Muscle Growth, Repair and Preservation: A Mechanistic Approach

Full text

1
Muscle Growth, Repair and Preservation: A Mechanistic Approach
Robert M. Erskine1,2 and Hans Degens3,4
1School of Sport and Exercise Sciences, Liverpool John Moores University, Liverpool, United
Kingdom; 2Institute of Sport, Exercise and Health, Division of Surgery and Interventional Sciences,
University College London, London, UK; 3School of Healthcare Science, Manchester Metropolitan
University, Manchester, United Kingdom; 4Institute of Sport Science and Innovations, Lithuanian
Sports University, Kaunas, Lithuania.
Address correspondence to: R.M. Erskine, PhD; School of Sport and Exercise
Sciences, Liverpool John Moores University, Byrom Street, Liverpool L3 3AF,
United Kingdom; Tel: +44 (0)151 904 6256; Fax: +44 (0)151 904 6284; Email:
R.M.Erskine@ljmu.ac.uk
Running title: Muscle Growth, Repair and Preservation

2
SUMMARY
Resistance exercise, amino acid ingestion and an anabolic hormone environment all
have the capacity to elevate muscle protein synthesis (MPS), while a catabolic
hormone environment, such as elevated pro-inflammatory cytokines as seen during
disuse, aging, and conditions such as cancer and AIDS, can cause an increase in
muscle protein degradation (MPD). When the rate of MPS exceeds that of MPD there
is a positive net protein balance (NPB) and over a prolonged period of time this
results in accretion of contractile material and muscle growth, or hypertrophy. In
contrast, when NPB is chronically negative, muscle atrophy occurs, i.e. muscle size
decreases. Various signaling pathways within the muscle fiber appear to play a crucial
role in the adaptive processes, and understanding how these pathways can be
modulated will help the design of therapies to prevent or reverse muscle atrophy in a
host of muscle wasting conditions.
Key words: skeletal muscle – protein synthesis – hypertrophy – atrophy – IGF-I –
mTOR – cytokine – TNF- – interleukin – myostatin

3
INTRODUCTION
Skeletal muscle comprises numerous bundles of long, thin, multinucleated cells called
muscle fibers, each containing a multitude of myofibrils. Each myofibril is composed
of myofilaments (comprising the contractile proteins, actin and myosin) and a variety
of structural proteins, all arranged in a regular configuration throughout the length of
the myofibril, so as to form a series of contractile components, or sarcomeres. The
maximum force that can be generated by a muscle fiber is proportional to the number
of sarcomeres arranged in parallel, or fiber cross-sectional area (CSA), and ultimately
the CSA of the whole muscle (Jones, Rutherford, Parker, 1989). Therefore, there is a
strong relationship between whole muscle CSA and maximum isometric force
measured in vivo (Bamman, Newcomer, Larson-Meyer, Weinsier, Hunter, 2000;
Fukunaga et al., 2001; Kanehisa, Ikegawa, Fukunaga, 1994).
Based on the relationship between muscle size and force generating capacity (Erskine,
Fletcher, Folland, 2014), it is not surprising that an increase in muscle size following
resistance training (RT) is accompanied by an increase in maximal muscle force
(Erskine, Jones, Williams, Stewart, Degens, 2010b; Jones, Rutherford, 1987; Narici et
al., 1996). Not only can this enhance the athletic performance of an individual but it
can also reduce the elevated risk of falling and bone fracture in older people that is
among other factors attributable to sarcopenia (the age-related loss of muscle mass).
The question thus arises as to the mechanisms underlying overload-induced muscle
hypertrophy.
A multitude of signaling molecules within the muscle fiber are thought to play an
integral role in stimulating muscle protein synthesis (MPS) and degradation (MPD). If

4
there is a positive net protein balance (NPB), i.e. when the rate of MPS exceeds that
of MPD, the amount of contractile material will increase, enabling the muscle to
hypertrophy and generate more force. Conversely, when NPB is negative, the muscle
will decrease in size, or atrophy, and become weaker. This chapter will explore the
specific signaling pathways involved in MPS and MPD, which help to explain how
skeletal muscle adapts to overload, disuse, ageing and muscle-wasting diseases.
Furthermore, strategies used to preserve or maintain muscle mass during periods of
disuse and wasting, such as RT and nutritional interventions, will be discussed.
In addition to the mechanisms underlying muscle growth and atrophy, there is still
more to be learned about the systems associated with repair following exercise-
induced muscle damage. Several studies have reported that disruption of the
cytoskeletal structure of muscle fibers is accompanied by impairment of muscular
function following damage-inducing exercise (Baumert, Lake, Stewart, Drust,
Erskine, 2016; Friden, Sjostrom, Ekblom, 1983; Lieber, Shah, Friden, 2002). As well
as structural damage to the sarcomere, eccentric exercise can cause raised intracellular
calcium ion (Ca2+) levels (Belcastro, Albisser, Littlejohn, 1996), decreased muscle
force production (Clarkson, Sayers, 1999; Friden et al., 1983), an increase in serum
levels of muscle-specific proteins (Ebbeling, Clarkson, 1989), an increase in muscle
specific inflammation (MacIntyre, Reid, McKenzie, 1995), an increase in proteolytic
enzyme activity (Evans, Cannon, 1991), and a delayed-onset muscle soreness
(MacIntyre et al., 1995). The ultimate repair of the muscle requires the activation of
satellite cells and in this chapter we will consider the various MPD systems and the
role of satellite cells thought to play a major role in muscle damage and repair
following exercise.

5
MUSCLE GROWTH
While prenatal muscle growth is largely the result of muscle fiber formation, postnatal
maturational muscle growth and that in response to RT is almost entirely attributable
to fiber hypertrophy. Prenatal myogenesis, i.e. the formation of muscle fibers during
embryonic development, involves the proliferation, migration, differentiation and
fusion of muscle precursor cells to form post-mitotic multinucleated myotubes.
Postnatal skeletal muscle growth is accompanied by an increase in the number of
myonuclei per muscle fiber (Delhaas, van der Meer, Schaart, Degens, Drost, In Press)
that requires the activation of muscle stem cells, or satellite cells (located at the basal
lamina that surrounds the muscle fiber), which proliferate and fuse with existing
muscle fibers (Jacquemin, Furling, Bigot, Butler-Browne, Mouly, 2004). Once fully
mature, skeletal muscle growth, or hypertrophy, is dependent upon a positive NPB,
i.e. MPS must be greater than MPD (Chesley, MacDougall, Tarnopolsky, Atkinson,
Smith, 1992; Phillips, Tipton, Aarsland, Wolf, Wolfe, 1997), a process that is driven
by an increase in the rate of MPS (Kumar, Atherton, Smith, Rennie, 2009). This leads
to an accretion of myofibrillar proteins and an increase in muscle fiber CSA, which in
turn leads to an increase in the overall CSA of the muscle, thus enabling more force to
be produced. Resistance exercise, i.e. overloading the muscle, has been shown to
increase MPS (Chesley et al., 1992; Phillips et al., 1997), and chronic resistance
exercise, i.e. RT performed over many weeks, is a potent stimulus for skeletal muscle
hypertrophy and strength gains (Erskine et al., 2010b; Jones et al., 1987; Narici et al.,
1996). However, exactly how overloading the muscle leads to a positive NPB and
therefore an increase in muscle size has yet to be fully elucidated. It is thought that the
process necessary for inducing muscle hypertrophy involves a myriad of molecules

6
within the muscle fiber that form signaling cascades, eventually culminating in
increased MPS and/or decreased MPD. Here we will discuss how insulin-like growth
factor-I (IGF-I), mechanosensors, and amino acids might activate these specific
signaling pathways that lead to MPS and ultimately to muscle growth, or hypertrophy.
The role of IGF-I in muscle growth
IGF-I is produced by the liver and skeletal muscle and thus acts on muscle fibers in an
endocrine and autocrine/paracrine manner (Goldspink, 1999; Stewart, Rotwein,
1996). This growth factor appears to play an integral role in activating a specific
signaling pathway within the muscle fiber that stimulates MPS (Bodine et al., 2001;
Rommel et al., 2001). The local production and release of IGF-I during muscle
contraction (DeVol, Rotwein, Sadow, Novakofski, Bechtel, 1990) activates this
signaling cascade by binding to its receptor, located in the sarcolemma. This causes
autophosphorylation of the insulin receptor substrate (IRS1) and subsequent
phosphorylation of down-stream molecules within this signaling pathway, which
includes phosphatidylinositol-3 kinase (PI3K), protein kinase-B (PKB or Akt), the
mammalian target of rapamycin complex 1 (mTORC1), 70-kDa ribosomal S6 protein
kinase (p70S6K), and eukaryotic initiation factor 4E binding protein (4E-BP) (Fig. 1).
In fact, p70S6K activation is related to gains in skeletal muscle mass following RT,
both in rats (Baar, Esser, 1999) and humans (Terzis et al., 2008), with the increase
occurring mainly in type II fibers (Koopman, Zorenc, Gransier, Cameron-Smith, van
Loon, 2006). Together, these studies implicate mTORC1 and p70S6K as principal
downstream mediators of IGF-I stimulation of skeletal muscle growth.
[FIGURE 1 NEAR HERE]

7
There is evidence that IGF-I produced in skeletal muscle is more important for
developmental and exercise-induced muscle growth than IGF-I produced by the liver.
This is indicated by the greater muscle mass in transgenic mice over-expressing IGF-I
in skeletal muscle compared to wild-type mice (Coleman et al., 1995; Musaro et al.,
2001) despite normal serum IGF-I levels (Coleman et al., 1995). Furthermore, low
systemic IGF-I levels in liver-specific IGF-I knockout mice does not affect muscle
size (Ohlsson et al., 2009; Yakar et al., 1999). Also in young adult men, elevated
levels of circulating IGF-I do not influence MPS following an acute bout of resistance
exercise (West et al., 2009) or muscle hypertrophy in response to RT (West et al.,
2010). Although in rat skeletal muscle, local IGF-I gene expression increases
proportionately to the progressive increase in external load (DeVol et al., 1990), it is
equivocal whether this occurs in human muscle (Bamman et al., 2001; Bickel et al.,
2005; Bickel, Slade, Haddad, Adams, Dudley, 2003; Hameed, Orrell, Cobbold,
Goldspink, Harridge, 2003; Petrella, Kim, Cross, Kosek, Bamman, 2006; Psilander,
Damsgaard, Pilegaard, 2003). Some of this controversy might be explained by the
elevated expression of two isoforms of the Igf1 gene in animal skeletal muscle in
response to mechanical stimulation (McKoy et al., 1999; Yang, Alnaqeeb, Simpson,
Goldspink, 1996). Thus, at least two IGF-I isoforms exist: (i) IGF-IEa, which is
similar to the hepatic endocrine isoform, and (ii) the less abundant IGF-IEb (in rats)
or IGF-IEc (in humans), otherwise known as mechanical growth factor (MGF).
However, it is not always clear which IGF-I isoform has been measured in the muscle
(Bamman et al., 2001). Furthermore, the age of the participants also influences the
findings, with MGF increasing in young but not old people following an exercise
bout. Interestingly, IGF-IEa does not appear to change in young or older people

8
following resistance exercise (Hameed et al., 2003), despite its apparent hypertrophic
effect (Musaro et al., 2001).
Skeletal muscle-derived IGF-I does not appear to be the only regulator of adult
muscle mass and function, as unloading induces skeletal muscle atrophy in mice
whose muscles over-express IGF-I (Criswell et al., 1998), and overload induces
hypertrophy even in transgenic mice, which express a dominant negative IGF-I
receptor in skeletal muscle (Spangenburg, Le Roith, Ward, Bodine, 2008). Also in
older people, enhanced muscle strength can be attained following RT without a
significant change in muscle IGF-I gene expression (Taaffe, Jin, Vu, Hoffman,
Marcus, 1996). Thus, it appears that for the development of hypertrophy in adult
muscles, loading is more important than alterations in local and systemic IGF-I levels.
This fits the notion that activation of mTORC1 and p70S6K can also occur
independently of PI3K activation following muscle overload (Hornberger, Sukhija,
Wang, Chien, 2007).
The role of mechanosensors in muscle growth
The process that couples the mechanical forces during a muscle contraction with cell
signaling and ultimately protein synthesis is called mechanotransduction. It has been
shown in rats that passive stretch induces an increase in the expression of myogenic
regulatory factors (Kamikawa, Ikeda, Harada, Ohwatashi, Yoshida, 2013) that may
well underlie the muscle fiber atrophy seen after passive stretching (Coutinho,
DeLuca, Salvini, Vidal, 2006). These responses are probably at least partly mediated
by stretch-activated channels (SACs), which are calcium (Ca2+) and sodium
permeable channels that increase their open probability (the fraction of time spent in

9
the open state) in response to mechanical loading of the sarcolemma (Franco,
Lansman, 1990a; Franco, Lansman, 1990b; Guharay, Sachs, 1984). It has been
proposed that SACs function as mechanosensors by allowing an influx of Ca2+ into
the muscle fiber (Yeung et al., 2005) following mechanical changes in the
sarcolemma, which activate mTORC1 (Gulati et al., 2008), leading to an increase in
MPS (Kameyama, Etlinger, 1979). Correspondingly, inhibition of SACs by
streptomycin reduces skeletal muscle hypertrophy in response to mechanical overload
(Butterfield, Best, 2009; Spangenburg, McBride, 2006) via attenuation of mTORC1
and p70S6K activation (Spangenburg et al., 2006).
Other mechanosensors might exist in the form of costameres, intra-sarcolemmal
protein complexes that are circumferentially aligned along the length of the muscle
fiber (Pardo, Siliciano, Craig, 1983). Costameres mechanically link peripheral
myofibrils via the Z-disks to the sarcolemma (Fig. 2), thus maintaining the integrity of
the muscle fiber during contraction and relaxation (Pardo et al., 1983). An individual
costamere contains many proteins arranged in a complex structure (Ervasti, 2003;
Patel, Lieber, 1997), which comprises two different laminin receptors, a
dystrophin/glycoprotein complex and an integrin-associated complex, which are
localised in the sarcolemma and bound to intra and extra-cellular structural proteins
(Fig. 2). Thus, the force-producing contractile material is connected to the basal
membrane and ultimately to adjacent muscle fibers (Morris, Fulton, 1994; Patel et al.,
1997; Rybakova, Patel, Ervasti, 2000).
[FIGURE 2 NEAR HERE]

10
Costameres are receptive to mechanical, electrical and chemical stimuli (Ervasti,
2003). Indeed, mechanical tension is essential in regulating costameric protein
expression, stability and organization, with talin and vinculin, for instance, being up-
regulated in response to muscle contraction (Tidball, Spencer, Wehling, Lavergne,
1999). Regular contractions, as experienced during RT, increase the expression of
costameric proteins, such as desmin (Woolstenhulme, Conlee, Drummond, Stites,
Parcell, 2006), alpha-1-syntrophin and dystrophin (Kosek, Bamman, 2008) in
humans, while focal adhesion kinase (FAK) and paxillin activity are increased in
stretch-induced hypertrophied avian skeletal muscle (Fluck, Carson, Gordon,
Ziemiecki, Booth, 1999). The forces exerted on both the intracellular contractile
proteins and the basal membrane during periods of loading are required to cause
binding of basal membrane laminin to the receptors on the  and  integrins and on
the dystrophin/glycoprotein complex (Fluck, Ziemiecki, Billeter, Muntener, 2002).
Interaction between integrins and the extra-cellular matrix causes rapid
phosphorylation of FAK (Cary, Guan, 1999), which subsequently activates p70S6K
independently of Akt (Durieux et al., 2009; Klossner, Durieux, Freyssenet, Flueck,
2009). This probably occurs via the phosphorylation and, thus, inactivation of
tuberous sclerosis complex 2 (Gan, Yoo, Guan, 2006; Malik, Parsons, 1996), thus
activating mTORC1 as shown in Fig. 1.
The role of amino acids in muscle growth
Both resistance exercise (Biolo, Maggi, Williams, Tipton, Wolfe, 1995; Phillips et al.,
1997) and amino acid/protein ingestion (Tang, Moore, Kujbida, Tarnopolsky, Phillips,
2009; Yang et al., 2012b) stimulate MPS independently, while a combination of the
two augments MPS even further (Biolo, Tipton, Klein, Wolfe, 1997; Tipton,

11
Ferrando, Phillips, Doyle, Wolfe, 1999). Both stimuli cause an increase in mTORC1
activation (Apro, Blomstrand, 2010; Moore, Atherton, Rennie, Tarnopolsky, Phillips,
2011) but it is unclear whether they stimulate MPS via different signaling pathways,
or whether the combination of the two stimulates the same pathway more than either
stimulant on its own. It is thought that amino acids cause Rag GTPases to interact
with raptor (a regulatory protein associated with mTORC1), leading to the
translocation of mTORC1 to the lysosomal membrane, where Rheb (a Ras GTPase)
activates mTORC1 (Kim, Guan, 2009; Sancak et al., 2010; Sancak et al., 2008), as
shown in Fig. 1. Of the essential amino acids (EAAs), the branched-chain amino acids
(BCAAs: isoleucine, leucine, and valine), and particularly leucine, are the most potent
stimulators of the mTORC1 signaling pathway (Anthony et al., 2000). Leucine
supplementation stimulates muscle protein accretion in cultured cells (Haegens,
Schols, van Essen, van Loon, Langen, 2012) and can also reduce MPD in healthy men
(Nair, Schwartz, Welle, 1992), and it is possible that the action of leucine occurs via
its metabolite, β-hydroxy-β-methylbutyrate (HMB) (Aversa et al., 2011; Pimentel et
al., 2011).
The timing of amino acid ingestion appears crucial for an optimal anabolic response
to a single bout of resistance exercise: ingesting amino acids immediately before an
exercise bout promotes a greater increase in MPS compared to ingestion immediately
after the bout (Tipton et al., 2001). This effect was attributed to an increased blood
flow during exercise, and therefore an increased delivery of amino acids to the active
muscle when they were ingested prior to exercise (Tipton et al., 2001). As well as the
timing, the amount of protein ingested is integral in producing an optimal anabolic
environment following resistance exercise (Moore et al., 2009; Witard et al., 2014;

12
Yang et al., 2012a). For example, the MPS dose response to ingested protein after a
single bout of resistance exercise in healthy young men is saturated at 20 g protein,
and any additional ingested protein is simply oxidized (Moore et al., 2009; Witard et
al., 2014). This suggests that if the rate of protein ingestion after resistance exercise
exceeds the rate at which it can be incorporated into the muscle, the excess protein is
not used for MPS. This dose-response relationship seems to be altered with age, as in
older men increased rates of MPS were found when participants ingested 40 g protein
following a resistance exercise bout (Yang et al., 2012a). Therefore, older muscle
appears to be less sensitive to amino acids, which has been termed ‘anabolic
resistance’, something that may be attributable to impaired ribosome genesis in older
muscle (Chaillou, 2017). Finally, the type and quality of the ingested protein appears
to be important when it comes to MPS (Tang et al., 2009; Yang et al., 2012b).
Following a single bout of resistance exercise and the ingestion of whey, soy or
casein, each containing 10 g EAA, larger increases in blood EAA, BCAA, and leucine
concentrations were found following the ingestion of whey compared to either soy or
casein (Tang et al., 2009), suggesting a greater availability of these amino acids for
protein synthesis following whey protein ingestion. This may be a reflection of the
different rate of protein digestion and absorption of amino acids between the protein
types (Boirie et al., 1997; Dangin et al., 2001; Dangin et al., 2003) and explain why
MPS was greater following ingestion of whey compared to casein, both at rest and
after exercise (Tang et al., 2009). In older muscle, the rate of MPS appears to be
greater with whey than soy protein ingestion following resistance exercise (Yang et
al., 2012b), which could be due to the ~28% greater leucine content in whey versus
soy protein (Drummond, Rasmussen, 2008) as well as differences in digestion and
absorption rate.

13
There is therefore striking evidence to support the acute effects of amino acid
ingestion and resistance exercise on MPS via their independent and complementary
effects on mTORC1 activation. It is also well known that repeated bouts of resistance
exercise over a prolonged period of time, i.e. a RT program, leads to gains in both
muscle size and strength (Erskine et al., 2010b; Jones et al., 1987; Narici et al., 1996).
Therefore, the amplification of the anabolic environment within the muscle seen with
the combination of both amino acid/protein ingestion and resistance exercise (Biolo et
al., 1997; Tipton et al., 1999) suggests that RT with protein supplementation should
confer greater gains in skeletal muscle size and strength than RT alone. However, the
evidence for protein supplementation enhancing the increases in muscle size and
strength following longer term RT programs in young (Erskine, Fletcher, Hanson,
Folland, 2012; Hartman et al., 2007) and older (Candow et al., 2008; Verdijk et al.,
2009b) individuals is equivocal. The controversy surrounding the longer-term RT
studies could be due to methodological differences/limitations between studies. For
example, considerable inter-individual variability exists in the response to RT
(Erskine, Jones, Williams, Stewart, Degens, 2010a; Hubal et al., 2005) and yet many
studies have used small sample sizes (Godard, Williamson, Trappe, 2002; Hulmi et
al., 2009; Willoughby, Stout, Wilborn, 2007) that may have limited the statistical
power required to detect an influence of protein supplementation. Different measures
of muscle hypertrophy may also compound this discrepancy. For example, some
studies have determined muscle thickness using ultrasonography (Candow et al.,
2008; Vieillevoye, Poortmans, Duchateau, Carpentier, 2010) or whole body fat-free
mass assessed via dual-energy X-ray absorptiometry (Hartman et al., 2007), while
others have used magnetic resonance imaging to provide a more precise assessment of

14
muscle size, but still found no effect of protein supplementation on muscle
hypertrophy following RT (Coburn et al., 2006; Erskine et al., 2012; Holm et al.,
2008; Hulmi et al., 2009). There are, however, circumstances where protein
supplementation may have a beneficial effect on muscle hypertrophy and strength
gains. For example, whole body RT (incorporating multiple muscle groups rather than
an individual muscle) could create a requirement for an increase in exogenous protein
(due to a greater absolute MPD) that might not be satisfied by habitual protein intake
alone. This might be particularly beneficial in the early phase of a RT program when
MPS and MPD are likely to be higher than towards the end (Hartman, Moore,
Phillips, 2006; Phillips, Tipton, Ferrando, Wolfe, 1999). In addition, older individuals
need to ingest at least twice as much protein (40 g) to maximally stimulate MPS
(Moore et al., 2015; Yang et al., 2012a) compared to the 20 g dose in younger people
(Moore et al., 2009; Witard et al., 2014), which may reflect an ‘anabolic resistance’ in
old age. Therefore, previous studies that have not shown a beneficial effect of protein
supplementation on muscle size and strength gains in older people may have been
administering suboptimal amounts and types of protein per RT session. In frail elderly
individuals, whose protein requirements are probably higher than in old, healthy
people, more recent evidence supports the combination of 60 g/day milk protein
supplementation and resistance exercise (twice weekly for six months) in increasing
total leg lean mass composition over resistance training alone (Tieland et al., 2012).
MUSCLE ATROPHY
Skeletal muscle atrophy is known to occur in response to disuse and numerous
chronic conditions, such as cancer, AIDS, and senile sarcopenia (the age-related loss
of muscle mass). Here we will focus on the mechanisms underlying sarcopenia, which

15
is the major cause of muscle weakness in older individuals (Evans, 1995). Although
the causes of sarcopenia are not fully understood, disuse, chronic systemic
inflammation and neuropathic changes leading to motoneuron death are thought to
play an integral role (Degens, 2010). Motoneuron death results in denervation of
muscle fibers, and ultimately the loss of muscle fibers (hypoplasia). Selective atrophy
of type II fibers (Lexell, Taylor, Sjostrom, 1988) and a decrease in the proportion of
type II fibers (Jakobsson, Borg, Edstrom, Grimby, 1988; Larsson, 1983; Larsson,
Sjodin, Karlsson, 1978) are thought to be caused by denervation accompanied by
reinnervation of these fibers by axonal sprouting from adjacent slow-twitch motor
units (Brooks, Faulkner, 1994; Faulkner, Larkin, Claflin, Brooks, 2007).
Chronic low-grade inflammation and sarcopenia
Physiological aging is associated with chronic low-grade inflammation, a condition
that has been termed ‘inflammaging’ (Franceschi et al., 2000). Inflammaging is
characterized by elevated serum levels of pro-inflammatory cytokines such as
interleukin 1 (IL-1), IL-6 and tumor necrosis factor- (TNF-), as well as acute
phase proteins such as C-reactive protein, or CRP (Bartlett et al., 2012; Erskine et al.,
2017; Franceschi et al., 2000), and increased circulating levels are associated with
lower muscle mass and weakness in old age (Erskine et al., 2017; Visser et al., 2002).
Furthermore, the levels of cytokines that counteract the inflammatory state, such as
IL-10, are reduced with age (Bartlett et al., 2012; Lio et al., 2002). The pro-
inflammatory cytokines, IL-1, IL-6 and TNF- are produced by both skeletal muscle
fibers and adipose cells, and are therefore also members of the adipokine family. As
we age, we accumulate more adipose tissue, which is deposited in the subcutaneous,
visceral and intramuscular regions, and it is particularly the visceral fat that appears to

16
contribute to the inflammatory environment (Pedersen, 2009).
TNF- induces the production of reactive oxygen species (ROS), altering vascular
permeability, which leads to leukocyte infiltration of the muscle fiber (Evans et al.,
1991) and further ROS generation by the leucocytes. This release of ROS also
activates NF-κB via degradation of I-κB (Fig. 1), which results in the increased
expression of key enzymes of the ubiquitin-proteasome MPD system (Li, Schwartz,
Waddell, Holloway, Reid, 1998). In addition, TNF- interferes with satellite cell
differentiation and therefore muscle growth and regeneration in old age by reducing
the expression of myogenic regulatory factors (MRFs) (Degens, 2010). The MRFs are
a family of muscle-specific transcription factors (MyoD, myogenin, MRF4 and myf-
5) that regulate the transition from proliferation to differentiation of the satellite cell
(Langen et al., 2004; Szalay, Razga, Duda, 1997). The TNF--induced activation of
NF-κB results in a loss of MyoD mRNA (Guttridge, Mayo, Madrid, Wang, Baldwin,
2000) and, via activation of the ubiquitin–proteasome pathway (UPP) (Reid, Li, 2001;
Saini, Al-Shanti, Faulkner, Stewart, 2008), breakdown of MyoD and myogenin. Thus,
part of the attenuated hypertrophic response in elderly compared to younger muscle
(Welle, Totterman, Thornton, 1996) could be due to a decrease in MRFs, as seen in
overload-induced hypertrophy in older rats (Alway, Degens, Krishnamurthy, Smith,
2002a). TNF- also stimulates the release of proteolytic enzymes, such as lysozymes
(e.g. cathepsin-B) from neutrophils (Farges et al., 2002), which are thought to
contribute to the MPD process (Friden, Kjorell, Thornell, 1984; Kasperek, Snider,
1985), but the main action of the pro-inflammatory cytokines in muscle atrophy is
thought to occur via the UPP.

17
It has been suggested that the UPP cannot breakdown intact sarcomeres, so additional
mechanisms are proposed to be involved (Solomon, Baracos, Sarraf, Goldberg, 1998).
For example, the activation of caspase-3 is thought to lead to cleavage of the
myofilaments, actin and myosin, which are then degraded by the UPP (Du et al.,
2004; Lee et al., 2004; Ottenheijm et al., 2006). The UPP involves numerous
enzymes, or ligases, that regulate the ubiquitination (the coupling of ubiquitin to
protein substrates), so that the ‘tagged’ protein fragments can be identified by the
proteasome for the final step of MPD (DeMartino, Ordway, 1998). These ubiquitin
ligases may also degrade MyoD and inhibit subsequent satellite cell activation and
differentiation (see above), thus exacerbating the effects of MPD on muscle size by
impairing satellite cell associated muscle growth. Expression of two genes that encode
the E3 ubiquitin ligases, muscle-specific atrophy F box (MAFbx, also known as
Atrogin-1) and muscle RING Finger 1 (MuRF1), has been shown to increase during
different types of muscle atrophy (Gomes, Lecker, Jagoe, Navon, Goldberg, 2001;
Lecker et al., 2004). The structure and function of the UPP will be discussed in more
detail, below, with regard to muscle damage and repair following exercise.
The role of myostatin in muscle atrophy
Myostatin, otherwise known as growth differentiation factor-8 (GDF-8), is part of the
transforming growth factor β superfamily and is produced in skeletal muscle. The role
of myostatin as a negative regulator of muscle mass has been demonstrated by
knocking out the gdf-8 gene in mice, which leads to a 2-3 fold increase in skeletal
muscle mass (McPherron, Lawler, Lee, 1997). Conversely, administering myostatin to
wild-type mice induces substantial muscle wasting (Zimmers et al., 2002). Further
examples of myostatin’s regulatory effect can be seen in bovine (McPherron, Lee,

18
1997) and human (Schuelke et al., 2004) cases, where mutation of the gdf -8 gene
leads to a reduction in myostatin production and considerably enlarged skeletal
muscles.
Once bound to the activin IIB receptor (ActRIIB), a signaling cascade is activated that
leads to MPD. Key signaling proteins in this pathway include SMAD 2 and 3 (Sartori
et al., 2009), which form a complex with SMAD 4 that then translocates to the
nucleus where it targets genes encoding MRFs (Rodino-Klapac et al., 2009), and
inhibits differentiation via the reduction of MyoD expression (Langley et al., 2002)
(Fig. 1). In addition, myostatin reduces Akt/ mTORC1/p70S6K signaling (Amirouche
et al., 2009; Trendelenburg et al., 2009) (Fig. 1) and is associated with smaller
myotube size (Trendelenburg et al., 2009). Accordingly, the inhibition of myostatin in
mature mice leads to increased activation of p70S6K, ribosomal protein S6 and skeletal
muscle MPS (Welle, Burgess, Mehta, 2009). Myostatin appears to not only inhibit
satellite cell differentiation and MPS, but also to induce the expression of atrogenes
(genes associated with muscle atrophy) via activation of the p38 mitogen-activated
protein (MAP) kinase, Erk1/2, Wnt and c-Jun N-terminal kinase (JNK) signaling
pathways (Huang et al., 2007; Philip, Lu, Gao, 2005; Yang et al., 2006) (Fig. 1). This
is in accord with the observation that myostatin induces cachexia by activating the
UPP, i.e. phosphorylating the ubiquitin E3 ligases MAFbx and MuRF1, via FOXO1
activation rather than via the NF-κB pathway (McFarlane et al., 2006). However,
myostatin-induced atrophy persists despite inhibiting the expression of the two E3
ligases (Trendelenburg et al., 2009). Therefore, it is likely that myostatin negatively
regulates muscle growth via multiple pathways (Fig. 1). A lower expression of
myostatin may therefore help to maintain muscle mass at old age, a situation reflected

19
by the attenuated loss of muscle mass and regenerative capacity in old myostatin-null
mice compared to age-matched wild-type mice (Siriett et al., 2006).
Combating sarcopenia
RT has been shown to increase muscle size and strength in old (Reeves, Narici,
Maganaris, 2004), very old (Harridge, Kryger, Stensgaard, 1999) and frail (Fiatarone
et al., 1994) individuals. This beneficial effect of RT is attributable to an increase in
MPS in the atrophied muscle (Kimball, O'Malley, Anthony, Crozier, Jefferson, 2004)
and a reduction in MPD as a result of a reduced MAFbx and MuRF-1 gene expression
(Jones et al., 2004; Mascher et al., 2008). A reduction in atrogene expression can be
realized by the ability of phosphorylated Akt to block FoxO1, which would suppress
the transcription of MuRF1 and MAFbx (Sandri et al., 2004), as shown in Fig. 1. In
addition to the Akt/FoxO-1 pathway, mTORC1 also blocks MuRF1 and MAFbx
transcription (Sandri et al., 2004). Therefore, while a degree of MPD is required for
muscle remodeling, RT appears to reverse atrophy via the inhibiting effect of
Akt/mTORC1 on MPD and the positive effect of mTORC1 activation on MPS (Apro
et al., 2010; Moore et al., 2011), thus resulting in net protein synthesis (albeit to a
lesser extent in elderly compared to younger muscle).
The apparent ‘anabolic resistance’ to RT in older compared to young muscle (Kimball
et al., 2004; Welle et al., 1996) may not be due to an age-related reduction in the
mechanosensitivity of the mTORC1 signaling pathway (Hornberger, Mateja, Chin,
Andrews, Esser, 2005), although others do see a reduction in the translational
signaling during overload (Thomson, Gordon, 2006). It could therefore be that the

20
rates of transcription and translation are reduced during overload, which may be a
consequence of impaired ribosome biogenesis in older muscle (Chaillou, 2017).
Another factor that might be considered is the proposed requirement of satellite cell
recruitment for the development of hypertrophy. An impaired satellite cell recruitment
would then result in impaired hypertrophy and part of the problem might be a decline
in satellite cell number (Shefer, Van de Mark, Richardson, Yablonka-Reuveni, 2006),
particularly in type II muscle fibers of older people (Verdijk et al., 2007). Yet,
paradoxically, older muscle appears to have an increased regenerative drive and
protein synthesis that is more pronounced the more severe the sarcopenia. For
instance, muscle mass is lower despite higher p70S6K activation and MPS in old
compared to young adult rats (Kimball et al., 2004). IGF-I appears to play a key role
in the activation and proliferation of satellite cells (Scime, Rudnicki, 2006), while
differentiation is regulated by MRFs (Langen et al., 2004; Szalay et al., 1997).
However, despite an increased regenerative drive as reflected by elevated IGF-I
expression (Edstrom, Ulfhake, 2005) and MRF mRNA expression (Alway, Degens,
Lowe, Krishnamurthy, 2002b) in old rat muscle, MRF protein levels are reduced
(Alway et al., 2002b). This reduction might be caused by a concomitant increase in Id
protein expression (Alway et al., 2002b), which inhibits MRF expression and DNA
binding capacity. The result is that during an hypertrophic stimulus, satellite cell
activation and proliferation can occur in older rat skeletal muscle but with limited
differentiation (Edstrom et al., 2005). In elderly human skeletal muscle, however, the
capacity for satellite cells to proliferate and differentiate in response to RT does not
appear to be diminished (Mackey et al., 2007; Verdijk et al., 2009a). It should be
noted, however, that the relative age in the human studies was less than that in the rat

21
studies and it may be that beyond a given age in humans, the differentiation of
satellite cells may also be diminished, particularly when associated with chronic low-
grade systemic inflammation.
Although satellite cells are generally considered to play an important role in muscle
hypertrophy, the conditional ablation of satellite cells by tamoxifen in Pax7-DTA
mice did not attenuate the development of 100% hypertrophy induced by overload
(McCarthy et al., 2011). It may well be that the microcirculation is crucial for the
development of hypertrophy as it has been found that in genetically modified mice
with muscle hypertrophy (due to myostatin knock out) and high-oxidative fibers
(overexpression of oestrogen-related receptor gamma) the regenerative capacity was
normal despite a lower satellite cell content, but higher capillary density (Omairi et
al., 2016). Also in aged mice, the blunted hypertrophic response was associated not
only associated with a lower satellite cell content (Ballak et al., 2015), but also with
impaired angiogenesis (Ballak et al., 2016). These data suggest that impaired
angiogenesis and/or reductions in the capillarization, that would result in an impaired
delivery of substrates, of the muscle may underlie the impaired regenerative capacity
and anabolic resistance in old age
Extra stimulation of the mTORC1 pathway may overcome the anabolic blunting. As
discussed above, older people need to ingest more protein than younger individuals to
stimulate maximal MPS (Moore et al., 2015; Moore et al., 2009; Volpi, Mittendorfer,
Rasmussen, Wolfe, 2000; Yang et al., 2012a). The greater activation of the mTORC1
pathway when combining RT with protein/amino acid ingestion may inhibit MPD and
augment MPS, thereby improving the hypertrophic response. The observation that

22
BCAA administration attenuates the loss of body mass in mice bearing a cachexia-
inducing tumor (Eley, Russell, Tisdale, 2007) is promising and suggests that it may
also enhance the hypertrophic response in this condition. Furthermore, HMB has been
shown to attenuate the reduction in MPS in rodents following the administration of a
cachectic stimulant (Aversa et al., 2011; Eley, Russell, Baxter, Mukerji, Tisdale,
2007).
In addition to RT and amino acid supplementation, various pharmaceutical therapies
have been proposed to combat sarcopenia. Supplementation of the anabolic steroid,
testosterone, augments muscle mass in older men, healthy hypogonadal men, older
men with low testosterone levels, and men with chronic illness and low testosterone
levels (Bhasin et al., 2006). It is thought that testosterone can reverse sarcopenia by
suppressing skeletal muscle myostatin expression, while simultaneously stimulating
the Akt pathway (Kovacheva, Hikim, Shen, Sinha, Sinha-Hikim, 2010) to increase
MPS and decrease MPD. Furthermore, administration of a myostatin antagonist has
led to satellite cell activation, increased MyoD protein expression, and greater muscle
regeneration after injury in old murine skeletal muscle (Siriett et al., 2007).
MUSCLE DAMAGE AND REPAIR
In normal skeletal muscle, cytoskeletal proteins act as a framework that keeps the
myofibrils aligned in a lateral position by connecting the Z-disks to one another and to
the sarcolemma (Friden et al., 1984; Friden et al., 1983). Following eccentric exercise,
Z-disk streaming (disturbance of Z-disk configuration) and misalignment of the
myofibrils is a common characteristic (Friden et al., 1984). Eccentric contractions are
defined as contractions where muscles lengthen as they exert force and generally

23
result in more muscle damage than concentric contractions (Clarkson, Hubal, 2002). It
has been suggested that this is due to fewer motor units being recruited during
eccentric exercise, leading to a smaller CSA of muscle being activated than during a
concentric contraction at the same load (Enoka, 1996). It has also been demonstrated
that the extent of muscle damage is due to strain (the change in length) rather than the
amount of force generated by the muscle (Lieber, Friden, 1993; Lieber, Friden, 1999).
Eccentric exercise not only causes alterations in the cytoskeletal structure, but also
increases in the activity of proteolytic enzymes (Arthur, Booker, Belcastro, 1999;
Kasperek et al., 1985; Stupka, Tarnopolsky, Yardley, Phillips, 2001). The positive
correlation between the proteolytic enzyme activity and a rise in serum concentrations
of muscle-specific proteins, e.g. creatine kinase (CK), post exercise (Arthur et al.,
1999; Kasperek et al., 1985; Stupka et al., 2001), suggests that the degree of
activation of the proteolytic machinery is related to the degree of muscle damage.
Therefore, it is feasible that the activity of these proteolytic enzymes may be required
for the remodeling of skeletal muscle in response to exercise, where the regulated
degradation of cellular proteins (Ordway, Neufer, Chin, DeMartino, 2000) may be a
pre-requisite for subsequent adaptive repair and growth.
There are three main systems that contribute to the controlled MPD following muscle
damage; 1) the release of calpain, a non-lysosomal, Ca2+-dependent neutral protease
that mediates the dismantling of myofibrils (Belcastro, Shewchuk, Raj, 1998), 2) the
inflammatory response, which includes lysosomal proteolysis (Farges et al., 2002),
and 3) the ATP-dependent UPP, which coordinates the demolition of protein
fragments liberated by the aforementioned degradation systems (DeMartino et al.,

24
1998). Recent findings suggest that myostatin is also implicated in the MPD process
following damaging muscle contractions (Ochi et al., 2010).
The calpain protein degradation system
Calpain is a multidomain protein composed of two subunits, a catalytic 80-kDa
subunit and a regulatory 30-kDa subunit (Suzuki, Sorimachi, Yoshizawa, Kinbara,
Ishiura, 1995). In skeletal muscle, three homologous isozymes of calpain with
different Ca2+ sensitivities have been identified (DeMartino et al., 1998):

-calpain
(active at micromolar Ca2+ concentrations), m-calpain (active at millimolar Ca2+
concentrations), and n-calpain (requiring very high Ca2+ concentrations). It appears
that, although the

- and m-calpain 80-kDa subunits are quite different, both have
similar binding domains; the proteolytic site of a cysteine proteinase, the calpastatin
(an endogenous inhibitor of calpain activity) binding domain, and the Ca2+-binding
domain (Belcastro et al., 1996). The 30-kDa subunit is extremely hydrophobic, which
may help to act as an anchor to the membrane proteins (Belcastro et al., 1996). While
there is evidence to suggest calpain is localized and activated at or around the
sarcolemma, thus targeting the membrane-associated proteins (Belcastro et al., 1998),
others have demonstrated that calpain also targets Z-disk proteins, such as desmin and
-actinin (Goll, Dayton, Singh, Robson, 1991). The action of other proteolytic
complexes, including lysosomal enzymes and the UPP, may have a part to play in
MPD immediately after damaging eccentric muscle contractions, but as their activity
does not peak until later in the muscle damage/repair process (Belcastro et al., 1998;
Kasperek et al., 1985), it is more likely that calpain and/or mechanical stress is the
initial effector of cytoskeletal protein breakdown.

25
Calpain is activated by raised intracellular [Ca2+] (Belcastro et al., 1998). Initial
mechanical damage to the sarcoplasmic reticulum (SR) and muscle plasma membrane
caused by eccentric muscle contractions could lead to SR vacuolization and an
increase in intracellular [Ca2+] (Clarkson et al., 2002; Warren, Hayes, Lowe,
Armstrong, 1993). The intracellular [Ca2+] could rise further following an increased
open probability of SACs as a consequence of increased strain on the skeletal muscle
fibers during forced lengthening (Lieber et al., 1993). It is thought that the activation
of calpain is pivotal in the breakdown of cytoskeletal proteins, including desmin and
-actinin, rather than mechanical stress applied to the “over-stretched” sarcomeres
during eccentric contractions per se (Belcastro et al., 1998). The activity of calpain is
not only dependent on the intracellular [Ca2+] but also on the concentration of its
inhibitor, calpastatin, and condition of degradable substrates, i.e. the ultrastructural
proteins. To become fully active, calpain undergoes autolysis into its subunits. It is
likely that the influx of excess Ca2+ into the muscle fiber (via the SACs, SR calcium
channels and sarcolemmal lesions) binds to the specific domain on the 80-kDa calpain
subunit, thereby inhibiting calpastatin. Once calpain is free of calpastatin, it may
begin autolysis and/or bind to its substrate (with the help of Ca2+) and begin the
process of MPD (Belcastro et al., 1996). A positive relationship between calpain
activity and neutrophil accumulation within skeletal muscle after exercise suggests
that the calpain-degraded protein fragments act as chemoattractants, thus localizing
leukocytes to the site of muscle damage (Belcastro et al., 1998; Raj, Booker,
Belcastro, 1998).
The inflammatory response
Exercise-induced damage to muscle fibers elicits an inflammatory response that

26
results in movement of fluid, plasma proteins and leukocytes to the site of injury
(Clarkson et al., 2002). Leukocytes have the ability to break down intracellular
proteins with the aid of lysosomal enzymes (Friden et al., 1984), but exactly how the
inflammatory response regulates MPD and muscle repair following eccentric exercise
is not entirely clear. However, the purpose of the post-exercise-induced inflammatory
response is to promote clearance of damaged muscle tissue and prepare the muscle for
repair (MacIntyre et al., 1995), a process that is sub-classified into acute and
secondary inflammation.
The acute phase response in skeletal muscle begins with the ‘complement system’
when fragments from the damaged fiber(s) serve as chemoattractants, luring
leukocytes to the injured area (Belcastro et al., 1998; Evans et al., 1991). As a
consequence there is an accumulation of neutrophils, the histological hallmark of
acute inflammation (MacIntyre et al., 1995), in and around the site of injury, which
peaks around 4 hours after exercise-induced damage has occurred (Evans et al.,
1991). The accumulation of neutrophils has been reported to be more significant after
eccentric than concentric exercise, and is most likely related to the degree of damage
incurred (Evans et al., 1991). Pro-inflammatory cytokines, such as IL-1, IL-6, TNF-,
and acute phase proteins, e.g. CRP, act as mediators of inflammatory reactions. TNF-
 induces the production of ROS, altering vascular permeability, which leads to
leukocyte infiltration into the muscle fiber (Evans et al., 1991). TNF- also stimulates
the release of cytoxic factors from neutrophils, such as lysozymes and ROS (Friden et
al., 1984; MacIntyre et al., 1995), which are responsible for at least a part of the MPD
process following exercise-induced damage (Friden et al., 1984; Kasperek et al.,
1985).

27
It may take up to seven days to see a significant infiltration of monocytes (precursors
to macrophages) within the damaged muscle fiber (Evans et al., 1991), which carry
out further phagocytic activity inside the muscle fiber. Furthermore, considerable
increases in the quantity of lipofuscin granules (generally considered to be the
indigestible residue of lysosomal degradation) in sore muscles three days after
exercise suggests that lysozyme activity plays a major part in the secondary
inflammatory process (Farges et al., 2002; Friden, 1984). The role of the
inflammatory response in muscle regeneration is therefore thought to be the further
breakdown of damaged muscle proteins via lysozymes, the engulfing of protein
fragments by macrophages and the activation of the UPP by the pro-inflammatory
cytokines released from the neutrophils. These cytokines simultaneously stimulate the
proliferation of satellite cells, crucial for the regeneration of the damaged area (Chen,
Jin, Li, 2007).
The ubiquitin-proteasome pathway
This pathway is recognized as the major non-lysosomal complex responsible for the
degradation of cellular proteins. The UPP has received much attention due to its
involvement in cellular processes, where protein degradation is a key regulatory or
adaptive event (Attaix et al., 1998; Attaix, Combaret, Pouch, Taillandier, 2001;
Ciechanover, 1994). There are two types of ubiquitin in human skeletal muscle: free
and conjugated (Thompson, Scordilis, 1994). In its free state ubiquitin is a normal
component of the non-stressed muscle fiber but it also forms complexes, or
conjugations, with abnormal proteins and then returns to its free state (Fig. 3). The
conjugation of ubiquitin with denatured proteins within the muscle fiber “tags” these

28
proteins for recognition by a non-lysosomal protease to be subsequently degraded in a
process that requires ATP (Attaix et al., 1998). The UPP, therefore, consists of two
major components that represent the system’s functionally distinct parts. Ubiquitin is
the element that covalently binds to the protein due to be broken down, while the 26S
proteasome, a large protease complex, catalyses the degradation of the ubiquitin-
tagged proteins (DeMartino et al., 1998).
[FIGURE 3 NEAR HERE]
The cellular proteins are selected for degradation by the attachment of multiple
molecules of ubiquitin, or a polyubiquitin chain, which is built by repeated cycles of
conjugation via the action of E1, E2, and E3 conjugating enzymes (Fig. 3). Ubiquitin
is initially activated in the presence of ATP by the ubiquitin-activating enzyme, E1,
which then transfers ubiquitin to E2, one of the ubiquitin-conjugating enzymes. E2
then binds the ubiquitin molecule to the protein substrate, which is selected for
tagging by E3 (Attaix et al., 2001). The 26S proteasome is able to discriminate
between ubiquitinated and non-ubiquitinated proteins, and rapidly degrades the
polyubiquitinated proteins, deriving the energy for this process from ATP hydrolysis
(Hershko, Ciechanover, 1998).
The 26S proteasome is composed of a 20S proteasome and two 19S (PA700)
regulatory modules (Attaix et al., 2001; Ciechanover, 1994; DeMartino et al., 1998).
The 20S proteasome is the proteolytic core, containing multiple catalytic sites. PA700
binds to each end of the proteasome cylinder and elicits ATPase activity in order to
unfold and/or translocate the ubiquitinated proteins to the catalytic sites within the

29
20S proteasome (Fig. 3). PA700 is also thought to be responsible for disassembling
the polyubiquitinated chain, a process requiring its isopeptidase activity.
The total amount of ubiquitin found in skeletal muscle is muscle fiber-type specific,
with a greater abundance of ubiquitin found in type I fibers (Riley et al., 1992).
Furthermore, a three to seven times higher density of conjugated ubiquitin was found
at the Z-discs than anywhere else in the muscle fiber, which suggests that, like
calpain, ubiquitin targets the cytoskeletal proteins of muscle fibers. One main
difference between the two systems, however, is that calpain is activated a lot earlier
than the ubiquitin-proteasome pathway (Feasson et al., 2002; Stupka et al., 2001).
Furthermore, the action of the UPP may be prolonged post-exercise in order to
increase the intracellular concentration of amino acids (Tipton, Wolfe, 1998), which
would stimulate MPS via mTORC1 activation.
Repair following exercise-induced muscle damage
Following the orderly demolition of damaged/cleaved muscle proteins (via the
aforementioned MPD systems) in response to eccentric contractions, the damaged
muscle fibers need to undergo repair. The activation, proliferation and fusion of
satellite cells with damaged muscle fibers, and the subsequent differentiation into
myoblasts, are crucial for this repair process (McCarthy et al., 2011; Petrella et al.,
2006). In fact, it has been shown that while the hypertrophic response maybe
maintained in the absence of satellite cell recruitment, recovery from damage was
severely impaired under these conditions (McCarthy et al., 2011).

30
As previously discussed, IGF-I and MRFs are integral in the activation and
differentiation of satellite cells (Langen et al., 2004; Scime et al., 2006; Szalay et al.,
1997), which probably explains why skeletal muscle IGF-I and MRF expression are
increased following stretch-induced damage (Bickel et al., 2005; Petrella et al., 2006;
Yang, Alnaqeeb, Simpson, Goldspink, 1997; Yang, Creer, Jemiolo, Trappe, 2005). To
repair the muscle, satellite cells fuse with damaged fibers and differentiate into
myonuclei, but may even form new fibers in the case of complete fiber necrosis. The
orderly proliferation and subsequent differentiation is crucial for optimal repair. For
the initial proliferation of satellite cells the inflammatory environment is beneficial,
but this inflammation must be transient to allow the cells to differentiate (Pelosi et al.,
2007). Therefore, it may be that chronic low-grade systemic inflammation, e.g. during
ageing, may underlie the delay in muscle regeneration (Langen et al., 2006).
SUMMARY
Skeletal muscle is able to hypertrophy in response to a variety of anabolic stimuli,
which include resistance exercise, amino acid ingestion, and an increase in IGF-I
expression. All these stimuli are able to activate the mTORC1 signaling pathway,
which stimulates MPS and inhibits MPD. When the rate of MPS exceeds MPD, there
is a positive net protein balance (NPB) and an accretion of contractile material occurs,
leading to muscle hypertrophy and an increase in strength. Inducing muscle
hypertrophy can have beneficial effects on individuals suffering from cachectic
conditions, such as cancer, AIDS, and sarcopenia, where muscle atrophy can have
devastating effects on an individual’s quality of life. Muscle atrophy occurs when
there is a negative NPB, i.e. when the rate of MPD is greater than MPS. There are a
number of stimuli that have been associated with muscle atrophy, including

31
chronically elevated levels of pro-inflammatory cytokines (e.g. IL-1, IL-6 and TNF-
), a reduction in IGF-I and increased expression of myostatin. Furthermore,
strenuous unaccustomed exercise can cause mechanical damage to the muscle, which
activates MPD systems, including calpain, inflammation and the ubiquitin-proteasome
protein degradation pathway. Damaged proteins within the muscle fiber are broken
down, resulting in an increased intracellular amino acid concentration, which in turn
activates mTORC1 and increases MPS, thus helping to repair the muscle. Elevated
local IGF-I and MRF expression facilitates the repair process by activating satellite
cells and enabling fusion with existing fibers. Many of the molecular signaling
pathways associated with muscle hypertrophy, atrophy and repair have been
identified. However, there is still much to be learned about these pathways, and
understanding them may help us to prevent or reverse muscle atrophy associated with
a host of muscle wasting conditions.

32
REFERENCES
Alway, S.E., Degens, H., Krishnamurthy, G., Smith, C.A. 2002a. Potential role for Id
myogenic repressors in apoptosis and attenuation of hypertrophy in muscles of aged
rats. Am J Physiol Cell Physiol 283, C66-76.
Alway, S.E., Degens, H., Lowe, D.A., Krishnamurthy, G. 2002b. Increased myogenic
repressor Id mRNA and protein levels in hindlimb muscles of aged rats. Am J Physiol
Regul Integr Comp Physiol 282, R411-422.
Amirouche, A., Durieux, A.C., Banzet, S., Koulmann, N., Bonnefoy, R., Mouret, C.,
Bigard, X., Peinnequin, A., Freyssenet, D. 2009. Down-regulation of Akt/mammalian
target of rapamycin signaling pathway in response to myostatin overexpression in
skeletal muscle. Endocrinology 150, 286-294.
Anthony, J.C., Yoshizawa, F., Anthony, T.G., Vary, T.C., Jefferson, L.S., Kimball,
S.R. 2000. Leucine stimulates translation initiation in skeletal muscle of
postabsorptive rats via a rapamycin-sensitive pathway. J Nutr 130, 2413-2419.
Apro, W., Blomstrand, E. 2010. Influence of supplementation with branched-chain
amino acids in combination with resistance exercise on p70S6 kinase phosphorylation
in resting and exercising human skeletal muscle. Acta Physiol (Oxf) 200, 237-248.
Arthur, G.D., Booker, T.S., Belcastro, A.N. 1999. Exercise promotes a subcellular
redistribution of calcium-stimulated protease activity in striated muscle. Can J Physiol
Pharmacol 77, 42-47.
Attaix, D., Aurousseau, E., Combaret, L., Kee, A., Larbaud, D., Ralliere, C.,
Souweine, B., Taillandier, D., Tilignac, T. 1998. Ubiquitin-proteasome-dependent
proteolysis in skeletal muscle. Reprod Nutr Dev 38, 153-165.
Attaix, D., Combaret, L., Pouch, M.N., Taillandier, D. 2001. Regulation of
proteolysis. Curr Opin Clin Nutr Metab Care 4, 45-49.
Aversa, Z., Bonetto, A., Costelli, P., Minero, V.G., Penna, F., Baccino, F.M., Lucia,
S., Rossi Fanelli, F., Muscaritoli, M. 2011. beta-hydroxy-beta-methylbutyrate (HMB)
attenuates muscle and body weight loss in experimental cancer cachexia. Int J Oncol
38, 713-720.
Baar, K., Esser, K. 1999. Phosphorylation of p70(S6k) correlates with increased
skeletal muscle mass following resistance exercise. Am J Physiol 276, C120-127.
Ballak, S.B., Buse-Pot, T., Harding, P.J., Yap, M.H., Deldicque, L., de Haan, A.,
Jaspers, R.T., Degens, H. 2016. Blunted angiogenesis and hypertrophy are associated
with increased fatigue resistance and unchanged aerobic capacity in old overloaded
mouse muscle. Age (Dordr) 38, 39.
Ballak, S.B., Jaspers, R.T., Deldicque, L., Chalil, S., Peters, E.L., de Haan, A.,
Degens, H. 2015. Blunted hypertrophic response in old mouse muscle is associated
with a lower satellite cell density and is not alleviated by resveratrol. Exp Gerontol
62, 23-31.

33
Bamman, M.M., Newcomer, B.R., Larson-Meyer, D.E., Weinsier, R.L., Hunter, G.R.
2000. Evaluation of the strength-size relationship in vivo using various muscle size
indices. Med Sci Sports Exerc 32, 1307-1313.
Bamman, M.M., Shipp, J.R., Jiang, J., Gower, B.A., Hunter, G.R., Goodman, A.,
McLafferty, C.L., Jr., Urban, R.J. 2001. Mechanical load increases muscle IGF-I and
androgen receptor mRNA concentrations in humans. Am J Physiol Endocrinol Metab
280, E383-390.
Bartlett, D.B., Firth, C.M., Phillips, A.C., Moss, P., Baylis, D., Syddall, H., Sayer,
A.A., Cooper, C., Lord, J.M. 2012. The age-related increase in low-grade systemic
inflammation (Inflammaging) is not driven by cytomegalovirus infection. Aging Cell
11, 912-915.
Baumert, P., Lake, M.J., Stewart, C.E., Drust, B., Erskine, R.M. 2016. Genetic
variation and exercise-induced muscle damage: implications for athletic performance,
injury and ageing. Eur J Appl Physiol 116, 1595-1625.
Belcastro, A.N., Albisser, T.A., Littlejohn, B. 1996. Role of calcium-activated neutral
protease (calpain) with diet and exercise. Can J Appl Physiol 21, 328-346.
Belcastro, A.N., Shewchuk, L.D., Raj, D.A. 1998. Exercise-induced muscle injury: a
calpain hypothesis. Mol Cell Biochem 179, 135-145.
Bhasin, S., Calof, O.M., Storer, T.W., Lee, M.L., Mazer, N.A., Jasuja, R., Montori,
V.M., Gao, W., Dalton, J.T. 2006. Drug insight: Testosterone and selective androgen
receptor modulators as anabolic therapies for chronic illness and aging. Nat Clin Pract
Endocrinol Metab 2, 146-159.
Bickel, C.S., Slade, J., Mahoney, E., Haddad, F., Dudley, G.A., Adams, G.R. 2005.
Time course of molecular responses of human skeletal muscle to acute bouts of
resistance exercise. J Appl Physiol 98, 482-488.
Bickel, C.S., Slade, J.M., Haddad, F., Adams, G.R., Dudley, G.A. 2003. Acute
molecular responses of skeletal muscle to resistance exercise in able-bodied and
spinal cord-injured subjects. J Appl Physiol 94, 2255-2262.
Biolo, G., Maggi, S.P., Williams, B.D., Tipton, K.D., Wolfe, R.R. 1995. Increased
rates of muscle protein turnover and amino acid transport after resistance exercise in
humans. Am J Physiol 268, E514-520.
Biolo, G., Tipton, K.D., Klein, S., Wolfe, R.R. 1997. An abundant supply of amino
acids enhances the metabolic effect of exercise on muscle protein. Am J Physiol 273,
E122-129.
Bodine, S.C., Stitt, T.N., Gonzalez, M., Kline, W.O., Stover, G.L., Bauerlein, R.,
Zlotchenko, E., Scrimgeour, A., Lawrence, J.C., Glass, D.J., Yancopoulos, G.D. 2001.
Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can
prevent muscle atrophy in vivo. Nat Cell Biol 3, 1014-1019.

34
Boirie, Y., Dangin, M., Gachon, P., Vasson, M.P., Maubois, J.L., Beaufrere, B. 1997.
Slow and fast dietary proteins differently modulate postprandial protein accretion.
Proc Natl Acad Sci U S A 94, 14930-14935.
Brooks, S.V., Faulkner, J.A. 1994. Skeletal muscle weakness in old age: underlying
mechanisms. Med Sci Sports Exerc 26, 432-439.
Butterfield, T.A., Best, T.M. 2009. Stretch-activated ion channel blockade attenuates
adaptations to eccentric exercise. Med Sci Sports Exerc 41, 351-356.
Candow, D.G., Little, J.P., Chilibeck, P.D., Abeysekara, S., Zello, G.A., Kazachkov,
M., Cornish, S.M., Yu, P.H. 2008. Low-dose creatine combined with protein during
resistance training in older men. Med Sci Sports Exerc 40, 1645-1652.
Cary, L.A., Guan, J.L. 1999. Focal adhesion kinase in integrin-mediated signaling.
Front Biosci 4, D102-113.
Chaillou, T. 2017. Impaired ribosome biogenesis could contribute to anabolic
resistance to strength exercise in the elderly. J Physiol 595, 1447-1448.
Chen, S.E., Jin, B., Li, Y.P. 2007. TNF-alpha regulates myogenesis and muscle
regeneration by activating p38 MAPK. Am J Physiol Cell Physiol 292, C1660-1671.
Chesley, A., MacDougall, J.D., Tarnopolsky, M.A., Atkinson, S.A., Smith, K. 1992.
Changes in human muscle protein synthesis after resistance exercise. J Appl Physiol
73, 1383-1388.
Ciechanover, A. 1994. The ubiquitin-proteasome proteolytic pathway. Cell 79, 13-21.
Clarkson, P.M., Hubal, M.J. 2002. Exercise-induced muscle damage in humans. Am J
Phys Med Rehabil 81, S52-69.
Clarkson, P.M., Sayers, S.P. 1999. Etiology of exercise-induced muscle damage. Can
J Appl Physiol 24, 234-248.
Coburn, J.W., Housh, D.J., Housh, T.J., Malek, M.H., Beck, T.W., Cramer, J.T.,
Johnson, G.O., Donlin, P.E. 2006. Effects of leucine and whey protein
supplementation during eight weeks of unilateral resistance training. J Strength Cond
Res 20, 284-291.
Coleman, M.E., DeMayo, F., Yin, K.C., Lee, H.M., Geske, R., Montgomery, C.,
Schwartz, R.J. 1995. Myogenic vector expression of insulin-like growth factor I
stimulates muscle cell differentiation and myofiber hypertrophy in transgenic mice. J
Biol Chem 270, 12109-12116.
Coutinho, E.L., DeLuca, C., Salvini, T.F., Vidal, B.C. 2006. Bouts of passive
stretching after immobilization of the rat soleus muscle increase collagen
macromolecular organization and muscle fiber area. Connect Tissue Res 47, 278-286.
Criswell, D.S., Booth, F.W., DeMayo, F., Schwartz, R.J., Gordon, S.E., Fiorotto,
M.L. 1998. Overexpression of IGF-I in skeletal muscle of transgenic mice does not
prevent unloading-induced atrophy. Am J Physiol 275, E373-379.

35
Dangin, M., Boirie, Y., Garcia-Rodenas, C., Gachon, P., Fauquant, J., Callier, P.,
Ballevre, O., Beaufrere, B. 2001. The digestion rate of protein is an independent
regulating factor of postprandial protein retention. Am J Physiol Endocrinol Metab
280, E340-348.
Dangin, M., Guillet, C., Garcia-Rodenas, C., Gachon, P., Bouteloup-Demange, C.,
Reiffers-Magnani, K., Fauquant, J., Ballevre, O., Beaufrere, B. 2003. The rate of
protein digestion affects protein gain differently during aging in humans. J Physiol
549, 635-644.
Degens, H. 2010. The role of systemic inflammation in age-related muscle weakness
and wasting. Scand J Med Sci Sports 20, 28-38.
Delhaas, T., van der Meer, S.F.T., Schaart, G., Degens, H., Drost, M.R. In Press.
Steep increase in myonuclear domain size during infancy. Anat Rec.
DeMartino, G.N., Ordway, G.A. 1998. Ubiquitin-proteasome pathway of intracellular
protein degradation: implications for muscle atrophy during unloading. Exerc Sport
Sci Rev 26, 219-252.
DeVol, D.L., Rotwein, P., Sadow, J.L., Novakofski, J., Bechtel, P.J. 1990. Activation
of insulin-like growth factor gene expression during work-induced skeletal muscle
growth. Am J Physiol 259, E89-95.
Drummond, M.J., Rasmussen, B.B. 2008. Leucine-enriched nutrients and the
regulation of mammalian target of rapamycin signalling and human skeletal muscle
protein synthesis. Curr Opin Clin Nutr Metab Care 11, 222-226.
Du, J., Wang, X., Miereles, C., Bailey, J.L., Debigare, R., Zheng, B., Price, S.R.,
Mitch, W.E. 2004. Activation of caspase-3 is an initial step triggering accelerated
muscle proteolysis in catabolic conditions. J Clin Invest 113, 115-123.
Durieux, A.C., D'Antona, G., Desplanches, D., Freyssenet, D., Klossner, S.,
Bottinelli, R., Fluck, M. 2009. Focal adhesion kinase is a load-dependent governor of
the slow contractile and oxidative muscle phenotype. J Physiol 587, 3703-3717.
Ebbeling, C.B., Clarkson, P.M. 1989. Exercise-induced muscle damage and
adaptation. Sports Med 7, 207-234.
Edstrom, E., Ulfhake, B. 2005. Sarcopenia is not due to lack of regenerative drive in
senescent skeletal muscle. Aging Cell 4, 65-77.
Eley, H.L., Russell, S.T., Baxter, J.H., Mukerji, P., Tisdale, M.J. 2007. Signaling
pathways initiated by beta-hydroxy-beta-methylbutyrate to attenuate the depression of
protein synthesis in skeletal muscle in response to cachectic stimuli. Am J Physiol
Endocrinol Metab 293, E923-931.
Eley, H.L., Russell, S.T., Tisdale, M.J. 2007. Effect of branched-chain amino acids on
muscle atrophy in cancer cachexia. Biochem J 407, 113-120.
Enoka, R.M. 1996. Eccentric contractions require unique activation strategies by the
nervous system. J Appl Physiol 81, 2339-2346.

36
Erskine, R.M., Fletcher, G., Folland, J.P. 2014. The contribution of muscle
hypertrophy to strength changes following resistance training. Eur J Appl Physiol
114, 1239-1249.
Erskine, R.M., Fletcher, G., Hanson, B., Folland, J.P. 2012. Whey protein does not
enhance the adaptations to elbow flexor resistance training. Med Sci Sports Exerc 44,
1791-1800.
Erskine, R.M., Jones, D.A., Williams, A.G., Stewart, C.E., Degens, H. 2010a. Inter-
individual variability in the adaptation of human muscle specific tension to
progressive resistance training. Eur J Appl Physiol 110, 1117-1125.
Erskine, R.M., Jones, D.A., Williams, A.G., Stewart, C.E., Degens, H. 2010b.
Resistance training increases in vivo quadriceps femoris muscle specific tension in
young men. Acta Physiol (Oxf) 199, 83-89.
Erskine, R.M., Tomlinson, D.J., Morse, C.I., Winwood, K., Hampson, P., Lord, J.M.,
Onambele, G.L. 2017. The individual and combined effects of obesity- and ageing-
induced systemic inflammation on human skeletal muscle properties. Int J Obes
(Lond) 41, 102-111.
Ervasti, J.M. 2003. Costameres: the Achilles' heel of Herculean muscle. J Biol Chem
278, 13591-13594.
Evans, W.J. 1995. What is sarcopenia? J Gerontol A Biol Sci Med Sci 50 Spec No, 5-
8.
Evans, W.J., Cannon, J.G. 1991. The metabolic effects of exercise-induced muscle
damage. Exerc Sport Sci Rev 19, 99-125.
Farges, M.C., Balcerzak, D., Fisher, B.D., Attaix, D., Bechet, D., Ferrara, M.,
Baracos, V.E. 2002. Increased muscle proteolysis after local trauma mainly reflects
macrophage-associated lysosomal proteolysis. Am J Physiol Endocrinol Metab 282,
E326-335.
Faulkner, J.A., Larkin, L.M., Claflin, D.R., Brooks, S.V. 2007. Age-related changes
in the structure and function of skeletal muscles. Clin Exp Pharmacol Physiol 34,
1091-1096.
Feasson, L., Stockholm, D., Freyssenet, D., Richard, I., Duguez, S., Beckmann, J.S.,
Denis, C. 2002. Molecular adaptations of neuromuscular disease-associated proteins
in response to eccentric exercise in human skeletal muscle. J Physiol 543, 297-306.
Fiatarone, M.A., O'Neill, E.F., Ryan, N.D., Clements, K.M., Solares, G.R., Nelson,
M.E., Roberts, S.B., Kehayias, J.J., Lipsitz, L.A., Evans, W.J. 1994. Exercise training
and nutritional supplementation for physical frailty in very elderly people. N Engl J
Med 330, 1769-1775.
Fluck, M., Carson, J.A., Gordon, S.E., Ziemiecki, A., Booth, F.W. 1999. Focal
adhesion proteins FAK and paxillin increase in hypertrophied skeletal muscle. Am J
Physiol 277, C152-162.

37
Fluck, M., Ziemiecki, A., Billeter, R., Muntener, M. 2002. Fibre-type specific
concentration of focal adhesion kinase at the sarcolemma: influence of fibre
innervation and regeneration. J Exp Biol 205, 2337-2348.
Franceschi, C., Bonafe, M., Valensin, S., Olivieri, F., De Luca, M., Ottaviani, E., De
Benedictis, G. 2000. Inflamm-aging. An evolutionary perspective on
immunosenescence. Ann N Y Acad Sci 908, 244-254.
Franco, A., Jr., Lansman, J.B. 1990a. Calcium entry through stretch-inactivated ion
channels in mdx myotubes. Nature 344, 670-673.
Franco, A., Jr., Lansman, J.B. 1990b. Stretch-sensitive channels in developing muscle
cells from a mouse cell line. J Physiol 427, 361-380.
Friden, J. 1984. Muscle soreness after exercise: implications of morphological
changes. Int J Sports Med 5, 57-66.
Friden, J., Kjorell, U., Thornell, L.E. 1984. Delayed muscle soreness and cytoskeletal
alterations: an immunocytological study in man. Int J Sports Med 5, 15-18.
Friden, J., Sjostrom, M., Ekblom, B. 1983. Myofibrillar damage following intense
eccentric exercise in man. Int J Sports Med 4, 170-176.
Fukunaga, T., Miyatani, M., Tachi, M., Kouzaki, M., Kawakami, Y., Kanehisa, H.
2001. Muscle volume is a major determinant of joint torque in humans. Acta Physiol
Scand 172, 249-255.
Gan, B., Yoo, Y., Guan, J.L. 2006. Association of focal adhesion kinase with tuberous
sclerosis complex 2 in the regulation of s6 kinase activation and cell growth. J Biol
Chem 281, 37321-37329.
Godard, M.P., Williamson, D.L., Trappe, S.W. 2002. Oral amino-acid provision does
not affect muscle strength or size gains in older men. Med Sci Sports Exerc 34, 1126-
1131.
Goldspink, G. 1999. Changes in muscle mass and phenotype and the expression of
autocrine and systemic growth factors by muscle in response to stretch and overload. J
Anat 194 ( Pt 3), 323-334.
Goll, D.E., Dayton, W.R., Singh, I., Robson, R.M. 1991. Studies of the alpha-
actinin/actin interaction in the Z-disk by using calpain. J Biol Chem 266, 8501-8510.
Gomes, M.D., Lecker, S.H., Jagoe, R.T., Navon, A., Goldberg, A.L. 2001. Atrogin-1,
a muscle-specific F-box protein highly expressed during muscle atrophy. Proc Natl
Acad Sci U S A 98, 14440-14445.
Guharay, F., Sachs, F. 1984. Stretch-activated single ion channel currents in tissue-
cultured embryonic chick skeletal muscle. J Physiol 352, 685-701.
Gulati, P., Gaspers, L.D., Dann, S.G., Joaquin, M., Nobukuni, T., Natt, F., Kozma,
S.C., Thomas, A.P., Thomas, G. 2008. Amino acids activate mTOR complex 1 via
Ca2+/CaM signaling to hVps34. Cell Metab 7, 456-465.

38
Guttridge, D.C., Mayo, M.W., Madrid, L.V., Wang, C.Y., Baldwin, A.S., Jr. 2000.
NF-kappaB-induced loss of MyoD messenger RNA: possible role in muscle decay
and cachexia. Science 289, 2363-2366.
Haegens, A., Schols, A.M., van Essen, A.L., van Loon, L.J., Langen, R.C. 2012.
Leucine induces myofibrillar protein accretion in cultured skeletal muscle through
mTOR dependent and -independent control of myosin heavy chain mRNA levels. Mol
Nutr Food Res 56, 741-752.
Hameed, M., Orrell, R.W., Cobbold, M., Goldspink, G., Harridge, S.D. 2003.
Expression of IGF-I splice variants in young and old human skeletal muscle after high
resistance exercise. J Physiol 547, 247-254.
Harridge, S.D., Kryger, A., Stensgaard, A. 1999. Knee extensor strength, activation,
and size in very elderly people following strength training. Muscle Nerve 22, 831-
839.
Hartman, J.W., Moore, D.R., Phillips, S.M. 2006. Resistance training reduces whole-
body protein turnover and improves net protein retention in untrained young males.
Appl Physiol Nutr Metab 31, 557-564.
Hartman, J.W., Tang, J.E., Wilkinson, S.B., Tarnopolsky, M.A., Lawrence, R.L.,
Fullerton, A.V., Phillips, S.M. 2007. Consumption of fat-free fluid milk after
resistance exercise promotes greater lean mass accretion than does consumption of
soy or carbohydrate in young, novice, male weightlifters. Am J Clin Nutr 86, 373-
381.
Hershko, A., Ciechanover, A. 1998. The ubiquitin system. Annu Rev Biochem 67,
425-479.
Holm, L., Olesen, J.L., Matsumoto, K., Doi, T., Mizuno, M., Alsted, T.J., Mackey,
A.L., Schwarz, P., Kjaer, M. 2008. Protein-containing nutrient supplementation
following strength training enhances the effect on muscle mass, strength, and bone
formation in postmenopausal women. J Appl Physiol 105, 274-281.
Hornberger, T.A., Mateja, R.D., Chin, E.R., Andrews, J.L., Esser, K.A. 2005. Aging
does not alter the mechanosensitivity of the p38, p70S6k, and JNK2 signaling
pathways in skeletal muscle. J Appl Physiol 98, 1562-1566.
Hornberger, T.A., Sukhija, K.B., Wang, X.R., Chien, S. 2007. mTOR is the
rapamycin-sensitive kinase that confers mechanically-induced phosphorylation of the
hydrophobic motif site Thr(389) in p70(S6k). FEBS Lett 581, 4562-4566.
Huang, Z., Chen, D., Zhang, K., Yu, B., Chen, X., Meng, J. 2007. Regulation of
myostatin signaling by c-Jun N-terminal kinase in C2C12 cells. Cell Signal 19, 2286-
2295.
Hubal, M.J., Gordish-Dressman, H., Thompson, P.D., Price, T.B., Hoffman, E.P.,
Angelopoulos, T.J., Gordon, P.M., Moyna, N.M., Pescatello, L.S., Visich, P.S.,
Zoeller, R.F., Seip, R.L., Clarkson, P.M. 2005. Variability in muscle size and strength
gain after unilateral resistance training. Med Sci Sports Exerc 37, 964-972.

39
Hulmi, J.J., Kovanen, V., Selanne, H., Kraemer, W.J., Hakkinen, K., Mero, A.A.
2009. Acute and long-term effects of resistance exercise with or without protein
ingestion on muscle hypertrophy and gene expression. Amino Acids 37, 297-308.
Jacquemin, V., Furling, D., Bigot, A., Butler-Browne, G.S., Mouly, V. 2004. IGF-1
induces human myotube hypertrophy by increasing cell recruitment. Exp Cell Res
299, 148-158.
Jakobsson, F., Borg, K., Edstrom, L., Grimby, L. 1988. Use of motor units in relation
to muscle fiber type and size in man. Muscle Nerve 11, 1211-1218.
Jones, D.A., Rutherford, O.M. 1987. Human muscle strength training: the effects of
three different regimens and the nature of the resultant changes. J Physiol 391, 1-11.
Jones, D.A., Rutherford, O.M., Parker, D.F. 1989. Physiological changes in skeletal
muscle as a result of strength training. Q J Exp Physiol 74, 233-256.
Jones, S.W., Hill, R.J., Krasney, P.A., O'Conner, B., Peirce, N., Greenhaff, P.L. 2004.
Disuse atrophy and exercise rehabilitation in humans profoundly affects the
expression of genes associated with the regulation of skeletal muscle mass. Faseb J
18, 1025-1027.
Kameyama, T., Etlinger, J.D. 1979. Calcium-dependent regulation of protein
synthesis and degradation in muscle. Nature 279, 344-346.
Kamikawa, Y., Ikeda, S., Harada, K., Ohwatashi, A., Yoshida, A. 2013. Passive
repetitive stretching for a short duration within a week increases myogenic regulatory
factors and myosin heavy chain mRNA in rats' skeletal muscles.
ScientificWorldJournal 2013, 493656.
Kanehisa, H., Ikegawa, S., Fukunaga, T. 1994. Comparison of muscle cross-sectional
area and strength between untrained women and men. Eur J Appl Physiol Occup
Physiol 68, 148-154.
Kasperek, G.J., Snider, R.D. 1985. Increased protein degradation after eccentric
exercise. Eur J Appl Physiol Occup Physiol 54, 30-34.
Kim, E., Guan, K.L. 2009. RAG GTPases in nutrient-mediated TOR signaling
pathway. Cell Cycle 8, 1014-1018.
Kimball, S.R., O'Malley, J.P., Anthony, J.C., Crozier, S.J., Jefferson, L.S. 2004.
Assessment of biomarkers of protein anabolism in skeletal muscle during the life span
of the rat: sarcopenia despite elevated protein synthesis. Am J Physiol Endocrinol
Metab 287, E772-780.
Klossner, S., Durieux, A.C., Freyssenet, D., Flueck, M. 2009. Mechano-transduction
to muscle protein synthesis is modulated by FAK. Eur J Appl Physiol 106, 389-398.
Koopman, R., Zorenc, A.H., Gransier, R.J., Cameron-Smith, D., van Loon, L.J. 2006.
Increase in S6K1 phosphorylation in human skeletal muscle following resistance
exercise occurs mainly in type II muscle fibers. Am J Physiol Endocrinol Metab 290,
E1245-1252.

40
Kosek, D.J., Bamman, M.M. 2008. Modulation of the dystrophin-associated protein
complex in response to resistance training in young and older men. J Appl Physiol
104, 1476-1484.
Kovacheva, E.L., Hikim, A.P., Shen, R., Sinha, I., Sinha-Hikim, I. 2010. Testosterone
supplementation reverses sarcopenia in aging through regulation of myostatin, c-Jun
NH2-terminal kinase, Notch, and Akt signaling pathways. Endocrinology 151, 628-
638.
Kumar, V., Atherton, P., Smith, K., Rennie, M.J. 2009. Human muscle protein
synthesis and breakdown during and after exercise. J Appl Physiol 106, 2026-2039.
Langen, R.C., Schols, A.M., Kelders, M.C., van der Velden, J.L., Wouters, E.F.,
Janssen-Heininger, Y.M. 2006. Muscle wasting and impaired muscle regeneration in a
murine model of chronic pulmonary inflammation. Am J Respir Cell Mol Biol 35,
689-696.
Langen, R.C., Van Der Velden, J.L., Schols, A.M., Kelders, M.C., Wouters, E.F.,
Janssen-Heininger, Y.M. 2004. Tumor necrosis factor-alpha inhibits myogenic
differentiation through MyoD protein destabilization. Faseb J 18, 227-237.
Langley, B., Thomas, M., Bishop, A., Sharma, M., Gilmour, S., Kambadur, R. 2002.
Myostatin inhibits myoblast differentiation by down-regulating MyoD expression. J
Biol Chem 277, 49831-49840.
Larsson, L. 1983. Histochemical characteristics of human skeletal muscle during
aging. Acta Physiol Scand 117, 469-471.
Larsson, L., Sjodin, B., Karlsson, J. 1978. Histochemical and biochemical changes in
human skeletal muscle with age in sedentary males, age 22--65 years. Acta Physiol
Scand 103, 31-39.
Lecker, S.H., Jagoe, R.T., Gilbert, A., Gomes, M., Baracos, V., Bailey, J., Price, S.R.,
Mitch, W.E., Goldberg, A.L. 2004. Multiple types of skeletal muscle atrophy involve
a common program of changes in gene expression. Faseb J 18, 39-51.
Lee, S.W., Dai, G., Hu, Z., Wang, X., Du, J., Mitch, W.E. 2004. Regulation of muscle
protein degradation: coordinated control of apoptotic and ubiquitin-proteasome
systems by phosphatidylinositol 3 kinase. J Am Soc Nephrol 15, 1537-1545.
Lexell, J., Taylor, C.C., Sjostrom, M. 1988. What is the cause of the ageing atrophy?
Total number, size and proportion of different fiber types studied in whole vastus
lateralis muscle from 15- to 83-year-old men. J Neurol Sci 84, 275-294.
Li, Y.P., Schwartz, R.J., Waddell, I.D., Holloway, B.R., Reid, M.B. 1998. Skeletal
muscle myocytes undergo protein loss and reactive oxygen-mediated NF-kappaB
activation in response to tumor necrosis factor alpha. Faseb J 12, 871-880.
Lieber, R.L., Friden, J. 1993. Muscle damage is not a function of muscle force but
active muscle strain. J Appl Physiol 74, 520-526.

41
Lieber, R.L., Friden, J. 1999. Mechanisms of muscle injury after eccentric
contraction. J Sci Med Sport 2, 253-265.
Lieber, R.L., Shah, S., Friden, J. 2002. Cytoskeletal disruption after eccentric
contraction-induced muscle injury. Clin Orthop Relat Res, S90-99.
Lio, D., Scola, L., Crivello, A., Colonna-Romano, G., Candore, G., Bonafe, M.,
Cavallone, L., Franceschi, C., Caruso, C. 2002. Gender-specific association between -
1082 IL-10 promoter polymorphism and longevity. Genes Immun 3, 30-33.
MacIntyre, D.L., Reid, W.D., McKenzie, D.C. 1995. Delayed muscle soreness. The
inflammatory response to muscle injury and its clinical implications. Sports Med 20,
24-40.
Mackey, A.L., Esmarck, B., Kadi, F., Koskinen, S.O., Kongsgaard, M., Sylvestersen,
A., Hansen, J.J., Larsen, G., Kjaer, M. 2007. Enhanced satellite cell proliferation with
resistance training in elderly men and women. Scand J Med Sci Sports 17, 34-42.
Malik, R.K., Parsons, J.T. 1996. Integrin-dependent activation of the p70 ribosomal
S6 kinase signaling pathway. J Biol Chem 271, 29785-29791.
Mascher, H., Tannerstedt, J., Brink-Elfegoun, T., Ekblom, B., Gustafsson, T.,
Blomstrand, E. 2008. Repeated resistance exercise training induces different changes
in mRNA expression of MAFbx and MuRF-1 in human skeletal muscle. Am J
Physiol Endocrinol Metab 294, E43-51.
McCarthy, J.J., Mula, J., Miyazaki, M., Erfani, R., Garrison, K., Farooqui, A.B.,
Srikuea, R., Lawson, B.A., Grimes, B., Keller, C., Van Zant, G., Campbell, K.S.,
Esser, K.A., Dupont-Versteegden, E.E., Peterson, C.A. 2011. Effective fiber
hypertrophy in satellite cell-depleted skeletal muscle. Development 138, 3657-3666.
McFarlane, C., Plummer, E., Thomas, M., Hennebry, A., Ashby, M., Ling, N., Smith,
H., Sharma, M., Kambadur, R. 2006. Myostatin induces cachexia by activating the
ubiquitin proteolytic system through an NF-kappaB-independent, FoxO1-dependent
mechanism. J Cell Physiol 209, 501-514.
McKoy, G., Ashley, W., Mander, J., Yang, S.Y., Williams, N., Russell, B.,
Goldspink, G. 1999. Expression of insulin growth factor-1 splice variants and
structural genes in rabbit skeletal muscle induced by stretch and stimulation. J Physiol
516 ( Pt 2), 583-592.
McPherron, A.C., Lawler, A.M., Lee, S.J. 1997. Regulation of skeletal muscle mass
in mice by a new TGF-beta superfamily member. Nature 387, 83-90.
McPherron, A.C., Lee, S.J. 1997. Double muscling in cattle due to mutations in the
myostatin gene. Proc Natl Acad Sci U S A 94, 12457-12461.
Milacic, V., Dou, Q.P. 2009. The tumor proteasome as a novel target for gold(III)
complexes: implications for breast cancer therapy. Coord Chem Rev 253, 1649-1660.

42
Moore, D.R., Atherton, P.J., Rennie, M.J., Tarnopolsky, M.A., Phillips, S.M. 2011.
Resistance exercise enhances mTOR and MAPK signalling in human muscle over
that seen at rest after bolus protein ingestion. Acta Physiol (Oxf) 201, 365-372.
Moore, D.R., Churchward-Venne, T.A., Witard, O., Breen, L., Burd, N.A., Tipton,
K.D., Phillips, S.M. 2015. Protein ingestion to stimulate myofibrillar protein synthesis
requires greater relative protein intakes in healthy older versus younger men. J
Gerontol A Biol Sci Med Sci 70, 57-62.
Moore, D.R., Robinson, M.J., Fry, J.L., Tang, J.E., Glover, E.I., Wilkinson, S.B.,
Prior, T., Tarnopolsky, M.A., Phillips, S.M. 2009. Ingested protein dose response of
muscle and albumin protein synthesis after resistance exercise in young men. Am J
Clin Nutr 89, 161-168.
Morris, E.J., Fulton, A.B. 1994. Rearrangement of mRNAs for costamere proteins
during costamere development in cultured skeletal muscle from chicken. J Cell Sci
107 ( Pt 3), 377-386.
Musaro, A., McCullagh, K., Paul, A., Houghton, L., Dobrowolny, G., Molinaro, M.,
Barton, E.R., Sweeney, H.L., Rosenthal, N. 2001. Localized Igf-1 transgene
expression sustains hypertrophy and regeneration in senescent skeletal muscle. Nat
Genet 27, 195-200.
Nair, K.S., Schwartz, R.G., Welle, S. 1992. Leucine as a regulator of whole body and
skeletal muscle protein metabolism in humans. Am J Physiol 263, E928-934.
Narici, M.V., Hoppeler, H., Kayser, B., Landoni, L., Claassen, H., Gavardi, C., Conti,
M., Cerretelli, P. 1996. Human quadriceps cross-sectional area, torque and neural
activation during 6 months strength training. Acta Physiol Scand 157, 175-186.
Ochi, E., Hirose, T., Hiranuma, K., Min, S.K., Ishii, N., Nakazato, K. 2010. Elevation
of myostatin and FOXOs in prolonged muscular impairment induced by eccentric
contractions in rat medial gastrocnemius muscle. J Appl Physiol 108, 306-313.
Ohlsson, C., Mohan, S., Sjogren, K., Tivesten, A., Isgaard, J., Isaksson, O., Jansson,
J.O., Svensson, J. 2009. The role of liver-derived insulin-like growth factor-I. Endocr
Rev 30, 494-535.
Omairi, S., Matsakas, A., Degens, H., Kretz, O., Hansson, K.A., Solbra, A.V.,
Bruusgaard, J.C., Joch, B., Sartori, R., Giallourou, N., Mitchell, R., Collins-Hooper,
H., Foster, K., Pasternack, A., Ritvos, O., Sandri, M., Narkar, V., Swann, J.R., Huber,
T.B., Patel, K. 2016. Enhanced exercise and regenerative capacity in a mouse model
that violates size constraints of oxidative muscle fibres. Elife 5.
Ordway, G.A., Neufer, P.D., Chin, E.R., DeMartino, G.N. 2000. Chronic contractile
activity upregulates the proteasome system in rabbit skeletal muscle. J Appl Physiol
88, 1134-1141.
Ottenheijm, C.A., Heunks, L.M., Li, Y.P., Jin, B., Minnaard, R., van Hees, H.W.,
Dekhuijzen, P.N. 2006. Activation of the ubiquitin-proteasome pathway in the
diaphragm in chronic obstructive pulmonary disease. Am J Respir Crit Care Med 174,
997-1002.

43
Pardo, J.V., Siliciano, J.D., Craig, S.W. 1983. Vinculin is a component of an
extensive network of myofibril-sarcolemma attachment regions in cardiac muscle
fibers. J Cell Biol 97, 1081-1088.
Patel, T.J., Lieber, R.L. 1997. Force transmission in skeletal muscle: from actomyosin
to external tendons. Exerc Sport Sci Rev 25, 321-363.
Pedersen, B.K. 2009. The diseasome of physical inactivity--and the role of myokines
in muscle--fat cross talk. J Physiol 587, 5559-5568.
Pelosi, L., Giacinti, C., Nardis, C., Borsellino, G., Rizzuto, E., Nicoletti, C.,
Wannenes, F., Battistini, L., Rosenthal, N., Molinaro, M., Musaro, A. 2007. Local
expression of IGF-1 accelerates muscle regeneration by rapidly modulating
inflammatory cytokines and chemokines. Faseb J 21, 1393-1402.
Petrella, J.K., Kim, J.-S., Cross, J.M., Kosek, D.J., Bamman, M.M. 2006. Efficacy of
myonuclear addition may explain differential myofiber growth among resistance-
trained young and older men and women. Am J Physiol Endocrinol Metab 291, E937-
946.
Philip, B., Lu, Z., Gao, Y. 2005. Regulation of GDF-8 signaling by the p38 MAPK.
Cell Signal 17, 365-375.
Phillips, S.M., Tipton, K.D., Aarsland, A., Wolf, S.E., Wolfe, R.R. 1997. Mixed
muscle protein synthesis and breakdown after resistance exercise in humans. Am J
Physiol 273, E99-107.
Phillips, S.M., Tipton, K.D., Ferrando, A.A., Wolfe, R.R. 1999. Resistance training
reduces the acute exercise-induced increase in muscle protein turnover. Am J Physiol
276, E118-124.
Pimentel, G.D., Rosa, J.C., Lira, F.S., Zanchi, N.E., Ropelle, E.R., Oyama, L.M.,
Oller do Nascimento, C.M., de Mello, M.T., Tufik, S., Santos, R.V. 2011. beta-
Hydroxy-beta-methylbutyrate (HMbeta) supplementation stimulates skeletal muscle
hypertrophy in rats via the mTOR pathway. Nutr Metab (Lond) 8, 11.
Psilander, N., Damsgaard, R., Pilegaard, H. 2003. Resistance exercise alters MRF and
IGF-I mRNA content in human skeletal muscle. J Appl Physiol 95, 1038-1044.
Raj, D.A., Booker, T.S., Belcastro, A.N. 1998. Striated muscle calcium-stimulated
cysteine protease (calpain-like) activity promotes myeloperoxidase activity with
exercise. Pflugers Arch 435, 804-809.
Reeves, N.D., Narici, M.V., Maganaris, C.N. 2004. Effect of resistance training on
skeletal muscle-specific force in elderly humans. J Appl Physiol 96, 885-892.
Reid, M.B., Li, Y.P. 2001. Tumor necrosis factor-alpha and muscle wasting: a cellular
perspective. Respir Res 2, 269-272.
Riley, D.A., Ellis, S., Giometti, C.S., Hoh, J.F., Ilyina-Kakueva, E.I., Oganov, V.S.,
Slocum, G.R., Bain, J.L., Sedlak, F.R. 1992. Muscle sarcomere lesions and
thrombosis after spaceflight and suspension unloading. J Appl Physiol 73, 33S-43S.

44
Rodino-Klapac, L.R., Haidet, A.M., Kota, J., Handy, C., Kaspar, B.K., Mendell, J.R.
2009. Inhibition of myostatin with emphasis on follistatin as a therapy for muscle
disease. Muscle Nerve 39, 283-296.
Rommel, C., Bodine, S.C., Clarke, B.A., Rossman, R., Nunez, L., Stitt, T.N.,
Yancopoulos, G.D., Glass, D.J. 2001. 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.
Rybakova, I.N., Patel, J.R., Ervasti, J.M. 2000. The dystrophin complex forms a
mechanically strong link between the sarcolemma and costameric actin. J Cell Biol
150, 1209-1214.
Saini, A., Al-Shanti, N., Faulkner, S.H., Stewart, C.E. 2008. Pro- and anti-apoptotic
roles for IGF-I in TNF-alpha-induced apoptosis: a MAP kinase mediated mechanism.
Growth Factors 26, 239-253.
Sancak, Y., Bar-Peled, L., Zoncu, R., Markhard, A.L., Nada, S., Sabatini, D.M. 2010.
Ragulator-Rag complex targets mTORC1 to the lysosomal surface and is necessary
for its activation by amino acids. Cell 141, 290-303.
Sancak, Y., Peterson, T.R., Shaul, Y.D., Lindquist, R.A., Thoreen, C.C., Bar-Peled,
L., Sabatini, D.M. 2008. The Rag GTPases bind raptor and mediate amino acid
signaling to mTORC1. Science 320, 1496-1501.
Sandri, M., Sandri, C., Gilbert, A., Skurk, C., Calabria, E., Picard, A., Walsh, K.,
Schiaffino, S., Lecker, S.H., Goldberg, A.L. 2004. Foxo transcription factors induce
the atrophy-related ubiquitin ligase atrogin-1 and cause skeletal muscle atrophy. Cell
117, 399-412.
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, C1248-1257.
Schuelke, M., Wagner, K.R., Stolz, L.E., Hubner, C., Riebel, T., Komen, W., Braun,
T., Tobin, J.F., Lee, S.J. 2004. Myostatin mutation associated with gross muscle
hypertrophy in a child. N Engl J Med 350, 2682-2688.
Scime, A., Rudnicki, M.A. 2006. Anabolic potential and regulation of the skeletal
muscle satellite cell populations. Curr Opin Clin Nutr Metab Care 9, 214-219.
Shefer, G., Van de Mark, D.P., Richardson, J.B., Yablonka-Reuveni, Z. 2006.
Satellite-cell pool size does matter: defining the myogenic potency of aging skeletal
muscle. Dev Biol 294, 50-66.
Siriett, V., Platt, L., Salerno, M.S., Ling, N., Kambadur, R., Sharma, M. 2006.
Prolonged absence of myostatin reduces sarcopenia. J Cell Physiol 209, 866-873.
Siriett, V., Salerno, M.S., Berry, C., Nicholas, G., Bower, R., Kambadur, R., Sharma,
M. 2007. Antagonism of myostatin enhances muscle regeneration during sarcopenia.
Mol Ther 15, 1463-1470.

45
Solomon, V., Baracos, V., Sarraf, P., Goldberg, A.L. 1998. Rates of ubiquitin
conjugation increase when muscles atrophy, largely through activation of the N-end
rule pathway. Proc Natl Acad Sci U S A 95, 12602-12607.
Spangenburg, E.E., Le Roith, D., Ward, C.W., Bodine, S.C. 2008. A functional
insulin-like growth factor receptor is not necessary for load-induced skeletal muscle
hypertrophy. J Physiol 586, 283-291.
Spangenburg, E.E., McBride, T.A. 2006. Inhibition of stretch-activated channels
during eccentric muscle contraction attenuates p70S6K activation. J Appl Physiol
100, 129-135.
Stewart, C.E., Rotwein, P. 1996. Growth, differentiation, and survival: multiple
physiological functions for insulin-like growth factors. Physiol Rev 76, 1005-1026.
Stupka, N., Tarnopolsky, M.A., Yardley, N.J., Phillips, S.M. 2001. Cellular
adaptation to repeated eccentric exercise-induced muscle damage. J Appl Physiol 91,
1669-1678.
Suzuki, K., Sorimachi, H., Yoshizawa, T., Kinbara, K., Ishiura, S. 1995. Calpain:
novel family members, activation, and physiologic function. Biol Chem Hoppe Seyler
376, 523-529.
Szalay, K., Razga, Z., Duda, E. 1997. TNF inhibits myogenesis and downregulates
the expression of myogenic regulatory factors myoD and myogenin. Eur J Cell Biol
74, 391-398.
Taaffe, D.R., Jin, I.H., Vu, T.H., Hoffman, A.R., Marcus, R. 1996. Lack of effect of
recombinant human growth hormone (GH) on muscle morphology and GH-insulin-
like growth factor expression in resistance-trained elderly men. J Clin Endocrinol
Metab 81, 421-425.
Tang, J.E., Moore, D.R., Kujbida, G.W., Tarnopolsky, M.A., Phillips, S.M. 2009.
Ingestion of whey hydrolysate, casein, or soy protein isolate: effects on mixed muscle
protein synthesis at rest and following resistance exercise in young men. J Appl
Physiol 107, 987-992.
Terzis, G., Georgiadis, G., Stratakos, G., Vogiatzis, I., Kavouras, S., Manta, P.,
Mascher, H., Blomstrand, E. 2008. Resistance exercise-induced increase in muscle
mass correlates with p70S6 kinase phosphorylation in human subjects. Eur J Appl
Physiol 102, 145-152.
Thompson, H.S., Scordilis, S.P. 1994. Ubiquitin changes in human biceps muscle
following exercise-induced damage. Biochem Biophys Res Commun 204, 1193-1198.
Thomson, D.M., Gordon, S.E. 2006. Impaired overload-induced muscle growth is
associated with diminished translational signalling in aged rat fast-twitch skeletal
muscle. J Physiol 574, 291-305.
Tidball, J.G., Spencer, M.J., Wehling, M., Lavergne, E. 1999. Nitric-oxide synthase is
a mechanical signal transducer that modulates talin and vinculin expression. J Biol
Chem 274, 33155-33160.

46
Tieland, M., Dirks, M.L., van der Zwaluw, N., Verdijk, L.B., van de Rest, O., de
Groot, L.C., van Loon, L.J. 2012. Protein Supplementation Increases Muscle Mass
Gain During Prolonged Resistance-Type Exercise Training in Frail Elderly People: A
Randomized, Double-Blind, Placebo-Controlled Trial. J Am Med Dir Assoc.
Tipton, K.D., Ferrando, A.A., Phillips, S.M., Doyle, D., Jr., Wolfe, R.R. 1999.
Postexercise net protein synthesis in human muscle from orally administered amino
acids. Am J Physiol 276, E628-634.
Tipton, K.D., Rasmussen, B.B., Miller, S.L., Wolf, S.E., Owens-Stovall, S.K., Petrini,
B.E., Wolfe, R.R. 2001. Timing of amino acid-carbohydrate ingestion alters anabolic
response of muscle to resistance exercise. Am J Physiol Endocrinol Metab 281, E197-
206.
Tipton, K.D., Wolfe, R.R. 1998. Exercise-induced changes in protein metabolism.
Acta Physiol Scand 162, 377-387.
Trendelenburg, A.U., Meyer, A., Rohner, D., Boyle, J., Hatakeyama, S., Glass, D.J.
2009. Myostatin reduces Akt/TORC1/p70S6K signaling, inhibiting myoblast
differentiation and myotube size. Am J Physiol Cell Physiol 296, C1258-1270.
Verdijk, L.B., Gleeson, B.G., Jonkers, R.A., Meijer, K., Savelberg, H.H., Dendale, P.,
van Loon, L.J. 2009a. Skeletal muscle hypertrophy following resistance training is
accompanied by a fiber type-specific increase in satellite cell content in elderly men. J
Gerontol A Biol Sci Med Sci 64, 332-339.
Verdijk, L.B., Jonkers, R.A., Gleeson, B.G., Beelen, M., Meijer, K., Savelberg, H.H.,
Wodzig, W.K., Dendale, P., van Loon, L.J. 2009b. Protein supplementation before
and after exercise does not further augment skeletal muscle hypertrophy after
resistance training in elderly men. Am J Clin Nutr 89, 608-616.
Verdijk, L.B., Koopman, R., Schaart, G., Meijer, K., Savelberg, H.H., van Loon, L.J.
2007. Satellite cell content is specifically reduced in type II skeletal muscle fibers in
the elderly. Am J Physiol Endocrinol Metab 292, E151-157.
Vieillevoye, S., Poortmans, J.R., Duchateau, J., Carpentier, A. 2010. Effects of a
combined essential amino acids/carbohydrate supplementation on muscle mass,
architecture and maximal strength following heavy-load training. Eur J Appl Physiol
110, 479-488.
Visser, M., Pahor, M., Taaffe, D.R., Goodpaster, B.H., Simonsick, E.M., Newman,
A.B., Nevitt, M., Harris, T.B. 2002. Relationship of interleukin-6 and tumor necrosis
factor-alpha with muscle mass and muscle strength in elderly men and women: the
Health ABC Study. J Gerontol A Biol Sci Med Sci 57, M326-332.
Volpi, E., Mittendorfer, B., Rasmussen, B.B., Wolfe, R.R. 2000. The response of
muscle protein anabolism to combined hyperaminoacidemia and glucose-induced
hyperinsulinemia is impaired in the elderly. J Clin Endocrinol Metab 85, 4481-4490.
Warren, G.L., Hayes, D.A., Lowe, D.A., Armstrong, R.B. 1993. Mechanical factors in
the initiation of eccentric contraction-induced injury in rat soleus muscle. J Physiol
464, 457-475.

47
Welle, S., Burgess, K., Mehta, S. 2009. Stimulation of skeletal muscle myofibrillar
protein synthesis, p70 S6 kinase phosphorylation, and ribosomal protein S6
phosphorylation by inhibition of myostatin in mature mice. Am J Physiol Endocrinol
Metab 296, E567-572.
Welle, S., Totterman, S., Thornton, C. 1996. Effect of age on muscle hypertrophy
induced by resistance training. J Gerontol A Biol Sci Med Sci 51, M270-275.
West, D.W., Burd, N.A., Tang, J.E., Moore, D.R., Staples, A.W., Holwerda, A.M.,
Baker, S.K., Phillips, S.M. 2010. Elevations in ostensibly anabolic hormones with
resistance exercise enhance neither training-induced muscle hypertrophy nor strength
of the elbow flexors. J Appl Physiol 108, 60-67.
West, D.W., Kujbida, G.W., Moore, D.R., Atherton, P., Burd, N.A., Padzik, J.P., De
Lisio, M., Tang, J.E., Parise, G., Rennie, M.J., Baker, S.K., Phillips, S.M. 2009.
Resistance exercise-induced increases in putative anabolic hormones do not enhance
muscle protein synthesis or intracellular signalling in young men. J Physiol 587,
5239-5247.
Willoughby, D.S., Stout, J.R., Wilborn, C.D. 2007. Effects of resistance training and
protein plus amino acid supplementation on muscle anabolism, mass, and strength.
Amino Acids 32, 467-477.
Witard, O.C., Jackman, S.R., Breen, L., Smith, K., Selby, A., Tipton, K.D. 2014.
Myofibrillar muscle protein synthesis rates subsequent to a meal in response to
increasing doses of whey protein at rest and after resistance exercise. Am J Clin Nutr
99, 86-95.
Woolstenhulme, M.T., Conlee, R.K., Drummond, M.J., Stites, A.W., Parcell, A.C.
2006. Temporal response of desmin and dystrophin proteins to progressive resistance
exercise in human skeletal muscle. J Appl Physiol 100, 1876-1882.
Yakar, S., Liu, J.L., Stannard, B., Butler, A., Accili, D., Sauer, B., LeRoith, D. 1999.
Normal growth and development in the absence of hepatic insulin-like growth factor
I. Proc Natl Acad Sci U S A 96, 7324-7329.
Yang, H., Alnaqeeb, M., Simpson, H., Goldspink, G. 1997. Changes in muscle fibre
type, muscle mass and IGF-I gene expression in rabbit skeletal muscle subjected to
stretch. J Anat 190 ( Pt 4), 613-622.
Yang, S., Alnaqeeb, M., Simpson, H., Goldspink, G. 1996. Cloning and
characterization of an IGF-1 isoform expressed in skeletal muscle subjected to stretch.
J Muscle Res Cell Motil 17, 487-495.
Yang, W., Chen, Y., Zhang, Y., Wang, X., Yang, N., Zhu, D. 2006. Extracellular
signal-regulated kinase 1/2 mitogen-activated protein kinase pathway is involved in
myostatin-regulated differentiation repression. Cancer Res 66, 1320-1326.
Yang, Y., Breen, L., Burd, N.A., Hector, A.J., Churchward-Venne, T.A., Josse, A.R.,
Tarnopolsky, M.A., Phillips, S.M. 2012a. Resistance exercise enhances myofibrillar
protein synthesis with graded intakes of whey protein in older men. Br J Nutr, 1-9.

48
Yang, Y., Churchward-Venne, T.A., Burd, N.A., Breen, L., Tarnopolsky, M.A.,
Phillips, S.M. 2012b. Myofibrillar protein synthesis following ingestion of soy protein
isolate at rest and after resistance exercise in elderly men. Nutr Metab (Lond) 9, 57.
Yang, Y., Creer, A., Jemiolo, B., Trappe, S. 2005. Time course of myogenic and
metabolic gene expression in response to acute exercise in human skeletal muscle. J
Appl Physiol 98, 1745-1752.
Yeung, E.W., Whitehead, N.P., Suchyna, T.M., Gottlieb, P.A., Sachs, F., Allen, D.G.
2005. Effects of stretch-activated channel blockers on [Ca2+]i and muscle damage in
the mdx mouse. J Physiol 562, 367-380.
Zimmers, T.A., Davies, M.V., Koniaris, L.G., Haynes, P., Esquela, A.F., Tomkinson,
K.N., McPherron, A.C., Wolfman, N.M., Lee, S.J. 2002. Induction of cachexia in
mice by systemically administered myostatin. Science 296, 1486-1488.

49
FIGURE LEGENDS
Figure 1. The molecular signaling pathways associated with muscle hypertrophy and atrophy.
The binding of IGF-I to its receptor (IGF-IR) causes autophosphorylation of insulin receptor substrate
(IRS1). Phosphatidylinositol 3-kinase (PI3K) is a lipid kinase that phosphorylates phosphatidylinositol
(4,5)-bisphosphate, producing phosphatidylinositol (3,4,5)-trisphosphate, which is a membrane-binding
site for phosphoinositide-dependent protein kinase (PDK1). Upon translocation to the sarcolemma,
AKT (or protein kinase B, PKB) is phosphorylated by PDK1. Once activated, AKT phosphorylates
mammalian target of rapamycin complex 1 (mTORC1) directly and by phosphorylating and
inactivating the tuberous sclerosis complexes 1 and 2 (TSC1/2), which otherwise inhibit mTORC1
activation. Following resistance exercise, an influx of calcium ions (Ca2+) via stretch-activated
channels (SACs) and the activation of FAK in the costamere can inactivate TSC1/2, thus activating
mTORC1. Amino acids entering the muscle fiber cause RagGTPase-dependent translocation of
mTORC1 to the lysosome, where it is activated by ras homologous protein enriched in brain (Rheb).
mTORC1 subsequently activates 70KDa ribosomal S6 protein kinase (p70S6K), and inhibits 4E-BP
(also known as PHAS-1), which is a negative regulator of the eukaryotic translation initiation factor 4E
(eIF-4E). Phosphorylated AKT also inhibits glycogen-synthase kinase 3β (GSK3β), a substrate of AKT
that blocks protein translation initiated by the eIF-2B protein. All of these actions lead to increased
protein synthesis. However, protein degradation can be induced by pro-inflammatory cytokines, such
as tumour necrosis factor-α (TNF-α) and interleukin-6 (IL-6), which activate NF-κB via degradation of
I-κB, leading to increased transcription of the E3 ubiquitin-ligase, muscle RING-finger protein-1
(MuRF1). Another ligase, Atrogin1 (also known as MAFbx), is up-regulated by mitogen-activated
protein kinase (MAPK) p38, while both ligases are up-regulated by forkhead box (FOXO) transcription
factors. However, phosphorylated AKT blocks the transcriptional up-regulation of Atrogin1 and
MuRF1 by inhibiting FOXO1, while phosphorylated mTORC1 inhibits the up-regulation of Atrogin1
directly. Myostatin also increases protein degradation and decreases protein synthesis by activating
MAPKs and the SMAD complex, and by inhibiting PI3K. In addition, myostatin inhibits the myogenic
program, thus resulting in a decrease of myoblast proliferation.

50
Figure 2. Schematic representation of (A) the location of costameres within a skeletal muscle
fiber and (B) the proteins that constitute the costamere. Costameres are protein complexes
circumferentially aligned along the length of the muscle fiber that connect peripheral myofibrils at the
Z-disks to the sarcolemma and beyond to the extra-cellular matrix (ECM). The costamere comprises a
dystrophin/glycoprotein complex and a focal adhesion complex (FAC), which includes the integrin-
associated tyrosine kinase focal adhesion kinase (FAK). Fig. 2B modified from (Fluck et al., 2002)
with permission.
Figure 3. Breakdown of the protein fragment via the ubiquitin proteasome pathway. Multiple
ubiquitin (Ub) molecules form a polyubiquitin chain in a process involving the Ub-activating (E1), Ub-
conjugating (E2), and Ub-ligating (E3) enzymes. The targeted protein fragment is then selected for
degradation, or “tagged”, via the covalent attachment of the polyubiquitin chain. The tagging enables
the 19S (PA700) module to recognize the protein fragment, so that it can be further degraded by the
20S core into oligopeptides after it has been de-ubiquitinated and the Ub molecules released recycled.
Adapted from (Milacic, Dou, 2009) with permission.

51
Figure 1

52
Figure 2A
Figure 2B

53
Figure 3