular ligand signaling through ActRIIB mediates the liver–muscle crosstalk in this context.
3.9. Exogenous Gdf8 Disrupts Muscle Regeneration, Mimicking Liver Injury
Our in vitro study shows that, like ActRIIB-Fc, Gdf8 inhibiting antibody recovered the diameter of myotubes exposed to injured hepatocyte media (Figure 5C,D). This finding suggests that liver injury-derived changes in systemic Gdf8 could mediate the negative effect of injured liver on skeletal muscle repair. To demonstrate this, muscle injury was induced via CTX and, 2 h later, 5 µg of Gdf8 protein or BSA were injected directly into the muscle. The next day, 1 µg of Gdf8 protein was administered to the injury site. Doses were chosen to mimic the injury-mediated waning in ligand exposure, as the protein is quickly absorbed and degraded. Ten days post CTX injury, muscles treated with Gdf8 displayed a decrease in regenerating myofiber diameter compared to the BSA control group (Figure 10A,B), reflecting earlier observations with CCl4. This result strongly suggests that Gdf8 contributes to liver injury-induced disruption of myofiber regeneration.
These studies demonstrate that ActRIIB-Fc intervention prevents and reverses liver injury-induced muscle atrophy and regenerative capabilities. We show that injured liver produces significant Gdf8 in both humans and mice. Furthermore, neutralization of Gdf8 largely recapitulates the positive muscle effects of ActRIIB-Fc intervention. Thus, these findings allow us to propose that ActRIIB ligands, primarily Gdf8, are significant mediators transducing the adverse signaling effects of injured liver to muscle. We and others have previously demonstrated that activin A induces muscle atrophy and degeneration [44]. However, in the context of CCl4-induced liver injury, we did not observe overt effects of neutralization of activin A alone on muscle atrophy. These finding suggests that activin A may not contribute to the injured liver–muscle crosstalk in our preclinical model.
Previous studies by others have suggested several potential mediators connecting liver injury and muscle wasting, including hyperammonemia, insufficiency of growth hormone, and testosterone [40,45,46,47,48,49,50]. Liver dysfunction and portosystemic shunting causes impaired ureagenesis and thus hyperammonemia, a consistent metabolic complication in cirrhotic patients [51,52]. A NF-kB/myostatin pathway in muscle cells has been proposed to contribute to ammonium acetate-induced muscle degeneration [40]. In addition, growth hormone was shown to inhibit the expression of myostatin in skeletal muscle [53]. These findings propound Gdf8 as a signal mediator downstream of those potential mechanisms. Indeed, cirrhotic patients exhibit increased Gdf8 expression in both skeletal muscle and plasma [54,55]. Gdf8 is considered an autocrine/paracrine myokine and a potent suppressor of myogenesis [38,39]. Here, we demonstrate for the first time that liver injury/fibrosis drives the production of hepatic Gdf8 to negatively regulate skeletal muscle mass. We thereby propose Gdf8 as a candidate hepato-myokine during liver injury. The data prompt the question as to whether Gdf8/ActRIIB ligand signaling is a key central mechanism by which the liver communicates with muscle more broadly in hepatic injury and disease, including NASH and cirrhosis.
One intriguing finding from our studies identified a circulatory environment that impedes muscle injury and repair. Our pathological assessment of the combination of liver and muscle injuries unveiled a discovery very redolent of a myopathy where the presence of Ringbinden fibers exists (Figure 9A). Ringbinden fibers have been described as an aberrant form of myofibrils that encircle themselves around existing or even dead fibers. Ringbinden fibers have been found predominantly in fast-twitch fibers in mice that possess a mutation in the skeletal muscle α-actin gene (Acta1) [56], which would be consistent in our model, wherein CCl4-induced liver injury selectively affects glycolytic fibers and is reinforced by Gdf8′s ability to affect glycolytic composition and type II fibers.
Previously published findings have demonstrated muscle dysfunction in CCl4-induced liver injury. Specifically, Weber et al. showed an increase in muscle protein catabolism of rat hindlimb muscles in a manner dependent on the severity of liver injury in CCl4-injured animals [57]. In that study, the muscle effects of CCl4 were limited to fast glycolytic fibers and did not reflect a more diffuse toxic effect in the muscle even in the context of 10-fold increases in CCl4 exposure on the muscle due to phenobarbital administration. Additionally, myotoxin-induced damage and degeneration can be characterized by specific histopathological features consistent with myofiber damage and immune cell infiltration [58], and this was not observed in our CCl4 studies or in other publications to our knowledge. These findings reported by Weber et al. are in line with more recent published data by Giusto et al. showing increased muscle protein degradation as the mechanism responsible for muscle atrophy observed in a CCl4-induced liver injury model [33]. Giusto et al. also reported increases in muscle protein degradation following several weeks of repeated CCl4 administration attributable to NF-KB-mediated upregulation of the ubiquitin ligase trim63, which is known to be regulated in muscle by myostatin [59]. Importantly, Giusto et al. also showed reductions in muscle myostatin (Gdf8) protein content following several weeks of CCl4-induced liver injury [33]. Although this may seem initially contradictory to reported clinical reports of increased circulating myostatin levels in liver disease patients who present with muscle atrophy [40,41], our data suggest that muscle may not be the sole source of increased circulating Gdf8 observed in these patients. Ultimately, although we cannot exclude the direct effects of CCl4 on damaging muscle, we do conclude that muscle Gdf8 expression is not significantly increased following hepatotoxic injury with CCl4, and the observed increased circulating levels are primarily attributable to extra-skeletal muscle sources, i.e., liver and potentially other organs.
The mechanisms by which Gdf8 contributes to the pathogenesis of liver damage are not entirely clear at this time, but multiple studies have shown that Gdf8 functions as a profibrotic factor in various tissues [41,42,43,60,61,62]. Our data have corroborated this role in the liver by demonstrating the ability of Gdf8-Ab to reduce CCl4-induced liver fibrosis. We postulate that, in addition to the endocrine role in inducing muscle atrophy, Gdf8 may play an additional autocrine role by stimulating fibrosis. This was highlighted by the morphological and gene expression changes engendered by Gdf8 that are indicative of stellate cell activation in vitro, which revealed Gdf8 as a novel and pivotal mediator, which potentially originates from damaged hepatocytes, underlying the potential crosstalk between hepatocytes and HSCs in damaged liver. Other reports show that Gdf8 induces the expression of procollagen type 1, TGFβ1, and the tissue inhibitor of metalloproteinase-1 in LX-2 cells and stimulate the release of procollagen type I from these cells [42], in line with our findings.
Our studies demonstrate inhibition of multiple members of the TGFβ superfamily as a promising therapeutic strategy to treat both primary- and secondary-organ injury or multi-organ injury. Here we demonstrate that targeting ActRIIB ligands, especially Gdf8, generates dual beneficial effects on both injured liver and wasted muscle. Our findings highlight that in scenarios where liver injury exists, it is vital to not only provide therapies that protect the liver and augment its repair but also address the resultant muscle atrophy that is a consequence of factors released into the systemic circulation shortly after the liver insult. Here we focused our efforts mainly on evaluating the effects of ActRIIB ligands on muscle atrophy secondary to liver injury. We also revealed that ActRIIB ligands may modulate liver fibrogenesis by directly targeting HSCs.
4. Conclusions
Based on our findings, we identified a working hypothesis to provide a substrate for future investigations to define potential new therapies (Figure 11). Recently, cirrhotic patients and the coinciding muscle wasting have been well documented [4,5,6]. Here we revealed that livers in patients with ESLD also produce abundant Gdf8. Furthermore, we identified specific hepatic cell types in NASH patient liver biopsies that may drive increased Gdf8 expression in fibrotic tissue. It is worth noting that we cannot definitively confirm that increases in Gdf8 observed in those patients is solely attributed to hepatic expression. Indeed, the increases observed in our studies are still relatively low compared to basal Gdf8 expression in muscle, and there are likely additional tissues involved that warrant further investigation. Nonetheless, these findings demonstrate the future potential of therapeutics targeting TGFβ family members for the treatment and control of liver diseases associated with muscle atrophy. Indeed, clinical evidence already exists for this pathway regulating skeletal muscle via ActRIIB-Fc treatment in the form of ACE-031, which has been shown to increase total lean body mass and thigh muscle volume in patients with muscular dystrophy [16]. Furthermore, antibody therapy directed against ActRIIB (BYM338-Bimagrumab) also demonstrates the ability to increase lean body mass in the clinical setting [63]. Recent BYM388 data in type II diabetic and obese patients demonstrated efficacy to improve outcomes associated not only with glucose control but also with hepatic fat fraction reductions, a finding highly relevant to NAFLD and NASH pathologies [64]. Results from these clinical studies may provide information pertinent to the utility of such therapeutics in liver disease.
Conceptualization, G.D., B.C.Y. and N.P.C.; methodology, A.C., M.H. and Y.W.; software, A.C. and M.H.; validation, G.D. and B.C.Y.; formal analysis, A.C., M.H. and Y.W.; investigation, A.C., M.H., Y.W., H.J. and J.Y.; resources, E.W., S.G., R.K.V. and N.P.C.; data curation, A.C.; writing—original draft preparation, B.C.Y., G.D. and A.C.; writing—review and editing, G.D. and B.C.Y.; visualization, G.D. and B.C.Y.; supervision, G.D. and B.C.Y.; project administration, G.D. and B.C.Y.; funding acquisition, G.D. and B.C.Y. All authors have read and agreed to the published version of the manuscript.
All mouse experiments were performed with the approval of the Institutional Animal Care and Use Committee of Indiana University—Purdue University Indianapolis (IUPUI). All human samples were collected under IUPUI Institutional Review Board-approved protocols (IUPUI IRB: EX0904-11).
Not applicable.
The data presented in this study are available on request from the corresponding authors.
The authors declare no conflict of interest.
Footnotes
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Abstract
Patients with end-stage liver disease exhibit progressive skeletal muscle atrophy, highlighting a negative crosstalk between the injured liver and muscle. Our study was to determine whether TGFβ ligands function as the mediators. Acute or chronic liver injury was induced by a single or repeated administration of carbon tetrachloride. Skeletal muscle injury and repair was induced by intramuscular injection of cardiotoxin. Activin type IIB receptor (ActRIIB) ligands and growth differentiation factor 8 (Gdf8) were neutralized with ActRIIB-Fc fusion protein and a Gdf8-specific antibody, respectively. We found that acute hepatic injury induced rapid and adverse responses in muscle, which was blunted by neutralizing ActRIIB ligands. Chronic liver injury caused muscle atrophy and repair defects, which were prevented or reversed by inactivating ActRIIB ligands. Furthermore, we found that pericentral hepatocytes produce excessive Gdf8 in injured mouse liver and cirrhotic human liver. Specific inactivation of Gdf8 prevented liver injury-induced muscle atrophy, similar to neutralization of ActRIIB ligands. Inhibition of Gdf8 also reversed muscle atrophy in a treatment paradigm following chronic liver injury. Direct injection of exogenous Gdf8 protein into muscle along with acute focal muscle injury recapitulated similar dysregulated muscle regeneration as that observed with liver injury. The results indicate that injured liver negatively communicate with the muscle largely via Gdf8. Unexpectedly, inactivation of Gdf8 simultaneously ameliorated liver fibrosis in mice following chronic liver injury. In vitro, Gdf8 induced human hepatic stellate (LX-2) cells to form a septa-like structure and stimulated expression of profibrotic factors. Our findings identified Gdf8 as a novel hepatomyokine contributing to injured liver–muscle negative crosstalk along with liver injury progression.
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; Chalasani, Naga P 3 ; Dai, Guoli 1 ; Yaden, Benjamin C 1 1 Department of Biology, School of Science, Center for Developmental and Regenerative Biology, Indiana University-Purdue University Indianapolis, Indianapolis, IN 46202, USA;
2 Department of Biological Sciences, College of Science, Purdue University, West Lafayette, IN 46202, USA;
3 Division of Gastroenterology and Hepatology, School of Medicine, Indiana University, Indianapolis, IN 46202, USA;