Skip to main content
NCBI home page
Physiology logoLink to Physiology
. 2019 Jun 5;34(4):232–239. doi: 10.1152/physiol.00003.2019

Skeletal Muscle Atrophy: Discovery of Mechanisms and Potential Therapies

Scott M Ebert 1,2, Asma Al-Zougbi 1, Sue C Bodine 1,2, Christopher M Adams 1,2,3,✉
PMCID: PMC6863373  PMID: 31165685

Abstract

Skeletal muscle atrophy proceeds through a complex molecular signaling network that is just beginning to be understood. Here, we discuss examples of recently identified molecular mechanisms of muscle atrophy and how they highlight an immense need and opportunity for focused biochemical investigations and further unbiased discovery work.

Introduction

Malnutrition, muscle disuse, aging, and essentially any serious illness or injury cause adult skeletal muscle fibers to become atrophic, ultimately leading to a loss of muscle mass. This disease process, called skeletal muscle atrophy, is highly prevalent, especially in older adults, and can significantly impair both health and quality of life, causing weakness and fatigability, reduced activity, derangements in whole-body metabolism, delayed recovery from acute illness and injury, increased morbidity and mortality from chronic disease, falls, extended hospital stays, and loss of independent living.

Despite its broad clinical impact, skeletal muscle atrophy is a relatively understudied and under-developed area of biomedical research. For example, muscle atrophy still lacks a pharmacological therapy, even though it is clear that therapies are desperately needed. Skeletal muscle atrophy also remains largely unexplored at the molecular level; indeed, unbiased genome- and proteome-wide analyses have revealed thousands of molecular changes in skeletal muscle that are strongly associated with muscle atrophy, but only a handful of these molecular events have been investigated at a mechanistic level.

Since 2001, molecular investigations of skeletal muscle atrophy have primarily centered around the E3 ubiquitin ligases MuRF1 (muscle RING finger 1) and MAFbx (muscle atrophy F-box, AKA atrogin-1), which are required for skeletal muscle atrophy during a wide range of stress conditions, including starvation, muscle disuse, glucocorticoid excess, and aging (8) (FIGURE 1). During those stress conditions, the glucocorticoid receptor and Foxo transcription factors activate the MuRF1 gene, and Foxo transcription factors activate the MAFbx gene, thereby increasing expression of MuRF1 and MAFbx proteins and promoting muscle atrophy. In the absence of stress conditions (i.e., in healthy young adult skeletal muscle), insulin/IGF-I signaling and a low level of glucocorticoids repress the MuRF1 and MAFbx genes by decreasing the expression and activity of Foxo transcription factors (44, 49, 50). In addition to its role in inhibiting muscle atrophy, insulin/IGF-I signaling also stimulates anabolic processes such as protein synthesis and skeletal muscle hypertrophy (10, 45). The key anabolic mediators of insulin/IGF-I signaling are Akt/PKB (protein kinase B), a protein kinase that directly inhibits Foxo transcription factors and contributes to mTORC1 activation, and mTORC1 (mechanistic target of rapamycin complex 1), a protein kinase that stimulates global protein synthesis and cell growth (29, 42). Resistance exercise and nutrient signals such as leucine also play important roles in stimulating the anabolic process in skeletal muscle.

FIGURE 1.

FIGURE 1.

Open in a new tab

A well-established pathway to skeletal muscle atrophy, involving MuRF1 and MAFbx

When muscles are deprived of nutrients, external loading, or neural activity, or when muscles are exposed to excess glucocorticoids or advanced age, expression of the the E3 ubiquitin ligase genes, MuRF1 and MAFbx, increases. Increased transcription of MuRF1 and MAFbx is controlled, in part, by activation of Foxo transcription factors and the glucocorticoid receptor (GR). Increased expression of MuRF1 and MAFbx promotes muscle atrophy via biochemical mechanisms that are not yet well defined. In healthy young adult skeletal muscle, activity of Foxo transcription factors is inhibited by insulin/IGF-I/Akt signaling and a low level of glucocorticoids. Insulin/IGF-I signaling has dual roles in that it can inhibit muscle atrophy through inhibition of Foxo transcription factors and stimulate protein synthesis and skeletal muscle hypertrophy via Akt and mTORC1.

The elegant and pioneering work surrounding the discovery of MuRF1 and MAFbx is clearly important to understanding how muscle atrophy occurs at the molecular level. However, it is also becoming increasingly clear that this well-defined signaling module is actually part of a much larger and more complex signaling network that controls skeletal muscle mass in mammals. In this review, we will briefly discuss examples of work we are pursuing to search for novel molecular mechanisms of muscle atrophy and new therapeutic approaches.

Discovery of a Different Molecular Signaling Pathway to Skeletal Muscle Atrophy

Our exploration into alternative potential mechanisms of muscle atrophy began with ATF4 (activating transcription factor 4), a rate-limiting subunit of several different heterodimeric basic leucine zipper (bZIP) transcription factors (4, 40). The biological effects of ATF4 are complex and highly context-dependent. Most of our current knowledge comes from work performed in transformed cultured cell lines, where ATF4 participates in anti-anabolic cellular stress responses as a downstream mediator of eIF2alpha kinases (4, 40), and ATF4 also independently participates in the anabolic response to insulin/IGF-I signaling as a downstream mediator of mTORC1 (2, 6, 38). Thus work in cultured cell models suggested that ATF4 could potentially have either a negative (anti-anabolic) or a positive (anabolic) effect on skeletal muscle mass. In skeletal muscle, the effect of ATF4 was unknown, but unbiased microarray analyses described an association between muscle atrophy and an increase in the level of ATF4 mRNA during a variety of conditions that cause muscle atrophy (e.g., starvation, cancer, renal failure, Type 1 diabetes, and muscle disuse) (43). Furthermore, increases in the level of ATF4 mRNA occurred alongside increases in MuRF1, MAFbx, and many other atrophy-associated mRNAs.

To determine the effect of ATF4 in skeletal muscle, we used molecular genetic methods to specifically manipulate the level of ATF4 in mouse skeletal muscle fibers in vivo. We found that ATF4 expression in skeletal muscle fibers is non-essential for skeletal muscle development and the maintenance of adult skeletal muscle mass and function (15, 16, 20). Interestingly, however, inhibition of ATF4 expression in young adult muscle fibers is partially protective against muscle atrophy during fasting and limb immobilization (16, 17, 20). In addition, mice lacking ATF4 in skeletal muscle fibers maintain young adult levels of muscle strength, quality, and mass into old age, elucidating ATF4 as the first example of a skeletal muscle protein that is required for the loss of strength, muscle quality, and muscle mass during aging in mammals (15). Conversely, forced expression of ATF4 in skeletal muscle fibers potently induces muscle fiber atrophy, even in the absence of aging or an acute stress condition (15–17, 20). Thus, in skeletal muscle fibers, ATF4 is not required for growth or maintenance, but it is both necessary and sufficient for atrophy during fasting, limb immobilization, and aging.

Although ATF4 is a transcriptional activator, ATF4-mediated muscle atrophy is not associated with increased expression of either MuRF1 or MAFbx mRNAs (16, 17). This finding suggested that ATF4 promotes muscle atrophy by activating genes that encode novel mediators of muscle atrophy. To find those genes, we performed an unbiased genome-wide search for mRNA transcripts that are increased by ATF4 overexpression and decreased by ATF4 gene excision in mouse skeletal muscle fibers. We identified Gadd45a (Growth arrest and DNA damage inducible 45 alpha) and p21/Cdkn1a (Cyclin-dependent kinase 1a) as two mRNAs that meet those criteria (16, 17, 20). Consistent with that finding, the Gadd45a and p21 genes are direct ATF4 targets (26, 28). Moreover, Gadd45a and p21 are among the most highly induced skeletal muscle mRNAs during skeletal muscle atrophy in humans, mice, and other mammalian species (5, 9, 16–18, 20, 23–25, 35, 37, 47, 52, 53).

The functional consequences of Gadd45a and p21 in skeletal muscle were unknown. Thus, to determine whether Gadd45a and p21 might play a causal role in muscle atrophy, we specifically manipulated the level of these proteins in mouse skeletal muscle fibers in vivo. We found that increased expression of either Gadd45a or p21 within mouse muscle fibers is sufficient to induce muscle fiber atrophy and required for muscle fiber atrophy during stress conditions that require ATF4 (16, 20). Furthermore, because ATF4 lies upstream of Gadd45a and p21, ATF4 is not required for Gadd45a- or p21-mediated atrophy, but Gadd45a and p21 are required for ATF4-mediated atrophy (16, 20). Thus ATF4 promotes muscle atrophy at least in part by activating the Gadd45a and p21 genes (FIGURE 2).

FIGURE 2.

FIGURE 2.

Open in a new tab

A different and more recently identified pathway to skeletal muscle atrophy, involving ATF4, Gadd45a, and p21

During fasting, immobilization, and aging, an ATF4 heterodimer activates the Gadd45a and p21 genes within skeletal muscle fibers, leading to increased expression of Gadd45a and p21 proteins. Gadd45a is a small globular protein that specifically interacts with at least 67 skeletal muscle proteins, including 11 protein kinases and 3 protein tyrosine phosphatases. To date, one Gadd45a-interacting protein has been studied in detail: the MAP kinase kinase kinase MEKK4, which is allosterically activated by Gadd45a binding. The Gadd45a/MEKK4 complex activates four MAP kinase kinases (MKK3, 4, 6, and 7), leading to activation of p38 MAP kinase; this signaling cascade promotes muscle atrophy via biochemical mechanisms that are not yet understood. p21 is a small, intrinsically disordered protein that promotes muscle atrophy by an unknown biochemical mechanism. One important downstream effect of p21 is repression of the spermine oxidase (Smox) gene, which encodes an enzyme that protects against muscle atrophy. Additional regulators of the Gadd45a gene include histone deacetylase 4 (HDAC4; during muscle denervation) and perhaps Foxo transcription factors (during fasting); additional regulators of the p21 gene include p53 (during immobilization).

Subsequent studies have revealed that ATF4 is not the only transcriptional regulator that controls the Gadd45a and p21 genes during skeletal muscle atrophy. For example, Gadd45a is required for denervation-induced atrophy (16), and, in that context, histone deacetylase 4 (rather than ATF4) is responsible for increasing Gadd45a expression (11). There is also some evidence that Foxo transcription factors might contribute to Gadd45a gene expression during fasting (39). Finally, during muscle immobilization, ATF4 and p53 independently and additively increase p21 expression (20). These findings indicate that the Gadd45a and p21 genes are important convergence points for multiple transcriptional regulators that promote muscle atrophy (FIGURE 2).

Consistent with their roles as downstream mediators of the ATF4 pathway, Gadd45a and p21 promote muscle atrophy without increasing expression of either MuRF1 mRNA or MAFbx mRNA (12, 16). These findings provide strong evidence that the ATF4 pathway does not lie upstream of MuRF1 or MAFbx but rather must operate in parallel to MuRF1 and MAFbx or lie downstream of MuRF1 and MAFbx. Resolving the relationship between these two pathways is an important area that we are now pursuing. To date, these pathways have been investigated and considered in isolation not because they exist in isolation but simply because the connections are not yet known.

To begin to understand how Gadd45a (an 18-kDa globular protein) promotes muscle atrophy, we biochemically isolated and identified skeletal muscle proteins that interact with Gadd45a as it induces atrophy in mouse skeletal muscle fibers in vivo (13). We found that Gadd45a interacts with (i.e., specifically binds) ~3% of mammalian protein kinases (MEKK4, MKK3, MKK6, MKK7, Raf-1, A-Raf, JAK1, ILK, RSK2, SPEG, MSK1, MSK2, RIOK3, and RIPK3) and 3% of mammalian protein tyrosine phosphatases (ACP1, PTP-1B, and SHP-1), as well as 53 other low-abundance skeletal muscle proteins (13). All of these proteins interact with Gadd45a in skeletal muscle, but none of these proteins interact with ATF4 or p21 in skeletal muscle (Ebert SM, Adams CM, unpublished observations). Thus far, we have found that at least one of these Gadd45a-interacting protein kinases (MEKK4) is directly activated by Gadd45a binding in skeletal muscle fibers (13) (FIGURE 2).

MEKK4 (MAPK/ERK kinase kinase 4, also known as MAP3K4) is a member of the MAP kinase kinase kinase family. In healthy young adult skeletal muscle, MEKK4 resides in an inactive conformation, with its NH2-terminal autoinhibitory domain occluding its COOH-terminal kinase domain (13). However, in the presence of conditions that cause muscle atrophy, the Gadd45a gene becomes active, Gadd45a levels rise, and Gadd45a binds the NH2-terminal region of MEKK4, generating a conformational change in MEKK4 that relieves auto-inhibition of the COOH-terminal kinase domain (13). Thus the active protein kinase is actually a complex of two proteins, Gadd45a and MEKK4. Importantly, in mouse skeletal muscle fibers, the Gadd45a/MEKK4 kinase complex induces atrophy and is at least partially required for atrophy during Gadd45a overexpression and limb immobilization (13). This identified a direct biochemical mechanism by which Gadd45a induces muscle atrophy.

Based on these results, a newly emergent question is how does the Gadd45a/MEKK4 kinase complex promote skeletal muscle atrophy? As it induces muscle fiber atrophy, the Gadd45a/MEKK4 complex activates specific downstream MAP kinase kinases (MKK3, MKK4, MKK6, and MKK7), which in turn activate a downstream MAP kinase, p38 (13). Thus the Gadd45a/MEKK4 complex may induce muscle fiber atrophy via one or more of these downstream kinases. In potential support of that hypothesis, lifelong, constitutive activation of MKK6 within skeletal muscle fibers dramatically reduces skeletal muscle mass and integrity in a p38alpha-dependent manner (54). Furthermore, the capacity of Gadd45a to allosterically activate MEKK4 suggests that Gadd45a may allosterically regulate other Gadd45a-interacting kinases and phosphatases that we identified in skeletal muscle fibers (a substantial portion of the signal transduction network within skeletal muscle fibers). These investigations are ongoing.

At this point, the direct biochemical mechanism by which p21 promotes muscle atrophy is not yet known. Thus, using approaches similar to those used for Gadd45a, we are now searching for direct p21 targets in adult skeletal muscle fibers. From work in other cell types, we know that p21 is a tightly regulated, intrinsically disordered protein capable of binding and regulating a wide range of other proteins, leading to a wide range of cellular effects (1, 7). p21 is best known as a cyclin-dependent kinase (CDK) inhibitor, based on its capacity to bind and inhibit a subset of cyclin/CDK complexes (1, 7). Although cyclin/CDK complexes play critical roles in cell cycle progression, it is unlikely that p21 causes muscle atrophy by inhibiting the cell cycle (20). First, during muscle atrophy, p21 specifically increases in terminally differentiated skeletal muscle fibers (which are post-mitotic) but not in satellite cells or interstitial cells (which have mitotic potential) (27). Second, the interventions used in our studies specifically targeted terminally differentiated skeletal muscle fibers (20). These considerations suggest that p21 promotes skeletal muscle atrophy by binding and regulating one or more proteins within skeletal muscle fibers (which may include classical and/or novel p21-binding proteins), leading to cell cycle-independent cellular changes, and ultimately muscle fiber atrophy.

Since p21 can regulate gene expression via direct interactions with transcription regulatory proteins (1, 7), we performed an unbiased genome-wide analysis to search for mRNA transcripts that are regulated by p21 in mouse skeletal muscle fibers. Interestingly, we found that p21 strongly represses the mRNA encoding spermine oxidase (Smox), leading to a reduction in Smox protein (12). In addition, p21-dependent atrophy conditions (aging, fasting, and immobilization) strongly repress Smox mRNA and protein in skeletal muscle (12), and p21 is required for this repression (12).

Smox is an enzyme in polyamine metabolism that catabolizes spermine to spermidine, 3-aminopropanol, and hydrogen peroxide; it was not known to play a role in the control of muscle mass. In further studies, we found that a reduction in Smox expression is sufficient to induce muscle fiber atrophy (12) and required for p21-mediated muscle fiber atrophy (12). Moreover, by a mechanism that is not yet understood, a reduction of Smox expression increases mRNAs that promote muscle atrophy and decreases mRNAs that help to maintain muscle mass (12). Thus, in healthy, non-stressed skeletal muscle, a low level of p21 permits Smox expression, which helps to maintain basal muscle gene expression and fiber size. Conversely, during aging and other conditions that cause muscle atrophy, p21 expression rises, leading to reduced Smox expression, disruption of basal muscle gene expression, and muscle fiber atrophy (12) (FIGURE 2).

The results illustrated in FIGURE 2 outline a previously unrecognized molecular pathway that leads to skeletal muscle atrophy during fasting, limb immobilization, aging, and possibly other conditions. That said, our knowledge of this pathway is still evolving, and new discoveries always stimulate new mechanistic questions. It is also crucially important to understand that this pathway, like the pathway illustrated in FIGURE 1, represents a small portion of the big picture, and the underlying mechanism for muscle wasting may differ depending on the etiology; for example, muscle atrophy induced during highly inflammatory states (e.g., sepsis, burns) may differ from that seen in disuse or aging. The emerging paradigm is that skeletal muscle atrophy is the output of a highly complex signal transduction network in skeletal muscle fibers. That network is not yet well defined but includes the pathways illustrated in FIGURES 1 AND 2, as well as other pathways, some of which remain to be discovered.

Discovery of Small Molecule Inhibitors of Skeletal Muscle Atrophy

As discussed above, skeletal muscle atrophy currently lacks a pharmacological therapy. One common approach to developing a therapy is to target specific vulnerabilities in the signal transduction network that promotes muscle atrophy. To date, molecular genetic investigations have identified several proteins that are at least partially required for some forms of skeletal muscle atrophy, including but not limited to MuRF1, MAFbx, Foxo transcription factors, ATF4, Gadd45a, p21, and MEKK4. Targeted removal or inhibition of any of these proteins within skeletal muscle fibers partially protects against muscle atrophy without promoting muscle hypertrophy (9, 13, 16, 17, 20, 41, 46). There are also other potential molecular targets, including, most prominently, myostatin, whose targeted inhibition reduces muscle atrophy and promotes muscle hypertrophy by mechanisms that involve both skeletal muscle fibers and satellite cells (34, 51).

As an alternative approach, we developed and utilized a novel strategy based on mRNA expression signatures, i.e., the specific patterns of positive and negative changes in cellular mRNA levels that are elicited by small molecules or disease conditions (3, 32). Because muscle atrophy involves numerous positive and negative changes in skeletal muscle mRNA expression, and because at least some of those changes are required for muscle atrophy, we hypothesized that we might discover small molecule inhibitors of muscle atrophy by identifying small molecules whose mRNA expression signatures negatively correlate to mRNA expression signatures of muscle atrophy (32). Unlike a conventional drug discovery approach, this strategy does not rely on a predefined molecular target or pathway but rather is designed to identify small molecules that counter the entire spectrum of pathological changes in atrophic muscle.

To employ this strategy, we obtained skeletal muscle samples from humans and mice under basal conditions and during two conditions that cause muscle atrophy (fasting and spinal cord injury). We then used genome-wide mRNA expression arrays to analyze the skeletal muscle samples and build two distinct mRNA expression signatures of muscle atrophy. Our first signature consisted of evolutionarily conserved mRNAs that are induced or repressed by fasting in both human and mouse skeletal muscle (32). Our second signature consisted of mRNAs that are induced or repressed by two very different types of muscle atrophy stimuli (fasting and spinal cord injury) in human skeletal muscle (32). By developing and utilizing these two distinct signatures of muscle atrophy, we sought to increase the specificity of our screen and the chance of discovering broad-spectrum inhibitors of muscle atrophy that would be applicable to multiple atrophy conditions in both humans and animals (3).

Next, to identify candidate small molecule inhibitors of muscle atrophy, we compared our two mRNA expression signatures of muscle atrophy to mRNA expression signatures of small molecules in the Connectivity Map library (33), looking for compounds with favorable safety characteristics whose mRNA expression signatures negatively correlate to both mRNA expression signatures of muscle atrophy. Through these analyses, we identified two structurally dissimilar natural compounds whose signatures in human cell lines negatively correlate to both muscle atrophy signatures (14, 32) (FIGURE 3). The first compound, ursolic acid, is a pentacyclic triterpene acid found in several edible herbs and fruits, including apples, which on average contain ~50 mg of ursolic acid within their peels (22). The second compound, tomatidine, is a hexacyclic alkaloid that naturally occurs as a metabolite from green (unripe) tomatoes. Both of these compounds possess favorable safety profiles (19, 21, 30, 36), but their effects on skeletal muscle were not known.

FIGURE 3.

FIGURE 3.

Open in a new tab

Use of mRNA expression signatures of skeletal muscle atrophy to identify small molecule inhibitors of skeletal muscle atrophy

Skeletal muscle biopsies obtained from human subjects were used to determine mRNA expression signatures of skeletal muscle atrophy. We then searched for small molecules whose mRNA expression signatures in human cell lines negatively correlate to mRNA signatures of skeletal muscle atrophy. This unbiased search revealed two small molecules, ursolic acid and tomatidine, which inhibit skeletal muscle atrophy.

Using mouse models, we found that ursolic acid and tomatidine reduce skeletal muscle atrophy in several different scenarios, including starvation, muscle disuse, and advanced age (14, 15, 32). Furthermore, in healthy, young adult animals lacking skeletal muscle atrophy, both ursolic acid and tomatidine stimulate skeletal muscle hypertrophy (14, 32). Importantly, the positive effects of ursolic acid and tomatidine on muscle mass are accompanied by increased strength and exercise capacity, as well as increased specific force (i.e., strength per unit muscle mass), indicating a greater effect on strength than muscle mass (14, 15, 31, 32).

Consistent with their phenotypic effects and the way they were discovered, ursolic acid and tomatidine generate widespread changes in skeletal muscle mRNA expression, including repression of ATF4-dependent gene expression (15). Furthermore, both compounds stimulate hypertrophy of cultured skeletal myotubes from both humans and mice, indicating that ursolic acid and tomatidine act directly on muscle cells and that their mechanisms of action are evolutionarily conserved (14, 32). The direct molecular receptors are not yet known; however, ursolic acid- and tomatidine-mediated skeletal muscle hypertrophy is associated with increased mTORC1 activity in skeletal muscle (14, 32). In addition, ursolic acid strongly upregulates Smox mRNA, enhances insulin-mediated activation of the insulin receptor, enhances IGF-I-mediated activation of the IGF-I receptor, increases Akt activity, decreases activity of Foxo transcription factors, and decreases expression of MuRF1 and MAFbx mRNAs (32).

Deeper investigations of ursolic acid and tomatidine could potentially generate important new insights into the pathogenesis of skeletal muscle atrophy. In addition, as natural dietary compounds, ursolic acid and tomatidine may have utility, first as new potential nutritional agents for muscle health and wellness, and, second, as lead compounds for muscle atrophy pharmaceuticals. In our view, preventing and treating muscle atrophy on a population-wide basis will almost certainly require a repertoire of modalities that can be used alone or in combination, depending on the clinical circumstances, similar to other complex chronic diseases such as osteoporosis, hypertension, Type 2 diabetes, and dyslipidemia. In the case of muscle atrophy, the modalities should include exercise, optimized nutrition, and multiple classes of pharmaceuticals that target a variety of atrophy pathways.

Summary

Skeletal muscle atrophy proceeds through a complex molecular signaling network that is just beginning to be elucidated. Since the initial discoveries of MuRF1 and MAFbx in 2001, several additional mediators of muscle atrophy have been discovered, providing new insights into how muscle atrophy occurs at the molecular level. However, these new discoveries have also generated many new questions and illustrate how far we remain from having a comprehensive, detailed, and fully integrated understanding of skeletal muscle atrophy. It is also vitally important to remember that the vast majority of molecular events that accompany muscle atrophy have yet to be investigated at a mechanistic level. This being the case, there are abundant opportunities for original mechanistic and therapeutic discoveries, and molecular investigations of skeletal muscle atrophy should remain an exciting area of research for many years into the future.

Acknowledgments

This work was supported by the National Institutes of Health Grants R01 AR-071762, R01 AG-060637, and R44 AG-047684; the U.S. Department of Veterans Affairs Grant I01BX00976; the Fraternal Order of Eagles Diabetes Research Center at the University of Iowa; and Emmyon, Inc.

C.M.A. is an inventor on patents related to ursolic acid and tomatidine, which have been licensed to Emmyon, Inc. C.M.A. and S.M.E. are officers of Emmyon, Inc. S.C.B. is a consultant to Emmyon, Inc. C.M.A., S.M.E., and S.C.B. hold equity in Emmyon, Inc.

S.M.E., A.A.-Z., S.C.B., and C.M.A. prepared figures; S.M.E., A.A.-Z., S.C.B., and C.M.A. drafted manuscript; S.M.E., A.A.-Z., S.C.B., and C.M.A. edited and revised manuscript; S.M.E., A.A.-Z., S.C.B., and C.M.A. approved final version of manuscript.

References

  • 1.Abbas T, Dutta A. p21 in cancer: intricate networks and multiple activities. Nat Rev Cancer 9: 400–414, 2009. doi: 10.1038/nrc2657. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Adams CM. Role of the transcription factor ATF4 in the anabolic actions of insulin and the anti-anabolic actions of glucocorticoids. J Biol Chem 282: 16744–16753, 2007. doi: 10.1074/jbc.M610510200. [DOI] [PubMed] [Google Scholar]
  • 3.Adams CM, Ebert SM, Dyle MC. Use of mRNA expression signatures to discover small molecule inhibitors of skeletal muscle atrophy. Curr Opin Clin Nutr Metab Care 18: 263–268, 2015. doi: 10.1097/MCO.0000000000000159. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Baird TD, Wek RC. Eukaryotic initiation factor 2 phosphorylation and translational control in metabolism. Adv Nutr 3: 307–321, 2012. doi: 10.3945/an.112.002113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Banduseela VC, Ochala J, Chen YW, Göransson H, Norman H, Radell P, Eriksson LI, Hoffman EP, Larsson L. Gene expression and muscle fiber function in a porcine ICU model. Physiol Genomics 39: 141–159, 2009. doi: 10.1152/physiolgenomics.00026.2009. [DOI] [PubMed] [Google Scholar]
  • 6.Ben-Sahra I, Hoxhaj G, Ricoult SJH, Asara JM, Manning BD. mTORC1 induces purine synthesis through control of the mitochondrial tetrahydrofolate cycle. Science 351: 728–733, 2016. doi: 10.1126/science.aad0489. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Besson A, Dowdy SF, Roberts JM. CDK inhibitors: cell cycle regulators and beyond. Dev Cell 14: 159–169, 2008. doi: 10.1016/j.devcel.2008.01.013. [DOI] [PubMed] [Google Scholar]
  • 8.Bodine SC, Baehr LM. Skeletal muscle atrophy and the E3 ubiquitin ligases MuRF1 and MAFbx/atrogin-1. Am J Physiol Endocrinol Metab 307: E469–E484, 2014. doi: 10.1152/ajpendo.00204.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA, Poueymirou WT, Panaro FJ, Na E, Dharmarajan K, Pan ZQ, Valenzuela DM, DeChiara TM, Stitt TN, Yancopoulos GD, Glass DJ. Identification of ubiquitin ligases required for skeletal muscle atrophy. Science 294: 1704–1708, 2001. doi: 10.1126/science.1065874. [DOI] [PubMed] [Google Scholar]
  • 10.Bodine SC, Stitt TN, Gonzalez M, Kline WO, Stover GL, Bauerlein R, Zlotchenko E, Scrimgeour A, Lawrence JC, Glass DJ, Yancopoulos GD. Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat Cell Biol 3: 1014–1019, 2001. doi: 10.1038/ncb1101-1014. [DOI] [PubMed] [Google Scholar]
  • 11.Bongers KS, Fox DK, Ebert SM, Kunkel SD, Dyle MC, Bullard SA, Dierdorff JM, Adams CM. Skeletal muscle denervation causes skeletal muscle atrophy through a pathway that involves both Gadd45a and HDAC4. Am J Physiol Endocrinol Metab 305: E907–E915, 2013. doi: 10.1152/ajpendo.00380.2013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Bongers KS, Fox DK, Kunkel SD, Stebounova LV, Murry DJ, Pufall MA, Ebert SM, Dyle MC, Bullard SA, Dierdorff JM, Adams CM. Spermine oxidase maintains basal skeletal muscle gene expression and fiber size and is strongly repressed by conditions that cause skeletal muscle atrophy. Am J Physiol Endocrinol Metab 308: E144–E158, 2015. doi: 10.1152/ajpendo.00472.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Bullard SA, Seo S, Schilling B, Dyle MC, Dierdorff JM, Ebert SM, DeLau AD, Gibson BW, Adams CM. Gadd45a protein promotes skeletal muscle atrophy by forming a complex with the protein kinase MEKK4. J Biol Chem 291: 17496–17509, 2016. doi: 10.1074/jbc.M116.740308. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Dyle MC, Ebert SM, Cook DP, Kunkel SD, Fox DK, Bongers KS, Bullard SA, Dierdorff JM, Adams CM. Systems-based discovery of tomatidine as a natural small molecule inhibitor of skeletal muscle atrophy. J Biol Chem 289: 14913–14924, 2014. doi: 10.1074/jbc.M114.556241. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Ebert SM, Dyle MC, Bullard SA, Dierdorff JM, Murry DJ, Fox DK, Bongers KS, Lira VA, Meyerholz DK, Talley JJ, Adams CM. Identification and small molecule inhibition of an activating transcription factor 4 (ATF4)-dependent pathway to age-related skeletal muscle weakness and atrophy. J Biol Chem 290: 25497–25511, 2015. doi: 10.1074/jbc.M115.681445. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Ebert SM, Dyle MC, Kunkel SD, Bullard SA, Bongers KS, Fox DK, Dierdorff JM, Foster ED, Adams CM. Stress-induced skeletal muscle Gadd45a expression reprograms myonuclei and causes muscle atrophy. J Biol Chem 287: 27290–27301, 2012. doi: 10.1074/jbc.M112.374777. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Ebert SM, Monteys AM, Fox DK, Bongers KS, Shields BE, Malmberg SE, Davidson BL, Suneja M, Adams CM. The transcription factor ATF4 promotes skeletal myofiber atrophy during fasting. Mol Endocrinol 24: 790–799, 2010. doi: 10.1210/me.2009-0345. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Edwards MG, Anderson RM, Yuan M, Kendziorski CM, Weindruch R, Prolla TA. Gene expression profiling of aging reveals activation of a p53-mediated transcriptional program. BMC Genomics 8: 80, 2007. doi: 10.1186/1471-2164-8-80. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Ferreira DS, Esperandim VR, Toldo MP, Saraiva J, Cunha WR, de Albuquerque S. Trypanocidal activity and acute toxicity assessment of triterpene acids. Parasitol Res 106: 985–989, 2010. doi: 10.1007/s00436-010-1740-2. [DOI] [PubMed] [Google Scholar]
  • 20.Fox DK, Ebert SM, Bongers KS, Dyle MC, Bullard SA, Dierdorff JM, Kunkel SD, Adams CM. p53 and ATF4 mediate distinct and additive pathways to skeletal muscle atrophy during limb immobilization. Am J Physiol Endocrinol Metab 307: E245–E261, 2014. doi: 10.1152/ajpendo.00010.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Friedman M. Anticarcinogenic, cardioprotective, and other health benefits of tomato compounds lycopene, α-tomatine, and tomatidine in pure form and in fresh and processed tomatoes. J Agric Food Chem 61: 9534–9550, 2013. doi: 10.1021/jf402654e. [DOI] [PubMed] [Google Scholar]
  • 22.Frighetto RTS, Welendorf RM, Nigro EN, Frighetto N, Siani AC. Isolation of ursolic acid from apple peels by high speed counter-current chromatography. Food Chem 106: 767–771, 2008. doi: 10.1016/j.foodchem.2007.06.003. [DOI] [Google Scholar]
  • 23.Gonzalez de Aguilar JL, Niederhauser-Wiederkehr C, Halter B, De Tapia M, Di Scala F, Demougin P, Dupuis L, Primig M, Meininger V, Loeffler JP. Gene profiling of skeletal muscle in an amyotrophic lateral sclerosis mouse model. Physiol Genomics 32: 207–218, 2008. doi: 10.1152/physiolgenomics.00017.2007. [DOI] [PubMed] [Google Scholar]
  • 24.Hulmi JJ, Silvennoinen M, Lehti M, Kivelä R, Kainulainen H. Altered REDD1, myostatin, and Akt/mTOR/FoxO/MAPK signaling in streptozotocin-induced diabetic muscle atrophy. Am J Physiol Endocrinol Metab 302: E307–E315, 2012. doi: 10.1152/ajpendo.00398.2011. [DOI] [PubMed] [Google Scholar]
  • 25.Ibebunjo C, Chick JM, Kendall T, Eash JK, Li C, Zhang Y, Vickers C, Wu Z, Clarke BA, Shi J, Cruz J, Fournier B, Brachat S, Gutzwiller S, Ma Q, Markovits J, Broome M, Steinkrauss M, Skuba E, Galarneau JR, Gygi SP, Glass DJ. Genomic and proteomic profiling reveals reduced mitochondrial function and disruption of the neuromuscular junction driving rat sarcopenia. Mol Cell Biol 33: 194–212, 2013. doi: 10.1128/MCB.01036-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Inoue Y, Kawachi S, Ohkubo T, Nagasaka M, Ito S, Fukuura K, Itoh Y, Ohoka N, Morishita D, Hayashi H. The CDK inhibitor p21 is a novel target gene of ATF4 and contributes to cell survival under ER stress. FEBS Lett 591: 3682–3691, 2017. doi: 10.1002/1873-3468.12869. [DOI] [PubMed] [Google Scholar]
  • 27.Ishido M, Kami K, Masuhara M. In vivo expression patterns of MyoD, p21, and Rb proteins in myonuclei and satellite cells of denervated rat skeletal muscle. Am J Physiol Cell Physiol 287: C484–C493, 2004. doi: 10.1152/ajpcell.00080.2004. [DOI] [PubMed] [Google Scholar]
  • 28.Jiang HY, Jiang L, Wek RC. The eukaryotic initiation factor-2 kinase pathway facilitates differential GADD45a expression in response to environmental stress. J Biol Chem 282: 3755–3765, 2007. doi: 10.1074/jbc.M606461200. [DOI] [PubMed] [Google Scholar]
  • 29.Kimball SR, Jefferson LS. Control of translation initiation through integration of signals generated by hormones, nutrients, and exercise. J Biol Chem 285: 29027–29032, 2010. doi: 10.1074/jbc.R110.137208. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Koh E, Kaffka S, Mitchell AE. A long-term comparison of the influence of organic and conventional crop management practices on the content of the glycoalkaloid α-tomatine in tomatoes. J Sci Food Agric 93: 1537–1542, 2013. doi: 10.1002/jsfa.5951. [DOI] [PubMed] [Google Scholar]
  • 31.Kunkel SD, Elmore CJ, Bongers KS, Ebert SM, Fox DK, Dyle MC, Bullard SA, Adams CM. Ursolic acid increases skeletal muscle and brown fat and decreases diet-induced obesity, glucose intolerance and fatty liver disease. PLoS One 7: e39332, 2012. doi: 10.1371/journal.pone.0039332. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Kunkel SD, Suneja M, Ebert SM, Bongers KS, Fox DK, Malmberg SE, Alipour F, Shields RK, Adams CM. mRNA expression signatures of human skeletal muscle atrophy identify a natural compound that increases muscle mass. Cell Metab 13: 627–638, 2011. doi: 10.1016/j.cmet.2011.03.020. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Lamb J, Crawford ED, Peck D, Modell JW, Blat IC, Wrobel MJ, Lerner J, Brunet JP, Subramanian A, Ross KN, Reich M, Hieronymus H, Wei G, Armstrong SA, Haggarty SJ, Clemons PA, Wei R, Carr SA, Lander ES, Golub TR. The Connectivity Map: using gene-expression signatures to connect small molecules, genes, and disease. Science 313: 1929–1935, 2006. doi: 10.1126/science.1132939. [DOI] [PubMed] [Google Scholar]
  • 34.Latres E, Pangilinan J, Miloscio L, Bauerlein R, Na E, Potocky TB, Huang Y, Eckersdorff M, Rafique A, Mastaitis J, Lin C, Murphy AJ, Yancopoulos GD, Gromada J, Stitt T. Myostatin blockade with a fully human monoclonal antibody induces muscle hypertrophy and reverses muscle atrophy in young and aged mice. Skelet Muscle 5: 34, 2015. doi: 10.1186/s13395-015-0060-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Laure L, Suel L, Roudaut C, Bourg N, Ouali A, Bartoli M, Richard I, Danièle N. Cardiac ankyrin repeat protein is a marker of skeletal muscle pathological remodelling. FEBS J 276: 669–684, 2009. doi: 10.1111/j.1742-4658.2008.06814.x. [DOI] [PubMed] [Google Scholar]
  • 36.Lee AW, Chen TL, Shih CM, Huang CY, Tsao NW, Chang NC, Chen YH, Fong TH, Lin FY. Ursolic acid induces allograft inflammatory factor-1 expression via a nitric oxide-related mechanism and increases neovascularization. J Agric Food Chem 58: 12941–12949, 2010. doi: 10.1021/jf103265x. [DOI] [PubMed] [Google Scholar]
  • 37.Llano-Diez M, Gustafson AM, Olsson C, Goransson H, Larsson L. Muscle wasting and the temporal gene expression pattern in a novel rat intensive care unit model. BMC Genomics 12: 602, 2011. doi: 10.1186/1471-2164-12-602. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Malmberg SE, Adams CM. Insulin signaling and the general amino acid control response. Two distinct pathways to amino acid synthesis and uptake. J Biol Chem 283: 19229–19234, 2008. doi: 10.1074/jbc.M801331200. [DOI] [PubMed] [Google Scholar]
  • 39.Milan G, Romanello V, Pescatore F, Armani A, Paik JH, Frasson L, Seydel A, Zhao J, Abraham R, Goldberg AL, Blaauw B, DePinho RA, Sandri M. Regulation of autophagy and the ubiquitin-proteasome system by the FoxO transcriptional network during muscle atrophy. Nat Commun 6: 6670, 2015. doi: 10.1038/ncomms7670. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Pakos-Zebrucka K, Koryga I, Mnich K, Ljujic M, Samali A, Gorman AM. The integrated stress response. EMBO Rep 17: 1374–1395, 2016. doi: 10.15252/embr.201642195. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Reed SA, Sandesara PB, Senf SM, Judge AR. Inhibition of FoxO transcriptional activity prevents muscle fiber atrophy during cachexia and induces hypertrophy. FASEB J 26: 987–1000, 2012. doi: 10.1096/fj.11-189977. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Rommel C, Bodine SC, Clarke BA, Rossman R, Nunez L, Stitt TN, Yancopoulos GD, Glass DJ. Mediation of IGF-1-induced skeletal myotube hypertrophy by PI(3)K/Akt/mTOR and PI(3)K/Akt/GSK3 pathways. Nat Cell Biol 3: 1009–1013, 2001. doi: 10.1038/ncb1101-1009. [DOI] [PubMed] [Google Scholar]
  • 43.Sacheck JM, Hyatt JP, Raffaello A, Jagoe RT, Roy RR, Edgerton VR, Lecker SH, Goldberg AL. Rapid disuse and denervation atrophy involve transcriptional changes similar to those of muscle wasting during systemic diseases. FASEB J 21: 140–155, 2007. doi: 10.1096/fj.06-6604com. [DOI] [PubMed] [Google Scholar]
  • 44.Sandri M, Sandri C, Gilbert A, Skurk C, Calabria E, Picard A, Walsh K, Schiaffino S, Lecker SH, Goldberg AL. Foxo transcription factors induce the atrophy-related ubiquitin ligase atrogin-1 and cause skeletal muscle atrophy. Cell 117: 399–412, 2004. doi: 10.1016/S0092-8674(04)00400-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Schiaffino S, Mammucari C. Regulation of skeletal muscle growth by the IGF1-Akt/PKB pathway: insights from genetic models. Skelet Muscle 1: 4, 2011. doi: 10.1186/2044-5040-1-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Senf SM, Dodd SL, Judge AR. FOXO signaling is required for disuse muscle atrophy and is directly regulated by Hsp70. Am J Physiol Cell Physiol 298: C38–C45, 2010. doi: 10.1152/ajpcell.00315.2009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Stevenson EJ, Giresi PG, Koncarevic A, Kandarian SC. Global analysis of gene expression patterns during disuse atrophy in rat skeletal muscle. J Physiol 551: 33–48, 2003. doi: 10.1113/jphysiol.2003.044701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Stitt TN, Drujan D, Clarke BA, Panaro F, Timofeyva Y, Kline WO, Gonzalez M, Yancopoulos GD, Glass DJ. The IGF-1/PI3K/Akt pathway prevents expression of muscle atrophy-induced ubiquitin ligases by inhibiting FOXO transcription factors. Mol Cell 14: 395–403, 2004. doi: 10.1016/S1097-2765(04)00211-4. [DOI] [PubMed] [Google Scholar]
  • 50.Waddell DS, Baehr LM, van den Brandt J, Johnsen SA, Reichardt HM, Furlow JD, Bodine SC. The glucocorticoid receptor and FOXO1 synergistically activate the skeletal muscle atrophy-associated MuRF1 gene. Am J Physiol Endocrinol Metab 295: E785–E797, 2008. doi: 10.1152/ajpendo.00646.2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Wang Q, McPherron AC. Myostatin inhibition induces muscle fibre hypertrophy prior to satellite cell activation. J Physiol 590: 2151–2165, 2012. doi: 10.1113/jphysiol.2011.226001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Welle S, Brooks AI, Delehanty JM, Needler N, Bhatt K, Shah B, Thornton CA. Skeletal muscle gene expression profiles in 20-29 year old and 65-71 year old women. Exp Gerontol 39: 369–377, 2004. doi: 10.1016/j.exger.2003.11.011. [DOI] [PubMed] [Google Scholar]
  • 53.Welle S, Brooks AI, Delehanty JM, Needler N, Thornton CA. Gene expression profile of aging in human muscle. Physiol Genomics 14: 149–159, 2003. doi: 10.1152/physiolgenomics.00049.2003. [DOI] [PubMed] [Google Scholar]
  • 54.Wissing ER, Boyer JG, Kwong JQ, Sargent MA, Karch J, McNally EM, Otsu K, Molkentin JD. P38α MAPK underlies muscular dystrophy and myofiber death through a Bax-dependent mechanism. Hum Mol Genet 23: 5452–5463, 2014. doi: 10.1093/hmg/ddu270. [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from Physiology are provided here courtesy of American Physiological Society

RESOURCES