543fc6bb0cf21227a11b7b45

REVIEW ARTICLE

Mechanisms regulating skeletal muscle growth and atrophyStefano Schiaffino1,2, Kenneth A. Dyar1, Stefano Ciciliot1, Bert Blaauw1,3 and Marco Sandri1,3

1 Venetian Institute of Molecular Medicine, Padova, Italy

2 Consiglio Nazionale delle Ricerche, Institute of Neuroscience, Padova, Italy

3 Department of Biomedical Sciences, University of Padova, Italy

Keywords

FoxO; IGF1; mTOR; myostatin; muscle

atrophy; muscle hypertrophy; protein

degradation; protein synthesis; satellite cells

Correspondence

S. Schiaffino; M. Sandri, Venetian Institute

of Molecular Medicine, Via Orus 2, 35129

Padova, Italy

Fax: +39 49 7923 250

Tel: +39 49 7923 232; +39 49 7923 258

E-mail: [email protected]; marco.

[email protected]

Website: www.vimm.it

(Received 25 January 2013, revised 13

March 2013, accepted 14 March 2013)

doi:10.1111/febs.12253

Skeletal muscle mass increases during postnatal development through a

process of hypertrophy, i.e. enlargement of individual muscle fibers, and a

similar process may be induced in adult skeletal muscle in response to con-

tractile activity, such as strength exercise, and specific hormones, such as

androgens and b-adrenergic agonists. Muscle hypertrophy occurs when theoverall rates of protein synthesis exceed the rates of protein degradation.

Two major signaling pathways control protein synthesis, the IGF1AktmTOR pathway, acting as a positive regulator, and the myostatinSmad2/3 pathway, acting as a negative regulator, and additional pathways have

recently been identified. Proliferation and fusion of satellite cells, leading to

an increase in the number of myonuclei, may also contribute to muscle

growth during early but not late stages of postnatal development and in

some forms of muscle hypertrophy in the adult. Muscle atrophy occurs

when protein degradation rates exceed protein synthesis, and may be

induced in adult skeletal muscle in a variety of conditions, including starva-

tion, denervation, cancer cachexia, heart failure and aging. Two major pro-

tein degradation pathways, the proteasomal and the autophagiclysosomalpathways, are activated during muscle atrophy and variably contribute to

the loss of muscle mass. These pathways involve a variety of atrophy-

related genes or atrogenes, which are controlled by specific transcription

factors, such as FoxO3, which is negatively regulated by Akt, and NF-jB,which is activated by inflammatory cytokines.

Introduction

Skeletal muscle mass and muscle fiber size vary accord-

ing to physiological and pathological conditions. An

increase in muscle mass and fiber size, i.e. muscle growth

or hypertrophy, occurs during development and in

response to mechanical overload (incapacitation or

ablation of synergistic muscles, strength training,

Abbreviations

4E-BP1, eukaryotic translation initiation factor 4E-binding protein 1; ACVR2, activin receptor 2; ALK4/5, activin receptor-like kinase 4/5;

AMPK, AMP-activated protein kinase; BNIP3, BCL2/adenovirus E1B 19 kDa protein-interacting protein 3; Fbxo, F-box only protein; Fn14,

fibroblast growth factor-inducible 14; FoxO, Forkhead box O; HDAC, histone deacetylase; IGF1, insulin-like growth factor 1; IKKb, IjB

kinase b; IL, interleukin; KLF15, Kruppel-like factor 15; MAFbx, muscle atrophy F-box; mTOR, mammalian target of rapamycin; mTORC1/2,

mTOR complex 1/2; MuRF1, muscle RING finger 1; NF-jB, nuclear factor j light-chain enhancer of activated B cells; nNOS, neuronal nitric

oxide synthase; PPAR, peroxisome proliferator-activated receptor; PGC-1a, PPAR-c co-activator-1a; PI3K, phosphatidylinositide-3-kinase;

PINK1, phosphatase and tensin homolog-induced putative kinase 1; REDD1, regulated in development and DNA damage responses 1; SRF,

serum response factor; TGF, transforming growth factor; TNFa, tumor necrosis factor a; TRAF, TNF receptor-associated factor; Trim32,

tripartite motif-containing protein 32; TWEAK, TNF-like weak inducer of apoptosis; VPS34, vacuolar protein sorting 34; YY1, Yin Yang 1.

4294 FEBS Journal 280 (2013) 42944314 2013 The Authors Journal compilation 2013 FEBS

reloading after unloading) or anabolic hormonal stimu-

lation (testosterone or b2-adrenergic agonists). Adecrease in muscle mass and fiber size, i.e. muscle atro-

phy, results from aging, starvation, cancer, diabetes,

bed rest, loss of neural input (denervation, motor neu-

ron disease) or catabolic hormonal stimulation (corti-

costeroids). The regulation of muscle mass and fiber size

essentially reflects protein turnover, i.e. the balance

between protein synthesis and degradation within the

muscle fibers. However, skeletal muscle fibers are multi-

nucleated structures, thus protein turnover may also be

affected by cell or nuclear turnover, i.e. addition of new

myonuclei, due to fusion of satellite cells, or loss of

myonuclei, due to nuclear apoptosis. Before separately

considering conditions of muscle growth and muscle

atrophy, it is useful to make some general points. First,

skeletal muscles and muscle fiber types vary, often dras-

tically, in their response to the same stimulus. This point

is evident in many experimental models used in muscle

research, without considering extreme cases such as the

differential response to testosterone in sexually dimor-

phic, androgen-sensitive muscles. For example, denerva-

tion in the rat diaphragm muscle causes atrophy of type

2X and 2B fibers, no change in type 2A fibers and slight

hypertrophy of type 1 fibers [1]. Similar changes are

found in other fast rat muscles (S. Schiaffino and

S. Ciciliot, unpublished data); however, the type 1 fibers

of the slow soleus show marked atrophy after denerva-

tion, thus the same fiber type may undergo opposite

changes in different muscles. With regard to nutrient

deprivation, slow muscles, such as the soleus, are less

sensitive to starvation compared to fast muscles [2]. This

response is presumably related to the different sensitiv-

ity of fast and slow muscles to corticosteroids [3]. Even

within the same muscle, for example the rat diaphragm,

corticosteroid treatment causes atrophy of type 2B and

2X fibers but not type 2A and 1 fibers [4]. This differen-

tial response probably reflects the fact that contractile

activity, which is greater in the continuously active

type 1 and 2A fibers, opposes the atrophic process,

possibly via the transcriptional co-activator PPAR-cco-activator-1a (PGC-1a) [5].Second, changes in protein turnover leading to

muscle hypertrophy or atrophy do not always pro-

ceed according to the simplistic equations suggested

by the balance analogy, i.e. muscle hypertrophy

results from increased protein synthesis and decreased

protein degradation, while muscle atrophy results

from decreased protein synthesis and increased pro-

tein degradation. Goldbergs analyses of muscle

growth in hypophysectomized rats showed that, dur-

ing hypertrophy of the soleus muscle induced by ten-

otomy of the gastrocnemius, there is decreased

protein catabolism as well as increased synthesis of

new proteins, while during hypertrophy of the soleus

induced by growth hormone, there is increased pro-

tein synthesis without any change in protein degrada-

tion rates [6]. Starvation causes decreased protein

synthesis and increased protein degradation in both

fast and slow rat muscles [2]. However, muscle dener-

vation is accompanied by increased protein degrada-

tion and increased rather than decreased protein

synthesis [7,8]. One must also consider that fast and

slow muscles differ in their protein turnover rates,

with slow muscles showing higher rates of both pro-

tein synthesis and degradation [2].

Third, changes in protein turnover and cell/nuclear

turnover in skeletal muscle do not always proceed in

parallel, thus myonuclear domains may vary. The myo-

nuclear domain size, defined as the cytoplasmic volume

per myonucleus, varies among fiber types, being larger

in type 2B and 2X fibers compared to type 2A and 1

fibers, and, in contrast to initial reports, is not constant

in various conditions [8a]. For example, during muscle

atrophy caused by denervation or corticosteroids, there

is no loss of myonuclei; therefore the myonuclear

domain decreases in proportion to the decrease in

cross-sectional area [1,4,9,10]. Conversely, myonuclear

domains may increase during postnatal muscle growth

and in different hypertrophy models (see below).

Muscle growth

In this section, we focus on muscle growth processes

that take place after birth, including muscle growth

during postnatal development and the process of mus-

cle hypertrophy induced in adult muscle by functional

overload. We do not deal specifically with muscle

growth during regeneration, which has been discussed

previously [11].

Major signaling pathways controlling muscle

growth

Two major signaling pathways control skeletal muscle

growth: the insulin-like growth factor 1 phosphoino-sitide-3-kinaseAkt/protein kinase Bmammalian targetof rapamycin (IGF1PI3KAkt/PKBmTOR) path-way acts as a positive regulator of muscle growth, and

the myostatinSmad3 pathway acts as a negative regu-lator (Fig. 1A). The role of the IGF1 pathway has

been supported by a variety of gain- and loss-of-func-

tion genetic approaches [12]. For example, muscle-spe-

cific inactivation of the IGF1 receptor impairs muscle

growth due to reduced muscle fiber number and size

[13]. Conversely, muscle-specific over-expression of

FEBS Journal 280 (2013) 42944314 2013 The Authors Journal compilation 2013 FEBS 4295

S. Schiaffino et al. Mechanisms regulating muscle growth and atrophy

IGF1 causes muscle hypertrophy [14]. In vivo transfec-

tion studies in adult mouse and rat muscles have

helped to elucidate the pathways downstream of the

IGF1 receptor. IGF1 is known to activate both the

mitogen-activated protein kinase/extracellular signal-

regulated kinase (MAPK/ERK) and the PI3KAktpathways. However, only a Ras mutant that selectively

activates the PI3KAkt pathway was able to inducehypertrophy of transfected fibers, whereas a Ras

mutant acting specifically on the ERK pathway did

not [15]. Accordingly, constitutively active Akt results

in a striking hypertrophy of transfected muscle fibers

[16,17], with a similar effect being seen using inducible

muscle-specific transgenic models [1820].Akt stimulates protein synthesis by activating

mTOR and its downstream effectors. The kinase

mTOR interacts with several proteins to form two

complexes: mTOR complex 1 (mTORC1) containing

raptor and mTOR complex 2 (mTORC2) containing

rictor. Here, we focus on mTOR signaling as it relates

to skeletal muscle growth, as a detailed general discus-

sion of mTOR function and regulation has been pub-

lished previously [21]. It should be stressed that

mTOR responds to multiple upstream signals in addi-

tion to Akt, including amino acids, and it controls sev-

eral cellular processes in addition to protein synthesis,

including autophagy. The crucial role of mTOR in

mediating muscle growth is supported by genetic and

pharmacological evidence. Muscle-specific mTOR

knockout causes reduced postnatal growth, due to the

reduced size of fast but not slow muscle fibers, and

severe myopathy [22]. A similar phenotype is found in

A

B

Fig. 1. Signaling modules responsible for skeletal muscle growth during development, regeneration and overload-induced hypertrophy in the

adult. We postulate that all these modules converge to a final common pathway centered on mTOR and its effectors that control protein

synthesis. (A) Major signaling pathways. IGF1 stimulates mTOR activity and muscle growth via PI3KAkt. Follistatin induces muscle growth by

inhibiting myostatin and activin A. The two pathways cross-talk by direct interaction between Smad3 and Akt. In addition, transcriptional regulation

by Smad3/Smad4 heterodimers may repress mTOR and protein synthesis through mechanisms that have not yet been defined. The arrow

connecting mTOR with a myonucleus indicates transcriptional roles of mTOR. (b) Additional pathways controlling mTOR activity and protein

synthesis. The SRF (serum response factor), PA (phosphatidic acid) and nNOS (neuronal nitric oxide synthase) pathways may be activated by

mechanical overload. The dotted arrow connecting newly fused myonuclei (new mn) to the mTOR pathway indicates the postulated increase in

protein synthesis and myotube/myofiber growth associated with myoblast/satellite cell fusion. SC, satellite cell; SSC, satellite stem cell.


Mechanisms regulating muscle growth and atrophy S. Schiaffino et al.

mice lacking raptor in skeletal muscle, whereas those

lacking rictor have a normal phenotype, supporting a

major role of mTORC1 in mediating the effect of

mTOR on protein synthesis [23]. Rapamycin, a specific

mTOR inhibitor, acts especially on mTORC1,

although mTORC2 is also affected during chronic

treatment [21]. Rapamycin inhibits muscle growth dur-

ing postnatal development [24], muscle regeneration

[17] (Fig. 2) and compensatory muscle hypertrophy

induced by synergist elimination [16]. Muscle growth

during reloading of unloaded muscles is only partially

inhibited by rapamycin [24]. Muscle fiber hypertrophy

induced by transfection of adult muscles with a consti-

tutively active Akt construct is also blunted by rapa-

mycin [16,17].

Two major effectors of mTORC1 that promote pro-

tein synthesis are eukaryotic translation initiation fac-

tor 4E-binding protein 1 (4E-BP1) and S6 kinase 1.

Muscle growth is apparently unaffected by 4E-BP1

knockout [25]. In contrast, deletion of S6 kinase 1

causes muscle atrophy and partially prevents the

response to constitutively active Akt [26]. However,

the control of protein synthesis by mTOR is still

incompletely characterized [21]. The growth-promoting

effect of mTORC1 is repressed by AMP-activated pro-

tein kinase (AMPK), and hypertrophy of soleus mus-

cle has been described in AMPK-deficient mice [26].

Another aspect of mTOR function that is incom-

pletely understood is the role of mTOR in transcrip-

tional regulation. In both yeast and mammalian cells,

TOR/mTOR controls cell growth by coordinately reg-

ulating the synthesis of ribosomes and tRNAs and

activating transcription by all three nuclear RNA

polymerases (I, II and III). However, ribosomal RNA

accumulation induced in overloaded muscles after syn-

ergist ablation is only partially inhibited by rapamycin

[27]. On the other hand, blockade of mTOR by

rapamycin in cultured myotubes is sufficient to block

most IGF1-induced changes in transcription [28].

What are the downstream effectors of mTOR that reg-

ulate transcription? The transcription factor

Yin Yang 1 (YY1) physically interacts with mTORC1

and mediates mTOR-dependent regulation of mito-

chondrial gene expression via a YY1PGC-1a complex[29]. More recently, active mTORC1 was found to

induce YY1 phosphorylation, resulting in displacement

of the polycomb repressor complex, thereby activating

transcription of many genes of the insulin/IGF1Aktpathway, including Igf1, Irs1, Irs2, Akt1 and Akt2.

Conversely, mTORC1 inactivation induced by rapa-

mycin results in YY1 dephosphorylation and recruit-

ment of the polycomb repressor complex to the

promoter of these genes, with a consequent block of

transcription [30]. Muscle-specific inactivation of the

Yy1 gene leads to up-regulation of genes of the insu-

lin/IGF1Akt pathway [30]; however, the effect of thisknockout on muscle growth has not been described.

The second major signaling pathway that controls

skeletal muscle growth involves myostatin, a member

of the transforming growth factor b (TGFb) superfam-ily. Myostatin is produced by skeletal muscle and acts

as a negative regulator of muscle growth, as shown by

the finding that myostatin mutations in various mam-

malian species cause muscle hypertrophy [31]. Purified

myostatin inhibits protein synthesis and reduces myo-

tube size when added to differentiated myotubes in

culture [32]. Furthermore, muscle atrophy is induced

in mice by systemically administered myostatin [31].

Muscle hypertrophy may also be induced by inhibitory

extracellular binding proteins, such as follistatin,

whose effect is even greater than the lack of myostatin,

because it binds to other TGFb superfamily members,such as activin A, that act as negative regulators of

muscle growth like myostatin does (Fig. 1A). Myosta-

Fig. 2. Muscle growth in regenerating

skeletal muscle is dependent on mTOR

activity. Rapamycin, a specific mTOR

inhibitor, inhibits growth of regenerating

muscle. The section was stained with an

antibody specific for embryonic myosin

heavy chain [177]. Modified from [17].

Scale bar = 50 lm.



tin and activin A interact and activate a heterodimeric

receptor complex with serinethreonine kinase activity,comprising a type II receptor, activin receptor 2

(ACVR2 and ACVR2B), and a type I receptor, activin

receptor-like kinase 4 and 5 (ALK4 and ALK5). A

soluble form of ACVR2B acts as a myostatin/acti-

vin A inhibitor that is capable of inducing muscle

hypertrophy in adult mice. Myostatin/activin A signal-

ing in myofibers is mediated by phosphorylation and

nuclear translocation of Smad2 or Smad3 transcription

factors, and formation of heterodimers with Smad4.

Although the transcriptional targets of the Smad2/

Smad4 and Smad3/Smad4 complexes that mediate the

inhibitory effect on growth are not known, it is possi-

ble that myostatin/activin A signaling interferes with

the AktmTOR pathway [33,34]. For example, musclehypertrophy induced by transfection of dominant-neg-

ative ACVR2B is partially prevented by mTOR-spe-

cific siRNAs or by rapamycin [33]. Likewise,

follistatin-induced muscle hypertrophy is blunted by

blocking the IGF1AktmTOR pathway at the levelof the IGF1 receptor, via a dominant-negative IGF1

receptor, at the level of Akt, via dominant-negative

Akt, or at the level of mTOR, via rapamycin. How-

ever, follistatin-induced muscle hypertrophy was

unchanged in S6 kinase 1/2 knockout mice [35].

Another recent study showed that virus-mediated

over-expression of a form of follistatin that remains

localized within the injected muscle stimulates Akt

phosphorylation, mTOR signaling and protein synthe-

sis, leading to a striking muscle hypertrophy that is

inhibited by rapamycin [36]. Inhibition of Smad3 activ-

ity by follistatin is critical for activation of AktmTOR signaling, as constitutively active Smad3 was

found to suppress follistatin-induced muscle growth

and mTOR activation. It is also possible that a direct

interaction between Smad3 and Akt, as demonstrated

in other cell systems [37,38], may be involved in cross-

talk between the myostatin/activin A and IGF1 path-

ways in skeletal muscle.

Additional signaling pathways controlling muscle

growth

As shown in Fig. 1B, other important signaling path-

ways are also known to control skeletal muscle growth.

The transcription factor SRF (serum response factor) is

required for muscle growth during development, as

shown by muscle-specific SRF knockout [39,40]. Using

an inducible, muscle-specific knockout model, SRF was

also found to be required for muscle hypertrophy

induced by synergist elimination [41]. In this model, the

effect of SRF is apparently mediated via release of

interleukins 4 and 6 (IL-4 and IL-6), which act in a

paracrine manner to induce satellite cell proliferation

and fusion [41] (see below). Akt phosphorylation is

apparently unchanged in control and SRF mutant mus-

cles at various time points after synergist ablation.

However, a previous study reported that SRF is able

to activate the Akt pathway via a muscle-enriched

microRNA, miR-486, that targets the phosphatase and

tensin homolog PTEN, which negatively affects PI3KAkt signaling [42]. In addition, as activation of mTOR

and its downstream effectors were not examined in the

previous study [41], it is possible that mTOR activity

and protein synthesis are increased by SRF in muscles

undergoing overload-induced hypertrophy, either via a

transcriptional mechanism mediated by SRF itself or as

a secondary consequence of incorporation of new

myonuclei resulting from satellite cell activation and

fusion. Finally, it should be stressed that SRF is known

to control the transcription of several cytoskeletal and

sarcomeric protein genes, including those for a-actin,by binding to CArG box regulatory elements.

The AktmTOR pathway is also a point of conver-gence for additional signaling pathways that are

known to promote muscle growth. This appears to be

the case for androgens and b2-adrenergic agents, whichboth have well-known anabolic effects on skeletal mus-

cles. Androgens potently stimulate muscle growth: tes-

tosterone loss in male mice decreases muscle Igf1

mRNA, Akt phosphorylation and the rate of myofibr-

illar protein synthesis; these changes are all reversed

by nandrolone treatment [43]. Muscle hypertrophy

induced by b2-adrenergic agents, such as clenbuterolor formoterol, is accompanied by a significant increase

in Akt phosphorylation [44] and is completely blocked

by rapamycin [45]. Another signaling pathway that

controls muscle growth involves Wnt7a, an extracellu-

lar protein that acts both on satellite stem cells,

increasing their numbers, and on myofibers by activat-

ing the PI3KAkt pathway via its receptor Fzd7[46,47]. A pathway linked to mTOR activation

involves neuronal nitric oxide synthase (nNOS). When

activated in myofibers by functional overload, nNOS

generates nitric oxide (NO) and causes peroxynitrite-

dependent activation of the cation channel Trpv1,

located in the sarcoplasmic reticulum. The resulting

increase in intracellular Ca2+ induced by Trpv1-medi-

ated Ca2+ release triggers activation of mTOR [48].

Furthermore, mTOR is also activated by mechanical

stimulation via an Akt-independent pathway involving

phosphatidic acid and activated phospholipase D [49].

A novel form of PGC-1a (PGC-1a4), which resultsfrom alternative promoter usage and splicing of the

primary transcript, is involved in muscle growth, as



shown by the finding that mice with skeletal muscle-

specific transgenic expression of PGC-1a4 showincreased muscle mass and strength [50]. PGC-1a4,which is expressed at significant levels in skeletal mus-

cle, is a shorter, truncated form of the previously

described PGC-1a [50a], now referred to as PGC-1a1,which is involved in mitochondrial biogenesis and not

in muscle growth. In cultured muscle cells, PGC-1a4was found to induce IGF1 and repress myostatin, thus

promoting myotube hypertrophy, which was blocked

by an IGF1 receptor inhibitor. Myotube growth

induced by treatment with clenbuterol was also

blunted by PGC-1a4 knockdown.

Satellite cell fusion and increase of myonuclei

during muscle growth

The issue of whether satellite cell proliferation and

fusion contributes to muscle growth has been the sub-

ject of debate [51]. There is no doubt that myoblast

fusion is essential for muscle growth during early stages

of muscle differentiation. For example, myotube

growth in culture is impaired when myoblast fusion is

inhibited, either during formation of the nascent myo-

tube or during the transition from nascent to mature

myotube [52]. IL-6 and IL-4 released by the myotubes

act on myoblasts, promoting their proliferation and

fusion, respectively [53,54]. Muscle cells lacking IL-4 or

the IL-4a receptor subunit form smaller myotubes withfewer myonuclei [53]. Muscle growth during early post-

natal development (from P0 to approximately P21 in

mice and rats) is also accompanied by, and presumably

dependent on, a continuous increase in the number of

myonuclei resulting from satellite cell fusion [55] (an

approximately fivefold increase from P3 to P21 in

mouse extensor digitorum longus muscle [56]). Muscle

regeneration recapitulates many aspects of embryonic

and neonatal myogenesis, with satellite cells acting as a

major myogenic stem cell, and undergoing active prolif-

eration and fusion during formation of new myofibers

[11]. A distinct feature of muscle regeneration, which is

missing in normal muscle development, is the central

role of inflammation and of various macrophage popu-

lations in the muscle growth process [57].

On the other hand, muscle hypertrophy at late post-

natal stages takes place without a significant contribu-

tion of satellite cell fusion. For example, the

approximately twofold increase in myofiber cross-sec-

tional area from P21 to P56 in mouse extensor digito-

rum longus muscle occurs with a negligible change in

myonuclear number [56]. In adult skeletal muscle, clen-

buterol-induced hypertrophy does not involve satellite

cell fusion [58], although satellite cell activation is

induced by androgens [59]. Similarly, myonuclear

number is not increased during muscle growth upon

reloading of unloaded muscles [9,60]. Satellite cell pro-

liferation and fusion were not detected during muscle

hypertrophy induced by a muscle-specific, inducible

and constitutively active Akt1 transgene [18], or by

over-expression of JunB [61]. The contribution of

satellite cells to muscle hypertrophy induced by block-

ade of the myostatin/activin A pathway is controver-

sial. In two studies, satellite cell activation was not

detected after injection of vectors encoding the myost-

atin propeptide, which binds non-covalently to myost-

atin and inhibits its activity [62], or in hypertrophic

muscles expressing dominant-negative ACVR2B [33].

In contrast, another study reported an increase in

BrdU-positive myonuclei and increased numbers of

satellite cells when using a soluble ACVR2B to block

this pathway [63]. A recent detailed study concluded

that satellite cells play little or no role in myostatin/

activin A signaling in vivo, based on the finding that

satellite cell and myonuclear number were unchanged

in hypertrophic muscles after injection of soluble

ACVR2B, and that muscle hypertrophy induced by

over-expressing follistatin also occurs in mice lacking

syndecan4 or Pax7, which have compromised satellite

cell function or number, respectively [64].

A widely used model of muscle hypertrophy in adult

mice or rats is compensatory hypertrophy induced by

ablation of synergist muscles: for example hypertrophy

of the plantaris or soleus muscle after removal of the

gastrocnemius, or hypertrophy of the extensor digito-

rum longus after removal of the tibialis anterior mus-

cle. This model of acute functional overload causes

immediate satellite cell proliferation and fusion [65,66],

with a consequent increase in the number of myonuclei

[54,67]. A similarly dramatic increase in mechanical

load may be induced in human skeletal muscle by

high-intensity eccentric contractions, which also cause

proliferation of satellite cells [68] and, when repeated,

are known to induce muscle hypertrophy. Depending

on the intensity of the exercise and the trained/

untrained state of the host, eccentric contractions may

cause a spectrum of responses, ranging from severe

muscle damage and local inflammation to mild muscle

damage (myofibrillar disruptions) without inflamma-

tion, to remodeling of the extracellular matrix without

obvious damage to the muscle fibers. Similar changes

have been reported in rat or mouse muscles after elimi-

nation of synergists, and it is possible that the satellite

cell activation seen in each of these conditions may

reflect a response that is similar to that observed in

muscle regeneration. A role of satellite cells in IGF1-

induced muscle hypertrophy was suggested by the



inhibitory effect of gamma radiation [69]; however,

exposure to radiation may affect protein synthesis

within the myofibers, thus complicating interpretation

of this experiment [70].

IL-6 produced by myofibers and satellite cells is

increased in overloaded muscles, suggesting a role for

IL-6 and downstream signal transducer and activator

of transcription 3 (STAT3) signaling in satellite cell

proliferation, myonuclear accretion and hypertrophy

[54]. IL-4 was found to control myoblast fusion, thus

promoting the transition from nascent to mature myo-

tubes in cultured muscle cells and during postnatal

growth [53]. A recent study indicated that satellite cell

proliferation and fusion during overload-dependent

hypertrophy are induced by IL-6 and IL-4, respec-

tively, and that both cytokines are released by over-

loaded myofibers under the control of SRF [41]. IL-4

expression is decreased by muscle-specific SRF knock-

out, which causes reduced postnatal muscle growth

and reduced hypertrophy after synergist ablation [40],

with both defects being rescued by IL-4 but not IL-6

over-expression [41]. However, another recent study

reported that recruitment of satellite cells is not

required for muscle hypertrophy, because hypertrophy

was unchanged in mice in which more than 90% of

satellite cells were ablated by diphtheria toxin A using

an inducible Pax7diphtheria toxin A transgene [71].How may these opposite conclusions about the mecha-

nism of muscle hypertrophy induced by functional

overload be reconciled?

It is clear that skeletal muscle has the capacity to

activate two modes of hypertrophy, either with or

without satellite cell involvement, as clearly shown by

the two stages of muscle growth during postnatal

development. These two modes of hypertrophy are

also observed at later stages, and may be activated in

the adult in response to various stimuli. It is likely that

satellite cells are activated and contribute to hypertro-

phy when an acute stimulus is involved, such as after

elimination of synergistic muscles, or after strong exer-

cise with eccentric contractions, i.e. under conditions

when some form of muscle damage occurs. In con-

trast, more gradual exercise, or reloading of unloaded

muscles, does not trigger satellite cell activation and

fusion. It may be envisaged that, when one of the two

available modes of response is artificially impaired, as

is the case in gene or cell ablation models, the muscle

will use the remaining available mode. Thus, in the

absence of satellite cells, a growth response is still

induced by functional overload, but via increased pro-

tein synthesis alone. Loss-of-function approaches,

involving either genes or cells, should thus be inter-

preted with caution, because compensatory adapta-

tions may occur, not only when the perturbation is

produced during development, as in traditional knock-

out models, but also when it is produced in the adult

using inducible systems.

Muscle atrophy

Muscle atrophy involves the shrinkage of myofibers

due to a net loss of proteins, organelles and cytoplasm.

Acute muscle atrophy, as occurs in many pathological

conditions, is due to hyperactivation of the cells main

degradation pathways, including the ubiquitinprotea-some system and the autophagylysosome pathway.Recent studies have highlighted a complex scenario

whereby these catabolic pathways modulate one

another at different levels, and are also coupled at var-

ious points to biosynthetic pathways. The result is a

coordinated balance between protein degradation and

synthesis that reflects the physiological state of the

muscle fiber. As muscle accounts for such a large pro-

portion of total body mass, particularly total body

protein, this local balance has a significant effect on

general protein homeostasis.

The ubiquitinproteasome and autophagylysosome machinery are activated in atrophying

muscles

Activation of the cells proteolytic systems is transcrip-

tionally regulated, and a subset of genes that are com-

monly up- or down-regulated has been identified in

atrophying skeletal muscle, regardless of the catabolic

condition [7275]. These common genes are thought toregulate the loss of muscle components, and were thus

designated atrophy-related genes or atrogenes [7577]. Among the up-regulated atrophy-related genes are

transcripts belonging to the ubiquitinproteasome andautophagylysosome systems. The up-regulation ofseveral ubiquitinproteasome and autophagy-relatedgenes is normally blocked by Akt through negative

regulation of Forkhead box O (FoxO) transcription

factors [7779].In muscle, the ubiquitinproteasome system is

required to remove sarcomeric proteins in response to

changes in muscle activity. The rate-limiting step of

the ubiquitination process, which affects subsequent

proteasome-dependent degradation, is catalysed by the

E3 enzyme, which is a ubiquitin ligase. Among the

known E3s, only a few are both muscle-specific and

up-regulated during muscle loss. The first to be

identified were atrogin-1/MAFbx (muscle atrophy F-

box) and muscle RING finger 1 (MuRF1). Mice lack-

ing atrogin-1/MAFbx and MuRF1 are resistant to



muscle atrophy induced by denervation [72]. More-

over, knockdown of atrogin-1 prevents muscle loss

during fasting [80], whereas MuRF1 knockout mice

(but not atrogin-1 knockout mice) are also resistant to

dexamethasone-induced muscle atrophy [81]. So far,

very few muscle proteins have been identified as sub-

strates for atrogin-1, and those that have been identi-

fied appear to be involved in growth-related processes

or survival pathways. For example, atrogin-1 promotes

degradation of MyoD, a key muscle transcription fac-

tor, and of eukaryotic translation initiation factor 3

subunit F (eIF3-f), an important activator of protein

synthesis [82,83]. In the heart, atrogin-1 ubiquitinates

and reduces the levels of calcineurin A, an important

factor triggering cardiac hypertrophy in response to

pressure overload [84]. Interestingly, immunoprecipita-

tion experiments in C2C12 myoblasts and myotubes

have found that atrogin-1 interacts with sarcomeric

proteins, including myosins, desmin and vimentin, as

well as transcription factors, components of the trans-

lational machinery, enzymes involved in glycolysis and

gluconeogenesis, and mitochondrial proteins [85].

Whether atrogin-1 ubiquitinates these proteins has yet

to be proven. Conversely, MuRF1 was reported to

interact with and control the half-life of many impor-

tant muscle structural proteins, including troponin I

[86], myosin heavy chains [87,88], actin [89], myosin

binding protein C and myosin light chains 1 and 2

[90]. Presumably, additional E3s that have not yet

been identified are also activated during atrophy to

promote the clearance of soluble cellular proteins and

to limit/regulate anabolic processes. A recent paper

reported that Trim32 (tripartite motif-containing pro-

tein 32) is a crucial E3 ligase for the degradation of

thin filaments (actin, tropomyosin and troponins),

a-actinin and desmin [91]. However, Trim32 knockoutmice are not protected from atrophy, but instead show

impaired recovery of muscle mass after atrophy [92].

Another E3 ubiquitin ligase that has been found to

play a critical role in atrophy is TRAF6 (TNF recep-

tor-associated factor) [93], which mediates the conjuga-

tion of Lys63-linked polyubiquitin chains to target

proteins. Lys48-linked polyubiquitin chains are a sig-

nal for proteasome-dependent degradation, but Lys63-

linked polyubiquitin chains play other roles, such as

regulating autophagy-dependent cargo recognition by

interacting with the scaffold protein p62 (also known

as SQSTM1) [9496]. Muscle-specific TRAF6 knock-out mice have a decreased amount of polyubiquitinat-

ed proteins, almost no Lys63-polyubiquitinated

proteins in starved muscles [97], and are resistant to

muscle loss induced by denervation, cancer or starva-

tion [93,97,98]. The mechanism of this protection

involves both direct and indirect effects of TRAF6 on

protein breakdown. In fact, TRAF6-mediated ubiquiti-

nation is required for the optimal activation of c-Jun

N-terminal kinase, AMPK, FoxO3 and NF-jB [97].All of these factors are crucial regulators of atrogin-1

and MuRF1 expression and of several autophagy-

related genes. Inhibition of TRAF6 reduces the

induction of atrogin-1 and MuRF1, thereby preserving

muscle mass under catabolic conditions.

Specific ubiquitin ligases may be involved in differ-

ent models of muscle wasting and at different stages of

the atrophy process. For instance, the HECT domain

ubiquitin ligase Nedd4-1 has been reported to be up-

regulated mainly during muscle disuse. Indeed, dele-

tion of the Nedd4-1 gene specifically in skeletal muscle

results in partial protection from muscle atrophy in

denervated type II fibers. However, Nedd4-1 knockout

mice have smaller muscles, suggesting that this E3 may

play additional roles during myogenesis or in the con-

trol of protein synthesis [99].

Mul1 is a mitochondrial ubiquitin ligase that plays

an important role in mitochondrial network remodel-

ing. Mul1 is up-regulated by the FoxO family of tran-

scription factors under catabolic conditions, such as

fasting or denervation, and causes mitochondrial frag-

mentation and removal via autophagy (mitophagy)

[100]. Importantly, knocking down Mul1 spares muscle

mass during fasting. Mul1 ubiquitinates the mitochon-

drial pro-fusion protein mitofusin 2, causing its degra-

dation via the proteasome system. The exact

mechanism that triggers Mul1-dependent mitochon-

drial dysfunction and mitophagy is unclear, but it has

been reported that mitofusin degradation is permissive

for mitochondrial fission and mitophagy [101].

Carboxy terminus of Hsc70 interacting protein

(CHIP) is another ubiquitin ligase, which regulates

ubiquitination and lysosomal-dependent degradation

of filamin C, a muscle protein found in the Z-line

[102]. Filamins undergo unfolding and refolding cycles

during muscle contraction, and are therefore prone to

irreversible damage [102]. Alterations to filamin struc-

ture trigger binding of the co-chaperone BAG3, which

is a complex comprising the chaperones Hsc70 and

HspB8, as well as the ubiquitin ligase CHIP. CHIP

ubiquitinates BAG3 and filamin, which are recognized

and delivered to the autophagy system by p62 [102].

Interestingly, filamin B half life is controlled, at least

during myogenesis, by another ubiquitin ligase,

ASB2b, which is mainly expressed in muscle cells. Inthis case, ubiquitination of filamin B by ASB2b leadsto proteasome-dependent degradation [103].

In skeletal muscle, E3 ligases also have important

regulatory functions in signaling pathways. For



example, it was recently found that the ubiquitin ligase

Fbxo40 (F-box only protein) regulates anabolic signals

[104]. Fbxo40 ubiquitinates and affects the degradation

of insulin receptor substrate 1, a downstream effector

of insulin receptor-mediated signaling. Inhibition of

Fbxo40 by RNAi induces hypertrophy in myotubes,

and Fbxo40 knockout mice display bigger muscle

fibers [104].

Although some E3 ligases involved in muscle protein

ubiquitination and breakdown have been identified,

very little is known about how ubiquitinated proteins

are recognized and delivered to the proteasome.

ZNF216 has been identified as an important player in

the recognition and delivery of ubiquitinated proteins

to the proteasome during muscle atrophy. Interest-

ingly, ZNF216 is up-regulated by FoxO transcription

factors in atrophying muscles, and ZNF216-deficient

mice are partially resistant to muscle loss during dener-

vation. The absence of ZNF216 in muscle leads to

accumulation of polyubiquitinated proteins [105].

Another important system for extraction and degra-

dation of ubiquitinated proteins from larger structures

is the p97/valosin containing protein (VCP) ATPase

complex. p97/VCP is induced during denervation, and

over-expression of a dominant-negative p97/VCP

reduces overall proteolysis by the proteasome and

lysosome pathways, and blocks the accelerated protein

breakdown induced by FoxO3. Interestingly, p97 and

its co-factors, Ufd1 and p47, have been found to be

associated with specific myofibrillar proteins, suggest-

ing a role for p97 in extracting ubiquitinated proteins

from myofibrils [106].

Although a great body of research has focused on

the ubiquitination process, little is known about the

role of deubiquitination and its contribution to muscle

atrophy. The largest class of deubiquitinating enzymes

are ubiquitin-specific proteases. So far, only two

ubiquitin-specific proteases (USP14 and USP19) have

been found to be up-regulated in atrophying muscles

[73,107]. Knockdown of USP19 in myotubes results in

decreased protein degradation and reverts dexametha-

sone-induced loss of myosin heavy chain [108].

Macroautophagy, hereafter referred to as auto-

phagy, is the other proteolytic system that is acti-

vated in catabolic conditions and that is under FoxO

regulation [109]. The various types of autophagy,

including their regulation and involvement in muscle

homeostasis, have been reviewed recently [110].

Briefly, autophagy is a highly conserved homeostatic

mechanism that is used for the degradation and recy-

cling, through the lysosomal machinery, of bulk cyto-

plasm, long-lived proteins and organelles [111].

Although autophagy was initially considered a

non-selective degradation pathway, the presence of

more selective forms of autophagy is becoming

increasingly evident. Indeed, autophagy may trigger

the selective removal of specific organelles, such as

mitochondria, via mitophagy. In mammals, parkin,

PINK1 (phosphatase and tensin homolog-induced

putative kinase 1), and BCL2/adenovirus E1B 19 kDa

protein-interacting protein 3 (Bnip3 and Bnip3L) have

been shown to regulate mitophagy, and inactivation

of the genes encoding these proteins leads to mito-

chondrial abnormalities [112,113]. PINK1 is normally

absent in healthy mitochondria because it is constitu-

tively degraded by mitochondrial proteases. However,

once mitochondria are damaged, PINK1 is no longer

degraded and accumulates. PINK1 induces parkin

recruitment to mitochondria, promoting mitophagy

through ubiquitination of outer mitochondrial mem-

brane proteins that are recognized by p62, which then

brings autophagic vesicles to ubiquitinated mitochon-

drial proteins [114,115]. Bnip3 and Bnip3L are

BH3-only proteins that are localized at the outer

membrane of the mitochondria after cellular stress,

and reportedly bind directly to LC3 (MAP1LC3A

microtubule-associated protein 1 light chain 3 alpha),

thereby recruiting the autophagosome to damaged

mitochondria [116,117]. In atrophying muscle, the

mitochondrial network is dramatically remodeled fol-

lowing fasting or denervation, and autophagy via

Bnip3 contributes to mitochondrial remodeling

[101,118120]. Expression of the fission machinery issufficient to cause muscle wasting in mice, whereas

inhibition of mitochondrial fission prevents muscle

loss during denervation, indicating that disruption of

the mitochondrial network is a crucial amplificatory

loop of the muscle atrophy program [101,118]. Con-

versely, impairment of basal mitophagy is deleterious

to muscle homeostasis, and leads to the accumulation

of damaged and dysfunctional mitochondria [121].

Accordingly, the phenotype of mice with muscle-spe-

cific inactivation of various genes coding for auto-

phagy-related proteins, such as Atg7, Atg5 or

nutrient-deprivation autophagy factor-1 (NAF-1), a

Bcl-2-associated autophagy regulator, results in atro-

phy, weakness and various myopathic features

[122124]. In addition, altered regulation of auto-phagy-related genes leads to muscle dysfunction. His-

tone deacetylases 1 and 2 (HDACs) were found to

regulate muscle autophagy by controlling the expres-

sion of autophagy genes. Muscle-specific ablation of

both HDAC1 and HDAC2 results in partial perinatal

lethality, and the HDAC1/2 knockout mice that do

survive develop a progressive myopathy characterized

by impaired autophagy [125,126].



IGF1AKTFoxO signaling

Several studies have shown that the IGF1 and/or insu-

lin signaling suppress protein breakdown while pro-

moting muscle growth [127129]. Additional datasupporting the role of the IGF1 pathway in regulating

muscle atrophy have been obtained from studies of

Akt. Electroporation of constitutively active Akt in

adult myofibers completely blocks muscle atrophy

induced by denervation [16]. Akt transgenic mice dis-

play muscle hypertrophy and protection from denerva-

tion-induced atrophy [19,20,130], showing that the Akt

pathway promotes muscle growth and simultaneously

blocks protein degradation [20,33]. In particular, Akt

regulates both the ubiquitinproteasome system andthe autophagylysosome pathway, and this action ismediated by FoxO transcription factors. The FoxO

family members that are important for skeletal muscle

include three isoforms: FoxO1, FoxO3 and FoxO4.

Akt phosphorylates all FoxOs, promoting their export

from the nucleus to the cytoplasm. As predicted, the

reduced activity of the Akt pathway observed in vari-

ous models of muscle atrophy leads to decreased levels

of phosphorylated FoxO in the cytoplasm and a

marked increase in nuclear FoxO [131] (Fig. 3). The

translocation and transcriptional activity of FoxO

members is sufficient to promote atrogin-1 and

MuRF1 expression, and muscle atrophy. Studies utiliz-

ing FoxO3 over-expression in adult muscle or muscle-

specific FoxO1 transgenic mice showed markedly

reduced muscle mass and fiber atrophy [77,132,133]. In

contrast, FoxO knockdown by RNAi blocks the up-

regulation of atrogin-1 expression during atrophy and

prevents muscle loss [77,134].

Cross-talk between protein breakdown and protein

synthesis is not limited to Akt, but also involves

FoxO. Activation of FoxO in Drosophila muscle up-

regulates 4E-BP1 [135] and represses mTOR via sestrin

[136]. Consistently, in mammals, FoxO3 reduces total

protein synthesis in adult muscle [137]. Thus, when

Akt is active, protein breakdown is suppressed, and

when FoxO is induced, protein synthesis is further

suppressed. This is not trivial, as FoxO activity is reg-

ulated by several post-translational modifications,

including phosphorylation, acetylation and mono- and

polyubiquitination [138]. Adding an additional level of

complexity, the regulatory consequences of these

changes appear to be specific for individual FoxO

members. For example, recent evidence suggests that

acetylation negatively regulates FoxO3 activity, but

has no effect on FoxO1 [139]. Mutants of FoxO3 that

mimic the effect of acetylation have cytosolic localiza-

tion and a reduced capacity to induce transcription of

the gene encoding atrogin-1, and cause muscle atrophy

[140]. Most of these regulatory mechanisms are Akt-

independent, and may play a role in muscle atrophy

induced by oxidative or energy stress.

Other studies have revealed a connection between

AMPK and FoxO3. AMPK phosphorylates several

Akt-independent sites on FoxO3, thereby stimulating

its transcriptional activity [141,142]. Indeed, treatment

Fig. 3. Protein degradation regulates

protein synthesis. In the presence of

growth factors, the PI3KAkt/protein

kinase B pathway sequesters FoxO1/3/4

transcription factors in the cytoplasm. In

the absence of growth factors, Akt is

inactive, and therefore, FoxOs are

translocated into the nucleus and induce

the transcription of target genes that

regulate the ubiquitinproteasome and

autophagylysosome systems. mTOR

senses the amino acids derived from the

proteasome, or, when localized on

lysosomes, the amino acid flux derived

from lysosomal protein breakdown, and is

therefore activated.



of muscle cultures with 5-aminoimidazole-4-carboxa-

mide riboside (AICAR), an activator of AMPK,

increases protein breakdown and atrogin-1 expression

via the FoxO family [143]. It has recently been shown

that FoxO3 is activated via AMPK in myofibers to

induce expression of atrogin-1 and MuRF1 under con-

ditions of energy stress [101,144]. Activation of AMPK

also leads to induction of some autophagy-related

genes encoding proteins such as LC3 and Bnip3.

Increased oxidative stress occurs during denervation

and hindlimb suspension. During these disuse condi-

tions, nNOS moves from the sarcolemma, where it is

bound to the dystrophinglycoprotein complex, to thecytosol. Free cytosolic nNOS induces oxidative stress

and enhances FoxO3-mediated transcription of atro-

gin-1 and MuRF1, thereby causing muscle loss [145].

Interestingly, the NF-jB pathway is not involved innNOS-mediated muscle atrophy [145]. Similarly, when

dihydropyridine receptor (DHPR) is reduced in adult

muscle by RNAi, muscle atrophy is triggered via

nNOS relocalization and FoxO3 activation [146].

However, in this latter setting, the genes up-regulated

by FoxO3 are those encoding the autophagy regulators

LC3, vacuolar protein sorting 34 (VPS34) and Bnip3

as well as the lysosomal enzyme cathepsin L. In

humans, the diaphragm of patients that are mechani-

cally ventilated undergoes rapid atrophy caused by

activation of proteolytic systems, including autophagy,

through Akt inhibition and FoxO1 induction [147].

Interestingly, oxidative stress is increased and therefore

contributes to FoxO activation in this example of dis-

use-mediated atrophy.

FoxO activity is also modulated by direct or indirect

actions of co-factors and by interaction with other

transcription factors. FoxOs have been found to inter-

act with PGC-1a, a critical co-factor for mitochondrialbiogenesis [148,149]. Maintaining high levels of PGC-

1a under catabolic conditions (either in transgenicmice or by transfecting adult myofibers) spares muscle

mass during denervation, fasting, heart failure, aging

and sarcopenia similar to the effect observed forexpression of constitutively active FoxO3 [5,150,151].

Similar beneficial effects were recently obtained by

over-expression of PGC-1b, a homolog of PGC-1a[152]. The positive action on muscle mass of these co-

factors is due to inhibition of autophagylysosomeand ubiquitinproteasome degradation. PGC-1a andPGC-1b reduce protein breakdown by inhibiting thetranscriptional activity of FoxO3 and NF-jB, but donot affect protein synthesis. Thus, these co-factors pre-

vent the excessive activation of proteolytic systems by

inhibiting the action of the pro-atrophy transcription

factors without perturbing the translational machinery.

We recently reported that the transcription factor

JunB blocks atrophy and promotes hypertrophy in

adult muscles [61]. Indeed, JunB blocks myofiber atro-

phy of denervated tibialis anterior muscles and cul-

tured myotubes induced by FoxO3 over-expression,

dexamethasone treatment or starvation. Under these

conditions, JunB prevents activation of atrogin-1 and

partially prevents activation of MuRF1, thereby reduc-

ing the increase in overall protein degradation induced

by activated FoxO3. Further analysis revealed that

JunB does not inhibit FoxO3-mediated activation of

the autophagylysosome system, but only ubiquitinproteasome degradation, by inhibiting atrogin-1 and

MuRF1 induction under catabolic conditions. In fact,

JunB directly binds FoxO3, thereby preventing its

recruitment to the promoters of key atrogenes. More-

over, JunB over-expression is sufficient to induce dra-

matic hypertrophy of myotubes and adult muscle.

These hypertrophic changes depend on increased pro-

tein synthesis, without affecting the basal rate of pro-

tein degradation. The growth-promoting effects

mediated by JunB in muscle resemble the effects of

inhibiting the TGFb pathway [33,34]. Indeed, JunBover-expression markedly suppresses myostatin expres-

sion in transfected myotubes and decreases the phos-

phorylation of Smad3, the transcription factor

downstream of the myostatinTGFb signaling path-way [61].

Inflammatory cytokines and NF-jB signaling

NF-jB transcription factors are expressed in skeletalmuscle and are activated by inflammatory cytokines,

particularly tumor necrosis factor a (TNFa). Indeed,inflammation is a potent trigger of muscle wasting and

cachexia [153]. NF-jB is maintained in the inactivestate by binding of a family of inhibitory proteins

called IjB. The increase in the TNFa level inducesactivation of an IjB kinase (IKKb) complex thatphosphorylates IjB, resulting in its ubiquitination andproteasomal degradation. This leads to nuclear trans-

location of NF-jB and activation of NF-jB-mediatedgene transcription [153].

Transgenic mice that over-express IKKb specificallyin muscle show severe muscle wasting that is mediated,

at least in part, by the ubiquitin ligase MuRF1, but

not by atrogin-1 [154]. In contrast, muscle-specific

inhibition of NF-jB by transgenic expression of a con-stitutively active IjB mutant does not induce an overtphenotype, but denervation atrophy is substantially

reduced [155]. Mice deficient for the p105/p50 subunit

of NF-jB are partially resistant to muscle atrophyinduced by hindlimb unloading [156]. However, one of



the effects of TNFa and pro-inflammatory cytokines isto induce insulin resistance and suppression of the

IGF1Akt pathway [157159]. Therefore, Akt phos-phorylation should always be monitored when NF-jBsignaling is altered, as Akt inhibition may substantially

contribute to muscle wasting. Indeed IKKb condi-tional knockout mice are resistant to muscle atrophy

but show activation of Akt [160]. The significance of

decreased muscle atrophy following IKKb ablationand the degree to which this effect is Akt-dependent

remains unclear. Nevertheless, these findings highlight

the relevance of the cross-talk between the two path-

ways, and future studies are required to elucidate the

respective contributions of the IKKbNF-jB andAktFoxO pathways to muscle atrophy.A recent study revealed an unexpected connection

between TNFa signaling and myogenin on MuRF1 andatrogin-1 expression: TNFa treatment causes up-regula-tion of myogenin, MuRF1 and atrogin-1. Interestingly,

a G protein-coupled receptor blocks TNFa-mediatedmyogenin up-regulation by activating Gai2 [161] andexpression of muscle-specific ubiquitin ligases. However,

the precise mechanisms of TNFa-mediated myogeninregulation, the interplay with Gai2 and the implicationsfor muscle wasting are still far from fully understood.

TNF-like weak inducer of apoptosis (TWEAK) is a

member of the TNF superfamily that was recently

found to induce muscle atrophy [158,162]. TWEAK

acts on responsive cells by binding to fibroblast growth

factor-inducible 14 (Fn14), a small cell-surface recep-

tor. Fn14 is up-regulated in denervated muscle, allow-

ing NF-jB activation and consequently MuRF1 (butnot atrogin-1) expression [162]. TWEAK knockout

mice display reduced atrophy after denervation, as well

as reduced NF-jB activation and MuRF1 expression.However, Fn14 does not increase under all conditions

of muscle atrophy; for instance, it is not induced by

dexamethasone treatment. Another important player

in NF-jB signaling is the ubiquitin ligase TRAF6,which is required for Fn14 up-regulation during fast-

ing [97]. As noted earlier, TRAF6 is also required for

activation of FoxO3 and AMPK in starved muscles

and for induction of the ubiquitinproteasome andautophagylysosome systems [97].The pro-inflammatory cytokines TNFa, IL-6 and

IL-1 also activate the Janus kinase/signal transducer

and activator of transcription (JAK/STAT) pathway.

Interestingly, sepsis and cancer induce STAT3 phos-

phorylation in muscles, and STAT3 inhibition spares

muscle mass in tumor-bearing mice [163]. Moreover,

over-expression of Stat3 is sufficient to induce muscle

atrophy and to up-regulate atrogin-1. However,

another recent study identified an unexpected role of

Stat3 in autophagy regulation. Stat3 has been reported

to block VPS34 expression, resulting in alteration of

assembly of the Vps34Beclin1Vps15Atg14 complex,and therefore autophagy inhibition and muscle degen-

eration [164]. This Stat3-dependent regulation of auto-

phagy occurs downstream of Fyn tyrosine kinase.

Other signaling pathways

Myostatin inhibition and its role in muscle growth has

been described above; however, the mechanism of myo-

statin activation and its role and capacity to trigger

muscle atrophy remain unclear. Myostatin activation

has been reported to induce massive [165], mild or no

atrophy at all [166,167]. However, in muscle cell cul-

tures, myostatin was reported to up-regulate essential

atrophy-related ubiquitin ligases. This regulation was

found to be FoxO-dependent and NF-jB-independent[168]. Importantly, myostatin expression is controlled

by FoxO1, supporting the concept that the myostatin

pathway synergizes with AktFoxO signaling [169]. Arecent study showed that inhibition of myostatin by sol-

uble ACVR2B prevents and fully reverses skeletal mus-

cle loss and atrophy of the heart in tumor-bearing

animals [63]. Such treatment dramatically prolongs the

survival of these animals, suggesting potential therapeu-

tic efficacy in patients with cancer cachexia. Reports

attempting to dissect the downstream signaling have

shown that Smad2 and Smad3 are the principle tran-

scription factors that mediate myostatins effects on

muscle mass [33,34,36,100]. However, as mentioned

above, specific transcriptional targets of Smad2 and

Smad3 are still unknown, and mechanisms of Smad-

dependent atrophy remain to be established.

Glucocorticoid levels are increased in many patholog-

ical conditions associated with muscle loss. Glucocorti-

coid treatment induces atrogin-1 and MuRF1

expression and muscle wasting in cell culture and in vivo

[16,77,88,127,170]. In contrast, adrenalectomy or treat-

ment with a glucocorticoid receptor antagonist attenu-

ate muscle loss in some diseases [170,171]. The

mechanisms of glucocorticoid-mediated muscle atrophy

were recently unraveled. Once in the nucleus, the gluco-

corticoid receptor activates expression of two target

genes, encoding REDD1 (regulated in development and

DNA damage responses 1) and KLF15 (Kruppel-like

factor 15) [172]. REDD1 inhibits mTOR activity by

sequestering 14-3-3 and increasing TSC1/2 activity.

Inhibition of mTOR is permissive for activation of an

atrophy program via KLF15. Indeed, mTOR activation

attenuates glucocorticoid-induced muscle atrophy.

KLF15 is a transcription factor that is involved in sev-

eral metabolic processes in skeletal muscle, for instance



up-regulation of branched-chain aminotransferase 2.

KLF15 participates in muscle catabolism via transcrip-

tional regulation of FoxO1, atrogin-1 and MuRF1.

Moreover, KLF15 negatively affects mTOR through

up-regulation of branched-chain aminotransferase 2,

which in turn induces branched-chain amino acid degra-

dation. Interestingly, FoxO1 and glucocorticoid recep-

tor cooperate to up-regulate MuRF1 expression [173].

Proteolysis-dependent regulation ofprotein synthesis

Synthesis and degradation of proteins are two pro-

cesses that are intimately connected. Indeed, most of

the above-mentioned pathways concomitantly regulate

both synthesis and degradation, such that when pro-

tein synthesis is induced, degradation is suppressed

and vice versa. However, this control appears to be a

compensatory mechanism to limit energy expenditure

for the production of novel proteins under catabolic

conditions. As mentioned above, in denervated mus-

cles, net protein synthesis is increased rather than

decreased compared to innervated muscles [8]. This is

because a proportion of the amino acids released from

protein breakdown stimulate protein synthesis via

mTOR, and, if this mechanism is blocked, muscle loss

is exacerbated [8]. The direct action of amino acids on

translation plays an important role in the rewiring of

protein synthesis during catabolic conditions, changing

the metabolism and expression of sarcomeric proteins

in order to optimize muscle homeostasis and perfor-

mance. An important example of amino acid-depen-

dent regulation of gene transcription during a

catabolic state has recently been described [174] for

lysosomal-dependent protein degradation. Nutrients,

especially free amino acids, are sensed by the mTOR

kinase, which then inhibits autophagy by blocking for-

mation of the Atg1/unc-51-like kinase 1 complex, an

important regulatory step for autophagy initiation.

The mTORC1 complex is therefore at the center of a

variety of cellular process such as protein synthesis,

autophagy, aging, mitochondrial function and energy

production. These various actions of mTORC1 are

exploited by its localization/recruitment to various cel-

lular compartments. For instance, the Rag GTPase

complex, which senses lysosomal amino acids, pro-

motes localization of mTORC1 to the lysosomal sur-

face. Accumulation of amino acids within the

lysosomal lumen generates an activating signal that is

transmitted to the Rag GTPases via vacuolar H+-

adenosine triphosphatase ATPase (v-ATPase), recruit-

ing mTORC1 to the lysosomes. This mTOR localiza-

tion initiates amino acid signaling and protein

synthesis [174] (Fig. 3). Concomitantly, mTOR also

inhibits transcription factor EB, a master regulator of

lysosome biogenesis [175]. Activation of mTORC1

induces phosphorylation and localization of transcrip-

tion factor EB at the lysosomal membrane, thus inhib-

iting its transcriptional activity [176]. These data

indicate that the content/activity of the lysosome

directly regulates lysosome biogenesis via an mTORtranscription factor EB axis. The implication of this

signaling as it relates specifically to muscle homeostasis

has yet to be investigated.

Conclusions and perspectives

Our understanding of the mechanisms that control

muscle growth and atrophy has greatly advanced dur-

ing the last ten years. Major milestones in this progress

have been identification of the transduction pathways

that mediate myostatin and IGF1 signals, in particular

the crucial role of Akt and its main downstream effec-

tors, mTOR, which controls protein synthesis, and

FoxO3, which controls protein degradation via the pro-

teasomal and autophagic/lysosomal systems. In addi-

tion, other pathways, such as SRF, have emerged as

potential players in regulation of muscle fiber size, and

the contribution of satellite cells has been the object of

intensive investigation. It is clear that there is no com-

mon mechanism that applies to all models of muscle

growth or wasting, and an important objective for

future studies will be to define the pathways that are

operative in the various situations. For example, auto-

phagy is highly up-regulated in skeletal muscles during

starvation, but its induction after denervation, when

the ubiquitinproteasome pathway is strongly acti-vated, is much less important. Satellite cell activation

may likewise vary in different models of muscle hyper-

trophy, and its contribution to the increase in muscle

fiber size remains to be established.

Dissection of the pathways that control muscle mass

and function will provide useful indications for the

development of drugs that are able to boost muscle

growth and prevent muscle wasting, a target that is now

being actively pursued in both academic and biotech-

nology/pharmaceutical research, and may have great

therapeutic importance for treatment of neuromuscular

diseases, systemic disorders, muscle disuse and aging.

Acknowledgements

Original work reported here is supported by the EC

FP7 Project MYOAGE (grant number 223576 to S.S.

and M.S.) and the European Research Council (grant

number 282310-MyoPHAGY to M.S.).



References

1 Aravamudan B, Mantilla CB, Zhan WZ & Sieck GC

(2006) Denervation effects on myonuclear domain size

of rat diaphragm fibers. J Appl Physiol 100,

16171622.

2 Li JB & Goldberg AL (1976) Effects of food

deprivation on protein synthesis and degradation in rat

skeletal muscles. Am J Physiol 231, 441448.

3 Goldberg AL & Goodman HM (1969) Relationship

between cortisone and muscle work in determining

muscle size. J Physiol 200, 667675.

4 Verheul AJ, Mantilla CB, Zhan WZ, Bernal M,

Dekhuijzen PN & Sieck GC (2004) Influence of

corticosteroids on myonuclear domain size in the rat

diaphragm muscle. J Appl Physiol 97, 17151722.

5 Sandri M, Lin J, Handschin C, Yang W, Arany ZP,

Lecker SH, Goldberg AL & Spiegelman BM (2006)

PGC-1a protects skeletal muscle from atrophy bysuppressing FoxO3 action and atrophy-specific gene

transcription. Proc Natl Acad Sci USA 103,

1626016265.

6 Goldberg AL (1969) Protein turnover in skeletal

muscle. I. Protein catabolism during work-induced

hypertrophy and growth induced with growth

hormone. J Biol Chem 244, 32173222.

7 Argadine HM, Hellyer NJ, Mantilla CB, Zhan WZ &

Sieck GC (2009) The effect of denervation on protein

synthesis and degradation in adult rat diaphragm

muscle. J Appl Physiol 107, 438444.

8 Quy PN, Kuma A, Pierre P & Mizushima N (2012)

Proteasome-dependent activation of mammalian target

of rapamycin complex 1 (mTORC1) is essential for

autophagy suppression and muscle remodeling

following denervation. J Biol Chem 288, 11251143.

8a Van der Meer SF, Jaspers RT & Degens H (2011) Is

the myonuclear domain size fixed? J Musculoskelet

Neuronal Interact 11, 286297.

9 Bruusgaard JC, Egner IM, Larsen TK, Dupre-

Aucouturier S, Desplanches D & Gundersen K (2012)

No change in myonuclear number during muscle

unloading and reloading. J Appl Physiol 113, 290296.

10 Bruusgaard JC & Gundersen K (2008) In vivo time-

lapse microscopy reveals no loss of murine myonuclei

during weeks of muscle atrophy. J Clin Invest 118,

14501457.

11 Ciciliot S & Schiaffino S (2010) Regeneration of

mammalian skeletal muscle. Basic mechanisms and

clinical implications. Curr Pharm Des 16, 906914.

12 Schiaffino S & Mammucari C (2011) Regulation of

skeletal muscle growth by the IGF1Akt/PKB pathway:insights from genetic models. Skelet Muscle 1, 4.

13 Mavalli MD, DiGirolamo DJ, Fan Y, Riddle RC,

Campbell KS, van Groen T, Frank SJ, Sperling MA,

Esser KA, Bamman MM, et al. (2010) Distinct growth

hormone receptor signaling modes regulate skeletal

muscle development and insulin sensitivity in mice.

J Clin Invest 120, 40074020.

14 Musaro A, McCullagh K, Paul A, Houghton L,

Dobrowolny G, Molinaro M, Barton ER, Sweeney

HL & Rosenthal N (2001) Localized Igf-1 transgene

expression sustains hypertrophy and regeneration in

senescent skeletal muscle. Nat Genet 27, 195200.

15 Murgia M, Serrano AL, Calabria E, Pallafacchina G,

Lomo T & Schiaffino S (2000) Ras is involved in

nerve-activity-dependent regulation of muscle genes.

Nat Cell Biol 2, 142147.

16 Bodine SC, Stitt TN, Gonzalez M, Kline WO, Stover

GL, Bauerlein R, Zlotchenko E, Scrimgeour A,

Lawrence JC, Glass DJ, et al. (2001) Akt/mTOR

pathway is a crucial regulator of skeletal muscle

hypertrophy and can prevent muscle atrophy in vivo.

Nat Cell Biol 3, 10141019.

17 Pallafacchina G, Calabria E, Serrano AL, Kalhovde

JM & Schiaffino S (2002) A protein

kinase B-dependent and rapamycin-sensitive pathway

controls skeletal muscle growth but not fiber type

specification. Proc Natl Acad Sci USA 99, 92139218.

18 Blaauw B, Canato M, Agatea L, Toniolo L,

Mammucari C, Masiero E, Abraham R, Sandri M,

Schiaffino S & Reggiani C (2009) Inducible activation

of Akt increases skeletal muscle mass and force without

satellite cell activation. FASEB J 23, 38963905.

19 Izumiya Y, Hopkins T, Morris C, Sato K, Zeng L,

Viereck J, Hamilton JA, Ouchi N, Lebrasseur NK &

Walsh K (2008) Fast/glycolytic muscle fiber growth

reduces fat mass and improves metabolic parameters

in obese mice. Cell Metab 7, 159172.

20 Lai KM, Gonzalez M, Poueymirou WT, Kline WO,

Na E, Zlotchenko E, Stitt TN, Economides AN,

Yancopoulos GD & Glass DJ (2004) Conditional

activation of Akt in adult skeletal muscle induces

rapid hypertrophy. Mol Cell Biol 24, 92959304.

21 Laplante M & Sabatini DM (2012) mTOR signaling in

growth control and disease. Cell 149, 274293.

22 Risson V, Mazelin L, Roceri M, Sanchez H,

Moncollin V, Corneloup C, Richard-Bulteau H,

Vignaud A, Baas D, Defour A, et al. (2009) Muscle

inactivation of mTOR causes metabolic and

dystrophin defects leading to severe myopathy. J Cell

Biol 187, 859874.

23 Bentzinger CF, Romanino K, Cloetta D, Lin S,

Mascarenhas JB, Oliveri F, Xia J, Casanova E, Costa

CF, Brink M, et al. (2008) Skeletal muscle-specific

ablation of raptor, but not of rictor, causes metabolic

changes and results in muscle dystrophy. Cell Metab 8,

411424.

24 Bodine SC (2006) mTOR signaling and the molecular

adaptation to resistance exercise. Med Sci Sports Exerc

38, 19501957.



25 Le Bacquer O, Petroulakis E, Paglialunga S, Poulin F,

Richard D, Cianflone K & Sonenberg N (2007)

Elevated sensitivity to diet-induced obesity and insulin

resistance in mice lacking 4E-BP1 and 4E-BP2. J Clin

Invest 117, 387396.

26 Mounier R, Lantier L, Leclerc J, Sotiropoulos A,

Foretz M & Viollet B (2011) Antagonistic control of

muscle cell size by AMPK and mTORC1. Cell Cycle

10, 26402646.

27 Goodman CA, Frey JW, Mabrey DM, Jacobs BL,

Lincoln HC, You JS & Hornberger TA (2011) The

role of skeletal muscle mTOR in the regulation of

mechanical load-induced growth. J Physiol 589,

54855501.

28 Latres E, Amini AR, Amini AA, Griffiths J, Martin

FJ, Wei Y, Lin HC, Yancopoulos GD & Glass DJ

(2005) Insulin-like growth factor-1 (IGF-1) inversely

regulates atrophy-induced genes via the

phosphatidylinositol 3-kinase/Akt/mammalian target

of rapamycin (PI3K/Akt/mTOR) pathway. J Biol

Chem 280, 27372744.

29 Cunningham JT, Rodgers JT, Arlow DH, Vazquez F,

Mootha VK & Puigserver P (2007) mTOR controls

mitochondrial oxidative function through a YY1

PGC-1a transcriptional complex. Nature 450, 736740.30 Blattler SM, Cunningham JT, Verdeguer F, Chim H,

Haas W, Liu H, Romanino K, Ruegg MA, Gygi SP,

Shi Y, et al. (2012) Yin Yang 1 deficiency in skeletal

muscle protects against rapamycin-induced diabetic-

like symptoms through activation of insulin/IGF

signaling. Cell Metab 15, 505517.

31 Lee SJ (2004) Regulation of muscle mass by

myostatin. Annu Rev Cell Dev Biol 20, 6186.

32 Taylor WE, Bhasin S, Artaza J, Byhower F, Azam M,

Willard DH Jr, Kull FC Jr & Gonzalez-Cadavid N

(2001) Myostatin inhibits cell proliferation and protein

synthesis in C2C12 muscle cells. Am J Physiol

Endocrinol Metab 280, E221E228.

33 Sartori R, Milan G, Patron M, Mammucari C,

Blaauw B, Abraham R & Sandri M (2009) Smad2 and

3 transcription factors control muscle mass in

adulthood. Am J Physiol Cell Physiol 296,

C1248C1257.

34 Trendelenburg AU, Meyer A, Rohner D, Boyle J,

Hatakeyama S & Glass DJ (2009) Myostatin reduces

Akt/TORC1/p70S6K signaling, inhibiting myoblast

differentiation and myotube size. Am J Physiol Cell

Physiol 296, C1258C1270.

35 Kalista S, Schakman O, Gilson H, Lause P,

Demeulder B, Bertrand L, Pende M & Thissen JP

(2011) The type 1 insulin-like growth factor receptor

(IGF-IR) pathway is mandatory for the follistatin-

induced skeletal muscle hypertrophy. Endocrinology

153, 241253.

36 Winbanks CE, Weeks KL, Thomson RE, Sepulveda

PV, Beyer C, Qian H, Chen JL, Allen JM, Lancaster

GI, Febbraio MA, et al. (2012) Follistatin-mediated

skeletal muscle hypertrophy is regulated by Smad3 and

mTOR independently of myostatin. J Cell Biol 197,

9971008.

37 Remy I, Montmarquette A & Michnick SW (2004)

PKB/Akt modulates TGF-b signalling through a directinteraction with Smad3. Nat Cell Biol 6, 358365.

38 Conery AR, Cao Y, Thompson EA, Townsend CM Jr,

Ko TC & Luo K (2004) Akt interacts directly with

Smad3 to regulate the sensitivity to TGF-b inducedapoptosis. Nat Cell Biol 6, 366372.

39 Li S, Czubryt MP, McAnally J, Bassel-Duby R,

Richardson JA, Wiebel FF, Nordheim A & Olson EN

(2005) Requirement for serum response factor for

skeletal muscle growth and maturation revealed by

tissue-specific gene deletion in mice. Proc Natl Acad

Sci USA 102, 10821087.

40 Charvet C, Houbron C, Parlakian A, Giordani J,

Lahoute C, Bertrand A, Sotiropoulos A, Renou L,

Schmitt A, Melki J, et al. (2006) New role for serum

response factor in postnatal skeletal muscle growth

and regeneration via the interleukin 4 and insulin-like

growth factor 1 pathways. Mol Cell Biol 26,

66646674.

41 Guerci A, Lahoute C, Hebrard S, Collard L,

Graindorge D, Favier M, Cagnard N, Batonnet-

Pichon S, Precigout G, Garcia L, et al. (2012) Srf-

dependent paracrine signals produced by myofibers

control satellite cell-mediated skeletal muscle

hypertrophy. Cell Metab 15, 2537.

42 Small EM, ORourke JR, Moresi V, Sutherland LB,

McAnally J, Gerard RD, Richardson JA & Olson EN

(2010) Regulation of PI3-kinase/Akt signaling by

muscle-enriched microRNA-486. Proc Natl Acad Sci

USA 107, 42184223.

43 White JP, Gao S, Puppa MJ, Sato S, Welle SL &

Carson JA (2012) Testosterone regulation of Akt/

mTORC1/FoxO3a signaling in skeletal muscle. Mol

Cell Endocrinol 365, 174186.

44 Koopman R, Gehrig SM, Leger B, Trieu J, Walrand

S, Murphy KT & Lynch GS (2010) Cellular

mechanisms underlying temporal changes in skeletal

muscle protein synthesis and breakdown during

chronic b-adrenoceptor stimulation in mice. J Physiol588, 48114823.

45 Kline WO, Panaro FJ, Yang H & Bodine SC (2007)

Rapamycin inhibits the growth and muscle-sparing

effects of clenbuterol. J Appl Physiol 102, 740747.

46 Le Grand F, Jones AE, Seale V, Scime A & Rudnicki

MA (2009) Wnt7a activates the planar cell polarity

pathway to drive the symmetric expansion of satellite

stem cells. Cell Stem Cell 4, 535547.



47 von Maltzahn J, Renaud JM, Parise G & Rudnicki

MA (2011) Wnt7a treatment ameliorates muscular

dystrophy. Proc Natl Acad Sci USA 109,

2061420619.

48 Ito N, Ruegg UT, Kudo A, Miyagoe-Suzuki Y &

Takeda S (2012) Activation of calcium signaling

through Trpv1 by nNOS and peroxynitrite as a key

trigger of skeletal muscle hypertrophy. Nat Med 19,

101106.

49 Hornberger TA, Chu WK, Mak YW, Hsiung JW,

Huang SA & Chien S (2006) The role of

phospholipase D and phosphatidic acid in the

mechanical activation of mTOR signaling in skeletal

muscle. Proc Natl Acad Sci USA 103, 47414746.

50 Ruas JL, White JP, Rao RR, Kleiner S, Brannan KT,

Harrison BC, Greene NP, Wu J, Estall JL, Irving BA,

et al. (2012) A PGC-1a isoform induced by resistancetraining regulates skeletal muscle hypertrophy. Cell

151, 13191331.

50a Puigserver P, Wu Z, Park CW, Graves R, Wright M

& Spiegelman BM (1998) A cold-inducible coactivator

of nuclear receptors linked to adaptive thermogenesis.

Cell 92, 829839.

51 Pallafacchina G, Blaauw B & Schiaffino S (2013) Role

of satellite cells in muscle growth and maintenance of

muscle mass. Nutr Metab Cardiovasc Dis, in press.

52 Abmayr SM & Pavlath GK (2012) Myoblast fusion:

lessons from flies and mice. Development 139, 641656.

53 Horsley V, Jansen KM, Mills ST & Pavlath GK

(2003) IL-4 acts as a myoblast recruitment factor

during mammalian muscle growth. Cell 113,

483494.

54 Serrano AL, Baeza-Raja B, Perdiguero E, Jardi M &

Munoz-Canoves P (2008) Interleukin-6 is an essential

regulator of satellite cell-mediated skeletal muscle

hypertrophy. Cell Metab 7, 3344.

55 Moss FP & Leblond CP (1971) Satellite cells as the

source of nuclei in muscles of growing rats. Anat Rec

170, 421435.

56 White RB, Bierinx AS, Gnocchi VF & Zammit PS

(2010) Dynamics of muscle fibre growth during

postnatal mouse development. BMC Dev Biol 10, 21.

57 Saclier M, Cuvellier S, Magnan M, Mounier R &

Chazaud B (2013) Monocyte/macrophage interactions

with myogenic precursor cells during skeletal muscle

regeneration. FEBS J, doi:10.1111/febs.12166.

58 Maltin CA & Delday MI (1992) Satellite cells in

innervated and denervated muscles treated with

clenbuterol. Muscle Nerve 15, 919925.

59 Kadi F (2008) Cellular and molecular mechanisms

responsible for the action of testosterone on human

skeletal muscle. A basis for illegal performance

enhancement. Br J Pharmacol 154, 522528.

60 Jackson JR, Mula J, Kirby TJ, Fry CS, Lee JD, Ubele

MF, Campbell KS, McCarthy JJ, Peterson CA &

Dupont-Versteegden EE (2012) Satellite cell depletion

does not inhibit adult skeletal muscle regrowth

following unloading-induced atrophy. Am J Physiol

Cell Physiol 303, C854C861.

61 Raffaello A, Milan G, Masiero E, Carnio S, Lee D,

Lanfranchi G, Goldberg AL & Sandri M (2010) JunB

transcription factor maintains skeletal muscle mass

and promotes hypertrophy. J Cell Biol 191, 101113.

62 Amthor H, Otto A, Vulin A, Rochat A, Dumonceaux

J, Garcia L, Mouisel E, Hourde C, Macharia R,

Friedrichs M, et al. (2009) Muscle hypertrophy driven

by myostatin blockade does not require stem/

precursor-cell activity. Proc Natl Acad Sci USA 106,

74797484.

63 Zhou X, Wang JL, Lu J, Song Y, Kwak KS, Jiao Q,

Rosenfeld R, Chen Q, Boone T, Simonet WS, et al.

(2010) Reversal of cancer cachexia and muscle wasting

by ActRIIB antagonism leads to prolonged survival.

Cell 142, 531543.

64 Lee SJ, Huynh TV, Lee YS, Sebald SM, Wilcox-

Adelman SA, Iwamori N, Lepper C, Matzuk MM &

Fan CM (2012) Role of satellite cells versus myofibers

in muscle hypertrophy induced by inhibition of the

myostatin/activin signaling pathway. Proc Natl Acad

Sci USA 109, E2353E2360.

65 Schiaffino S, Bormioli SP & Aloisi M (1972) Cell

proliferation in rat skeletal muscle during early stages

of compensatory hypertrophy. Virchows Arch B Cell

Pathol 11, 268273.

66 Snow MH (1990) Satellite cell response in rat soleus

muscle undergoing hypertrophy due to surgical

ablation of synergists. Anat Rec 227, 437446.

67 Bruusgaard JC, Johansen IB, Egner IM, Rana ZA &

Gundersen K (2010) Myonuclei acquired by overload

exercise precede hypertrophy and are not lost on

detraining. Proc Natl Acad Sci USA 107, 1511115116.

68 Crameri RM, Langberg H, Magnusson P, Jensen CH,

Schroder HD, Olesen JL, Suetta C, Teisner B & Kjaer

M (2004) Changes in satellite cells in human skeletal

muscle after a single bout of high intensity exercise.

J Physiol 558, 333340.

69 Barton-Davis ER, Shoturma DI & Sweeney HL (1999)

Contribution of satellite cells to IGF-I induced

hypertrophy of skeletal muscle. Acta Physiol Scand

167, 301305.

70 Adams GR, Caiozzo VJ, Haddad F & Baldwin KM

(2002) Cellular and molecular responses to increased

skeletal muscle loading after irradiation. Am J Physiol

Cell Physiol 283, C1182C1195.

71 McCarthy JJ, Mula J, Miyazaki M, Erfani R,

Garrison K, Farooqui AB, Srikuea R, Lawson BA,

Grimes B, Keller C, et al. (2011) Effective fiber

hypertrophy in satellite cell-depleted skeletal muscle.

Development 138, 36573666.



72 Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez

L, Clarke BA, Poueymirou WT, Panaro FJ, Na E,

Dharmarajan K, et al. (2001) Identification of

ubiquitin ligases required for skeletal muscle atrophy.

Science 294, 17041708.

73 Gomes MD, Lecker SH, Jagoe RT, Navon A &

Goldberg AL (2001) Atrogin-1, a muscle-specific F-

box protein highly expressed during muscle atrophy.

Proc Natl Acad Sci USA 98, 1444014445.

74 Lecker SH, Jagoe RT, Gilbert A, Gomes M, Baracos

V, Bailey J, Price SR, Mitch WE & Goldberg AL

(2004) Multiple types of skeletal muscle atrophy

involve a common program of changes in gene

expression. FASEB J 18, 3951.

75 Sacheck JM, Hyatt JP, Raffaello A, Jagoe RT, Roy

RR, Edgerton VR, Lecker SH & Goldberg AL (2007)

Rapid disuse and denervation atrophy involve

transcriptional changes similar to those of muscle

wasting during systemic diseases. FASEB J 21,

140155.

76 Sandri M (2008) Signaling in muscle atrophy and

hypertrophy. Physiology (Bethesda) 23, 160170.

77 Sandri M, Sandri C, Gilbert A, Skurk C, Calabria E,

Picard A, Walsh K, Schiaffino S, Lecker SH &

Goldberg AL (2004) Foxo transcription factors induce

543fc6bb0cf21227a11b7b45

Documents

skeletal muscle growth

adult skeletal muscle

muscle fiber size

skeletal muscle mass

kda protein

protein fn14

rates of protein degradation

tothe loss of muscle