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! 1!
Muscle wasting in the presence of disease, why is it so variable? 1!
2!
Paul R. Kemp1,*, Mark Griffiths1 and Michael I. Polkey2 3!
4!
1National Heart & Lung Institute, Imperial College London, South Kensington Campus, 5!
London SW7 2AZ, UK 6!
2National Institute for Health Research Respiratory Biomedical Research Unit at Royal 7!
Brompton and Harefield NHS Foundation Trust and Imperial College, London, London SW3, 8!
6NP, UK 9!
10!
Running title: Susceptibility to muscle wasting in disease 11!
12!
*Author for correspondence (E-mail: [email protected]; Tel.: 020 7594 1716). 13!
14!
ABSTRACT 15!
Skeletal muscle wasting is a common clinical feature of many chronic diseases and also 16!
occurs in response to single acute events. The accompanying loss of strength can lead to 17!
significant disability, increased care needs and have profound negative effects on quality of 18!
life. As muscle is the most abundant source of amino acids in the body, it appears to function 19!
as a buffer for fuel and substrates that can be used to repair damage elsewhere and to feed the 20!
immune system. In essence, the fundamentals of muscle wasting are simple: less muscle is 21!
made than is broken down. However, although well-described mechanisms modulate muscle 22!
protein turnover, significant individual differences in the amount of muscle lost in the 23!
presence of a given severity of disease complicate the understanding of underlying 24!
mechanisms and suggest that individuals have different sensitivities to signals for muscle 25!
! 2!
loss. Furthermore, the rate at which muscle protein is turned over under normal conditions 26!
means that clinically significant muscle loss can occur with changes in the rate of protein 27!
synthesis and/or breakdown that are too small to be measurable. Consequently, the changes in 28!
expression of factors regulating muscle turnover required to cause a decline in muscle mass 29!
are small and, except in cases of rapid wasting, there is no consistent pattern of change in the 30!
expression of factors that regulate muscle mass. MicroRNAs are fine tuners of cell phenotype 31!
and are therefore ideally suited to cause the subtle changes in proteome required to tilt the 32!
balance between synthesis and degradation in a way that causes clinically significant wasting. 33!
Herein we present a model in which muscle loss as a consequence of disease in non-muscle 34!
tissue is modulated by a set of microRNAs, the muscle expression of which is associated with 35!
severity of disease in the non-muscle tissue. These microRNAs alter fundamental biological 36!
processes including the synthesis of ribosomes and mitochondria leading to reduced protein 37!
synthesis and increased protein breakdown, thereby freeing amino acids from the muscle. We 38!
argue that the variability in muscle loss observed in the human population arises from at least 39!
two sources. The first is from pre-existing or disease-induced variation in the expression of 40!
microRNAs controlling the sensitivity of muscle to the atrophic signal and the second is from 41!
the expression of microRNAs from imprinted loci (i.e. only expressed from the maternally or 42!
paternally inherited allele) and may control the rate of myonuclear recruitment. In the 43!
absence of disease, these factors do not correlate with muscle mass, since there is no 44!
challenge to the established balance. However, in the presence of such a challenge, these 45!
microRNAs determine the rate of decline for a given disease severity. Together these 46!
mechanisms provide novel insight into the loss of muscle mass and its variation in the human 47!
population. The involvement of imprinted loci also suggests that genes that regulate early 48!
development also contribute to the ability of individuals to resist muscle loss in response to 49!
disease. 50!
! 3!
Key words: microRNA, TGF-beta signalling, muscle wasting, susceptibility, protein turnover. 51!
52!
CONTENTS 53!
I. Introduction 54!
II. Muscle loss in disease: a problem with homeostasis? 55!
(1) Why does disease promote muscle loss? !56!
(2) What are the dynamics of muscle wasting?!57!
III. Mechanisms contributing to the maintenance of skeletal muscle mass 58!
(1) Growth factors and cytokines: how strong is the signal? 59!
(2) Synthesising muscle 60!
(a) IGF-1/Akt/mTOR pathway: sensing the signal 61!
(b) Ribosome synthesis: regulating synthetic capacity 62!
(c) Satellite/stem cell activation: the capacity to recruit myonuclei 63!
(3) Degrading muscle 64!
(a) Myostatin/SMAD/FOXO/inflammatory signals: sensing the signal 65!
(b) Proteolysis: the capacity to degrade protein 66!
(c) Autophagy and mitophagy: the capacity to degrade organelles 67!
(4) Integration of signalling 68!
IV. Fine-tuning the muscle proteome by microRNAs: do miRNAs contribute to 69!
muscle wasting? 70!
(1) The role of miRNAs 71!
(2) myomiRs 72!
(3) A cluster of atromiRs 73!
(4) miR-422a and miR-378 74!
(5) miR-128 75!
! 4!
(6) Imprinting-associated miRNAs 76!
V. A unified model 77!
VI. Future questions 78!
VII. Conclusions 79!
VIII. References 80!
81!
I. INTRODUCTION!82!
Skeletal muscle constitutes approximately 40% of the body. It enables all voluntary 83!
movement, including locomotion and breathing, but also contributes to many homeostatic 84!
mechanisms including glucose disposal (Evans, Murray & Kissebah, 1984) and 85!
thermogenesis (Rowland, Bal & Periasamy, 2015). The phenotype of skeletal muscle is 86!
plastic, responding to changes in demand, most obviously in response to increased or reduced 87!
physical activity where the relevant muscle will either hypertrophy or atrophy (Harridge, 88!
2007; Schiaffino et al., 2013). However, this plasticity also allows muscle to be used as a 89!
reserve that can be exploited during periods of starvation, for example supplying amino acids 90!
for the immune system and other tissues in response to infection or tissue damage (Biolo et 91!
al., 2000; Gore & Jahoor, 2000; Lightfoot, McArdle & Griffiths, 2009). This latter function 92!
forms the focus of this review. 93!
A marked reduction in muscle mass, especially in the locomotor muscles, occurs as a 94!
common co-morbidity of several chronic diseases including chronic obstructive pulmonary 95!
disease (COPD) (Remels et al., 2013) and chronic heart failure (CHF) (Collamati et al., 96!
2016), as well as in response to acute insults that lead to critical illness, such as in so-called 97!
intensive care unit acquired weakness (ICUAW) (Bloch et al., 2012). Muscle wasting can 98!
lead to a marked reduction in the quality of life of patients and is associated with increased 99!
mortality (Anker et al., 1997; Schols et al., 2005; Pocock et al., 2008). Since increasing 100!
! 5!
numbers of people now live with chronic conditions, understanding the processes that 101!
contribute to muscle wasting is important in reducing the overall burden of ill health. 102!
103!
II. MUSCLE LOSS IN DISEASE: A PROBLEM WITH HOMEOSTASIS?!104!
(1) Why does disease promote muscle loss? !105!
The presence of disease as a common factor in the development of clinically relevant wasting 106!
suggests that some component of disease outside the muscle tissue elicits an atrophic 107!
response in the muscle whereby muscle acts as a source of amino acids (in particular essential 108!
amino acids) to provide building blocks and fuel for other tissues (Biolo et al., 2000; Gore & 109!
Jahoor, 2000; Lightfoot et al., 2009). These amino acids contribute to meeting the increased 110!
demand by the liver in the acute-phase response (Kaysen, 2004) and to the immune system 111!
(Straub et al., 2010; Straub, 2011). However, identifying components of disease that 112!
contribute to this process is complicated by multiple factors. First, inactivity often 113!
accompanies chronic disease and is, itself, a cause of muscle loss (Breen et al., 2013; 114!
Dideriksen et al., 2016). Furthermore, increased activity can reverse muscle wasting in 115!
patients (Wagner, 2006; Slot et al., 2014; West et al., 2016) so inactivity has been considered 116!
the primary driver of muscle wasting. However, it has also been argued that the inactivity 117!
allows diversion of energy from muscle to the immune system (Straub et al., 2010; Straub, 118!
2011) making it difficult to separate cause from effect. Indeed, Straub (2011) describes 119!
diurnal neuro-hormonal regulation of energy usage where during active waking hours the 120!
muscle and brain are the major energy-utilising systems whereas, during inactive hours, 121!
energy is diverted to the immune system. Consequently, in chronic disease where the increase 122!
in energy usage by the immune system is marked (Segerstrom, 2007; Straub, 2011), reduced 123!
activity allows energy to be appropriately redistributed. In these circumstances, both reduced 124!
activity and increased levels of inflammatory cytokines contribute to the change in balance of 125!
! 6!
muscle turnover (Fig. 1). Second, in the diseases studied, the magnitude of wasting is not 126!
directly associated with disease severity (Anker & Sharma, 2002; Natanek et al., 2013a). For 127!
example, in COPD there is only a weak association of muscle mass with classical markers of 128!
disease severity and it is only in very large studies where this association becomes 129!
significant: for example compare Natanek et al. (2013a) with Agusti et al. (2010). Similarly, 130!
skeletal muscle loss and retention can be found in patients at both ends of the spectrum of 131!
severity of CHF (Fulster et al., 2013) and in patients undergoing aortic surgery, the loss of 132!
muscle mass does not associate with either pre-existing disease severity or surgical 133!
parameters (Bloch et al., 2013; Farre Garros et al., 2017). Third, the molecular mechanisms 134!
associated with muscle wasting in response to inactivity and starvation are similar to those 135!
that occur in wasting associated with disease (Schiaffino et al., 2013), making it difficult to 136!
identify disease-specific processes. 137!
The use of muscle as a source of amino acids is possibly most marked in critical illness where 138!
sepsis promotes a marked increase in inflammatory cytokines that are associated with muscle 139!
wasting (Roth, 2007; Lightfoot et al., 2009; Bloch et al., 2012; Hoffer & Bistrian, 2016). 140!
However, given that wasting also occurs in response to a range of infections that lead to an 141!
acute-phase response (Friman & Wesslen, 2000), and increased nitrogen excretion is seen in 142!
response to infection (Beisel, 1984), the loss of muscle in critical illness is likely to reflect an 143!
extreme end of this response. 144!
In this review, we build on the view that muscle atrophy, both accompanying chronic disease 145!
and more acutely in response to critical illness, is a consequence of chronic or over-146!
stimulation of systems that evolved to use muscle as a fuel and substrate buffer in times of 147!
short-term illness or injury (Reeds, Fjeld & Jahoor, 1994; Preiser et al., 2014; Argiles et al., 148!
2016). One aspect of this hypothesis is that muscle responds to the signals that drive 149!
inflammation and the acute-phase response, to release both amino acids and energy in a 150!
! 7!
continuous responsive fashion rather than simply providing amino acids when circulating 151!
levels are low. This mechanism explains wasting in the presence of adequate nutrition and the 152!
relative failure of amino acid supplementation to inhibit muscle atrophy (Stein & Blanc, 153!
2011). It also explains why individuals continue to waste if medical treatment stabilises 154!
symptoms but does not repair organ damage (e.g. chronic kidney disease, where patients are 155!
maintained by dialysis, COPD where medication provides symptomatic relief and chronic 156!
heart disease where pharmacological agents improve cardiac performance). Potential 157!
signalling mechanisms that may contribute to wasting include the chronic inflammation and 158!
oxidative stress that are commonly found in chronic disease [reviewed in Bistrian (2007) and 159!
Moylan & Reid (2007)]. In this continuous responsive mechanism, the muscle provides 160!
substrates for inflammation and repair, but incomplete repair means that the demand is never 161!
met. As the turnover of muscle occurs at a significant rate, marginal imbalances in muscle 162!
synthesis and degradation explain the long-term decline of muscle mass seen in chronic 163!
conditions and perhaps in otherwise healthy seniors. This concept is more plausible because 164!
of emerging data demonstrating that microRNAs (miRNAs) fine tune the proteome, making 165!
them ideal modulators of the synthesis/degradation balance, the sensitivity of the signalling 166!
systems and the ability to recruit myonuclei. This latter component is partly dependent on the 167!
expression of imprinted miRNAs. 168!
169!
(2) What are the dynamics of muscle wasting?!170!
Fearon, Evans & Anker (2011)coined the term myopenia to describe muscle loss in chronic 171!
disease rather than as a consequence of old age. One criterion suggested for this diagnosis 172!
was 5% muscle loss over 6–12 months. The gradual nature of the process together with the 173!
constant turnover of muscle suggests that any imbalance in muscle synthesis and breakdown 174!
is very small. For example, in disuse atrophy following complete immobilisation, muscle loss 175!
! 8!
occurs at approximately 0.6%/day for the first 30 days, giving an 18% reduction in muscle 176!
mass (Phillips, Glover & Rennie, 2009). The bulk of this loss (0.5%/day) results from 177!
reduced protein synthesis with minimal changes in proteolysis (Phillips et al., 2009). Yet this 178!
rate of loss is nearly 20 times faster than that leading to clinically significant muscle loss in 179!
response to disease (5% in 6 months). Consequently, the changes in protein synthesis and 180!
protein degradation required to achieve the loss of muscle typical of a chronic disease is 181!
minimal. Indeed, using data from tracer studies (Mittendorfer et al., 2005; Caso et al., 2006; 182!
Timmerman et al., 2010; Poortmans et al., 2012), it can be calculated that 5% muscle loss in 183!
6 months can occur with less than a 3% increase in the absolute rate of protein breakdown or 184!
a 3% reduction in protein synthesis (Fig. 2). Consequently, the required change in synthesis 185!
or degradation that can lead to significant muscle loss is much smaller than the measured 186!
variation in the rate of protein turnover in tracer studies (standard deviation approximately 187!
20%). As a result, measuring differences in the expression of atrogenes or components of the 188!
protein synthetic pathway is difficult, as exemplified by different studies showing altered or 189!
unchanged atrogene expression and signalling pathway activity in COPD (Doucet et al., 190!
2007; Lemire et al., 2012; Riddoch-Contreras et al., 2012; Natanek et al., 2013b), heart 191!
failure (Gielen et al., 2012; Forman et al., 2014) and by transcriptomic studies showing no 192!
difference in atrogene expression in diseases ranging from cancer to critical illness, and 193!
reductions in atrogene expression in mouse models of atrophy (Melov et al., 2007; Mo et al., 194!
2010; Stephens et al., 2010; Jespersen et al., 2011b). 195!
The calculations in Fig. 2 assume that only one process changes but in reality, both synthesis 196!
and degradation are likely to change in response to disease. The signalling systems that 197!
modulate protein turnover regulate synthesis and degradation in opposite directions, although 198!
instances of synthesis and degradation both increasing or reducing are not unknown 199!
! 9!
(Schiaffino et al., 2013). Consequently, chronic muscle loss can be caused by small, 200!
unmeasurable absolute differences. 201!
202!
III. MECHANISMS CONTRIBUTING TO THE MAINTENANCE OF SKELETAL 203!
MUSCLE MASS!204!
The processes contributing to muscle synthesis include both protein synthesis and the 205!
accretion of new nuclei. Muscle breakdown results from net proteolysis, autophagy and 206!
apoptosis. The details of these processes and their contribution to muscle mass loss have been 207!
reviewed extensively elsewhere [see, for example, Glass (2005); Sandri (2008); Schiaffino et 208!
al. (2013)], so are only covered briefly here. The absolute rates of protein synthesis, 209!
degradation, recruitment of new nuclei and apoptosis vary, within limits, among individuals 210!
as a consequence of their environment, genetics and epigenetics. This variation will 211!
contribute to differences in growth and adult muscle mass in the population, with genetics 212!
accounting for up to 65% of the variation in muscle mass (Abney, McPeek & Ober, 2001). 213!
Genetic differences include mutations that prevent myostatin production leading to increased 214!
muscle mass and function (Schuelke et al., 2004), and polymorphisms in the Angiotensin 215!
converting enzyme (ACE) promoter that modify the response to training (Charbonneau et al., 216!
2008) and are associated with muscle strength in COPD (Hopkinson et al., 2004). Reduced 217!
regeneration and a reduction in satellite cell number can be seen in patients with mutations in 218!
selenoprotein N1 (Sepn1)-related myopathies (Castets et al., 2011). A combination of 219!
genetics and epigenetics also contributes to muscle mass and size as exemplified by the 220!
higher prevalence of polymorphisms reducing the DNA methylation cycle in elite athletes 221!
(Terruzzi et al., 2011) and of the contribution of methylation on cytosines that are followed 222!
by guanines (CpGs) in imprinted genes to birth weight (Hoyo et al., 2014). 223!
224!
! 10!
(1) Growth factors and cytokines: how strong is the signal?!225!
Under normal conditions, the signals promoting muscle synthesis are balanced by those 226!
promoting degradation (Fig. 1). Several growth factors have been identified that control these 227!
processes and are altered in response to activity, inflammation or disease. Insulin-like growth 228!
factor-1 (IGF-1) promotes muscle protein synthesis and reduces breakdown (Rommel et al., 229!
2001; Stitt et al., 2004; Latres et al., 2005). Circulating IGF-1 increases in response to 230!
physical activity (Philippou et al., 2009; Schwarz et al., 2016) and is reduced in the elderly 231!
(Bucci et al., 2013), in response to a number of diseases (Saeki, Hamada & Hiwada, 2002; 232!
Kythreotis et al., 2009), after burns (Nedelec, de Oliveira & Garrel, 2003) and following 233!
cardiac surgery (Bloch et al., 2013). In response to inflammation, there is an increase in 234!
levels of circulating cytokines that increase muscle protein breakdown. These cytokines 235!
include: tumour necrosis factor α (TNFα) which promotes atrophy in vitro (Dehoux et al., 236!
2007) and in vivo (Subramaniam et al., 2015), and interleukin-6 (IL-6) (Haddad et al., 2005; 237!
Gao et al., 2017). 238!
In addition to cytokines, TGF-β family members are also key regulators of muscle mass in 239!
health and disease. Myostatin limits muscle growth during development as seen by the 240!
hypermuscular ‘double muscled’ phenotype of animals with natural inactivating mutations 241!
[e.g. Belgian Blue cows (Grobet et al., 1997) and bully whippets (Mosher et al., 2007)]. 242!
Myostatin promotes atrophy in animals (Reisz-Porszasz et al., 2003; Lee et al., 2015), and is 243!
increased in response to a range of atrophic stimuli [including starvation (Jeanplong et al., 244!
2003), detraining (Jespersen et al., 2011a) and oxidative stress (Sriram et al., 2011)] and in a 245!
number of diseases [e.g. myocardial infarction, CHF (Mahmoudabady et al., 2008; Breitbart 246!
et al., 2011) and COPD (Plant et al., 2010; Man et al., 2010)]. Conversely blockade of the 247!
activin IIb receptor, to which myostatin and other negative regulators of muscle mass bind, in 248!
humans results in hypertrophy (Polkey et al., 2018). 249!
! 11!
Increased circulating levels of another TGF-β family member, growth differentiation factor-250!
15 (GDF-15), are also increased in a range of diseases where muscle wasting occurs 251!
including pulmonary arterial hypertension (PAH) (Rhodes, Wharton & Wilkins, 2013), 252!
COPD (Freeman et al., 2015; Patel et al., 2016), CHF (George et al., 2016) and some cancers 253!
(Shnaper et al., 2009; Pavo et al., 2016; Zhang et al., 2016a). Circulating GDF-15 levels 254!
correlate with muscle mass and function in COPD (Patel et al., 2016) and are associated with 255!
muscle loss following aortic surgery (Bloch et al., 2013). GDF-15 contributes to loss of 256!
muscle mass both by suppressing appetite (Johnen et al., 2007) via the GDNF family alpha 257!
receptor like (GFRAL) receptor in the central nervous system (Emmerson et al., 2017; Yang 258!
et al., 2017) and by directly stimulating muscle atrophy (Patel et al., 2016) most likely via an 259!
alternative mechanism as GFRAL receptor expression is restricted to the hind brain 260!
(Emmerson et al., 2017). Another member of this family that appears to contribute to muscle 261!
loss is GDF-11. Like myostatin, GDF-11 signals through the small mothers against 262!
decapentaplegic (SMAD) pathway. Initial studies suggested that GDF-11 was protective to 263!
skeletal and cardiac systems and that there was an age-related decrease in circulating GDF-11 264!
(Loffredo et al., 2013). However, these data appear to have been influenced by reagent 265!
specificity (Harper et al., 2016). Recent studies using both recombinant GDF-11 (Egerman et 266!
al., 2015) and GDF-11 over-expression (Hammers et al., 2017) showed muscle loss and 267!
reduced regenerative capacity. The problems with assaying GDF-11 mean that the role of this 268!
protein in muscle loss in patients is unknown. 269!
270!
(2) Synthesising muscle!271!
Changes in the expression of individual proteins is predominantly regulated by changes in 272!
messenger RNA (mRNA) levels or translation for that protein. However, muscle hypertrophy 273!
or atrophy require a more general change in protein synthesis affecting the entire pool of 274!
! 12!
mRNAs. These changes are predominantly achieved by altering the rate of initiation of 275!
protein synthesis (Jackson, Hellen & Pestova, 2010; Gobet & Naef, 2017). The control of 276!
translational initiation has been extensively reviewed elsewhere (Jackson et al., 2010; 277!
Iadevaia, Liu & Proud, 2014). Translation is also dependent on a sufficient number of free, 278!
fully formed 40S subunits, the number and availability of which is not often discussed, but is 279!
variable as can be seen from the effects of physical exercise on ribosomal RNA (rRNA) and 280!
protein content and the activation of rRNA synthesis by mammalian Target of Rapamycin 281!
(mTOR) (von Walden et al., 2012; Figueiredo et al., 2015; West et al., 2016). 282!
283!
(a) IGF-1/Akt/mTOR pathway: sensing the signal!284!
The best-studied pathway increasing protein synthesis is the IGF-1/protein kinase 285!
B(Akt)/mTOR pathway (Quevedo, Alcazar & Salinas, 2000). Increased IGF-1 activity, either 286!
through addition of exogenous ligand or over-expression of the gene (Musaro et al., 2001; 287!
Schertzer & Lynch, 2006) increases new protein synthesis. The downstream components of 288!
this pathway have been extensively described and reviewed elsewhere (Glass, 2005, 2010; 289!
Cook & Morley, 2007; Esser, 2008). 290!
291!
(b) Ribosome synthesis: regulating synthetic capacity !292!
Ribosome synthesis is energetically expensive, so the number of ribosomes is co-ordinated 293!
with the synthetic requirements of the cell (Gobet & Naef, 2017). In eukaryotic cells there are 294!
two forms of ribosome, cytoplasmic and mitochondrial, which are made of different proteins 295!
and rRNAs. Cytoplasmic ribosome synthesis requires the production of multiple proteins and 296!
RNAs (Boisvert et al., 2007; Perry, 2007; Cmarko et al., 2008). The rRNAs are transcribed 297!
by RNA polymerase I rather than RNA polymerase II, which transcribes mRNAs. The 298!
activity of RNA polymerase I is dependent on a set of transcription cofactors including 299!
! 13!
upstream binding transcription factor (UBTF) (Chan et al., 1991; Comai, Tanese & Tjian, 300!
1992). Modulation of UBTF activity has been associated with altered rRNA expression in 301!
muscle following exercise, at least in part as a consequence of Akt/mTOR activity, indicating 302!
that pathways promoting hypertrophy also stimulate ribosome synthesis (Figueiredo et al., 303!
2015; West et al., 2016). This increased rRNA transcription increases total RNA in 304!
hypertrophying tissue (Stec et al., 2016). 305!
Reduced mitochondrial function is associated with muscle dysfunction and loss of muscle 306!
mass (Puente-Maestu et al., 2009; Romanello & Sandri, 2013). Although the majority of 307!
proteins in mitochondria are synthesised in the cytoplasm and subsequently translocated to 308!
the mitochondria, mitochondrial ribosomes are required for the synthesis of 13 proteins 309!
essential for the electron transport chain and ATP synthase (Alexeyev, Ledoux & Wilson, 310!
2004). The mitochondrial rRNAs are encoded on mitochondrial DNA with a 12S rRNA and a 311!
16S rRNA for the small and large subunits, respectively (Peralta, Wang & Moraes, 2012). 312!
Translation in the mitochondria is 5#-CAP independent and shows marked similarities to 313!
bacterial translation. Mutations in mitochondrial ribosomal proteins preventing ribosome 314!
formation cause growth retardation and fatal lactic acidosis (Miller et al., 2004; Baertling et 315!
al., 2015). 316!
317!
(c) Satellite/stem cell activation: the capacity to recruit myonuclei!318!
Skeletal muscle hypertrophy, beyond a given size or in repair following damage, requires the 319!
recruitment of extra nuclei, predominantly from satellite cells (reviewed in Yin, Price & 320!
Rudnicki, 2013). In response to a range of stimuli these cells become activated, proliferate 321!
and differentiate into myoblasts that either proliferate before fusing with the fibre or return to 322!
their niche to maintain the satellite cell pool. Muscle-resident stem cells and bone-marrow-323!
derived stem cells can also contribute to muscle regeneration (Yin et al., 2013). 324!
! 14!
Satellite cell activation and proliferation are controlled by both stimulatory and inhibitory 325!
growth factors including hepatocyte growth factor/scatter factor, which promotes activation 326!
(Anastasi et al., 1997; Tatsumi et al., 1998), fibroblast growth factors and IGF-1 327!
(Vandenburgh et al., 1991; Lefaucheur & Sebille, 1995), which promote proliferation, bone 328!
morphogenetic proteins (BMPs) which inhibit cell cycle exit and so maintain the proliferative 329!
phenotype (Friedrichs et al., 2011) and myostatin which inhibits satellite cell activation and 330!
myoblast proliferation (McCroskery et al., 2003). 331!
The processes of stem cell proliferation, differentiation and return to the satellite cell niche 332!
must be tightly regulated to allow sufficient myoblast proliferation for growth and repair 333!
without depleting the stem cell pool and reducing the potential for future repair (Yue et al., 334!
2016). Intrinsic differences in the balance of satellite cell proliferation, return to quiescence 335!
and myoblast differentiation among individuals are potential sources of variability in the 336!
long-term maintenance of muscle mass both with age and in response to a long-term 337!
challenge like chronic disease. 338!
339!
(3) Degrading muscle !340!
(a) Myostatin/SMAD/FOXO/inflammatory signals: sensing the signal!341!
Several pathways contribute to muscle degradation including myostatin, TNF-α, and the 342!
inflammatory cytokines. Signalling via these pathways has been reviewed elsewhere (Glass, 343!
2005; Sandri, 2008) and leads by increased SMAD activity, increased forkhead box protein O 344!
FOXO activity and/or increased nuclear factor-kappa B (NF-κB) activity, to increased 345!
expression of ubiquitin ligases and/or components of the autophagy pathway. In addition to 346!
growth factor- and cytokine-dependent signals, oxidative stress can stimulate atrophy by 347!
increasing the activity of the proteolytic systems (Powers, Smuder & Judge, 2012; Abrigo et 348!
al., 2018). Oxidative stress can arise from several different sources with mitochondria a 349!
! 15!
major source of reactive oxygen species (ROS) (Bhat et al., 2015). Interestingly, the 350!
mitochondrial ROS also have a signalling role in response to physical activity, increasing the 351!
expression of peroxisomal proliferator activated receptor gamma coactivator 1α (PGC1α) 352!
and peroxisomal proliferator receptor (PPAR) transcription factors that contribute to 353!
mitochondriogenesis and antioxidant genes (e.g. superoxide dismutase). This beneficial 354!
increase is due to a transient rather than the sustained increase in ROS that can occur in 355!
disease (Ristow et al., 2009). 356!
357!
(b) Proteolysis: the capacity to degrade protein!358!
The best-studied system involved in muscle protein breakdown is the ubiquitin-dependent 359!
proteolytic system (UPS). This system allows for targeted protein breakdown using 360!
individual ubiquitin ligases to label specific proteins for degradation. In muscle atrophy the 361!
ubiquitin ligases muscle ring finger protein 1 (MuRF1) and Atrogin-1 are key components 362!
that stimulate the degradation of both myofilament proteins and myogenic transcription 363!
factors, thereby degrading muscle proteins and reducing the synthesis of new ones (Bodine et 364!
al., 2001; Centner et al., 2001; Cohen et al., 2009; Lokireddy et al., 2012). It is commonly 365!
suggested that the expression of the genes encoding these proteins is increased in all atrophic 366!
conditions. However, as indicated above, a number of studies show no change or a reduction 367!
in their expression in atrophic conditions. The reasons for this lack of change are probably the 368!
marginal nature of the imbalance required to cause muscle loss, and the timing of biopsies 369!
with respect to any insult (de Boer et al., 2007). In addition to the UPS, calpains, caspases 370!
and lysosomal mechanisms contribute to muscle breakdown and turnover. 371!
372!
(c) Autophagy and mitophagy: the capacity to degrade organelles!373!
! 16!
Autophagy and mitophagy also contribute to muscle breakdown. As the muscle shrinks, the 374!
number of organelles required reduces and these are processed by the bulk breakdown 375!
mechanism: autophagy. Autophagy is required for normal muscle homeostasis and mice that 376!
lack specific components of the autophagic system are not protected from muscle wasting, 377!
indeed the animals have accelerated muscle wasting most probably due to the accumulation 378!
of dysfunctional organelles (Masiero & Sandri, 2010). Although autophagy occurs under 379!
normal conditions, it is accelerated in response to atrophic factors including myostatin (Lee, 380!
Hopkinson & Kemp, 2011). 381!
Mitophagy is a specialised form of autophagy that removes mitochondria and plays an 382!
important role in the quality control of mitochondria (Tolkovsky, 2009; Romanello & Sandri, 383!
2010). Mitophagy can be triggered by the loss of membrane potential and by oxidative stress 384!
(de Boer et al., 2007). The mechanisms contributing to autophagy and mitophagy have been 385!
reviewed previously (de Boer et al., 2007; Romanello & Sandri, 2013). 386!
387!
(4) Integration of signalling 388!
The signalling pathways that regulate the processes described above tend to operate through 389!
nodes leading to increased protein synthesis and reduced degradation or vice versa (Sandri, 390!
2008). The principle nodes that control muscle synthesis and breakdown are Akt and the 391!
FOXO transcription factors. In this axis Akt phosphorylates and inactivates FOXO whereas 392!
the FOXO transcription factors increase the expression of Eukaryotic translation initiation 393!
factor 4E-binding protein 1 (4E-BP1) and down-regulates mTOR complex 1 (mTORC1) (an 394!
activator of Akt) (Sandri, 2008). 395!
However, as indicated above, other than in cases of rapid changes in muscle mass, the 396!
changes may be very subtle as only a very small change in net protein synthesis or 397!
degradation is required for significant loss of muscle over a period of months. It should also 398!
! 17!
be remembered that the responses to individual growth factors (e.g. IGF-1) or activities of 399!
individual proteins (e.g. Akt) have been identified in experimental systems designed to 400!
minimise changes in other variables. The responses to disease or activity in vivo are not as 401!
straightforward, as they operate in the presence of other activators and homeostatic 402!
mechanisms that will modify the overall response. 403!
404!
IV. FINE-TUNING THE MUSCLE PROTEOME BY MICRORNAS: DO miRNAS 405!
CONTRIBUTE TO MUSCLE WASTING?!406!
(1) The role of miRNAs!407!
The ability of miRNAs to regulate the synthesis of individual proteins or subsets of proteins 408!
allows them to fine-tune the proteome thereby adapting cell phenotype. The mechanisms 409!
controlling the expression and processing of miRNAs to produce the mature miRNA, and 410!
those involved in targeting individual mRNAs have been widely reviewed (He & Hannon, 411!
2004). The selectivity of miRNAs is driven by the ‘seed sequence’, bases 2–8 of the mature 412!
miRNA (Brennecke et al., 2005). It is not necessary for a particular miRNA to target the 413!
whole pathway, although many miRNAs do regulate multiple components in pathways, 414!
rather it is important that they target the ‘rate-determining step’ to have the maximum 415!
influence. MicroRNAs regulate muscle biology at a number of steps, influencing many major 416!
cellular functions including metabolism, differentiation, hypertrophy and atrophy (Chen et 417!
al., 2006; Drummond et al., 2008; Chen, Callis & Wang, 2009; Guller & Russell, 2010; Dey, 418!
Pfeifer & Dutta, 2014; Kemp & Natanek, 2017). In muscle, studies of differential expression 419!
of miRNAs and the pathways they control have primarily focussed on the muscle specific 420!
miRNAS (myomiRs), a set of miRNAs that are enriched in skeletal muscle. miRNAs also 421!
circulate in the plasma and measurements of circulating levels of miRNAs have also yielded 422!
! 18!
information on muscle wasting (Cacchiarelli et al., 2011; Donaldson et al., 2013; Lee et al., 423!
2017). 424!
425!
(2) myomiRs !426!
The myomiRs include miR-1, -133, -206, and -499. miR-133 is co-expressed with either 427!
miR-1 or miR-206 in bi-cistronic RNAs (Chen et al., 2006; Ma et al., 2015; Dai et al., 2016). 428!
miR-499 is expressed from an intron within myosin heavy chain 7b (myH7b) (van Rooij et 429!
al., 2009). 430!
miR-1 and -206 are similar and share a seed sequence suggesting that they target many of the 431!
same mRNAs (McCarthy, 2008). Expression of miR-206 is almost completely restricted to 432!
skeletal muscle, whereas miR-1 is also expressed in the heart and vasculature (McCarthy, 433!
2008). Changes in miRNA expression occur in development, disease and in response to 434!
exercise as described below (Chen et al., 2006; Liu et al., 2008; McCarthy, 2008). 435!
Correspondingly, these miRNAs have important roles in the differentiation of muscle and in 436!
the control of fibre type, and miR-1 and miR-206 appear to have roles both during 437!
development and in the maintenance of muscle mass. For example, these miRNAs target 438!
Histone deacetylase 4 (HDAC4) and a DNA polymerase α subunit (Kim et al., 2006; Dai et 439!
al., 2016), so have been implicated in promoting regeneration and in the recruitment of new 440!
myonuclei. More recently, we have shown that miR-1 targets the TGF-β receptor 1 (TGF-441!
βR1), thereby reducing TGF-β levels and potentially myostatin signalling (M. Connolly and 442!
PK unpublished data,). The expression of these miRNAs is affected by contractile history and 443!
by disease. For example, miR-1 increases in response to a single bout of activity, but 444!
decreases after long-term training regimens (Nielsen et al., 2010). In light of these effects of 445!
exercise on miR-1 expression, changes in response to disease are difficult to predict and 446!
conflicting data have been reported. For example, miR-1 was suppressed in patients with 447!
! 19!
COPD and associated with smoking history, measured as cigarette-pack years (Lewis et al., 448!
2012) similarly miR-1 was suppressed in patients with established ICUAW (Bloch et al., 449!
2015). Other studies have reported that miR-1 is increased in patients with COPD (Puig-450!
Vilanova et al., 2014, 2015). These differences could be ascribed to the smoking or physical 451!
activity patterns of controls or patients, or their timing in relation to the biopsy and whether 452!
individuals fasted prior to study. The ability of miR-206 to regulate myonuclear recruitment 453!
may contribute to its potential for the treatment of amyotrophic lateral sclerosis (ALS). 454!
Patients and animal models of ALS have shown markedly reduced expression of miR-206 455!
and an associated reduction in muscle regeneration. In G93A-SOD1 mice lacking miR-206, 456!
progression of ALS is accelerated indicating a role for the miRNA in stabilising muscle–457!
nerve interactions (Williams et al., 2009; Ma et al., 2015; Park, 2015). 458!
miR-133 is a well-studied miRNA that has three isoforms: miR-133a1, miR-133a2 and miR-459!
133b. Early studies showed that miR-133a1/2 inhibited myoblast differentiation and 460!
promoted myoblast proliferation by targeting serum response factor, a transcription factor 461!
that promotes the expression of many components of the contractile apparatus (Chen et al., 462!
2006). More recently this role has been questioned by data showing that miR-133 inhibits 463!
proliferation by targeting specificity protein 1 (SP1) (Zhang et al., 2012) and that expression 464!
of miR-133a or b reduced the proportion of cells in the S phase of the cell cycle by targeting 465!
fibroblast growth factor receptor 1 (FGFR1) and the catalytic subunits of protein phosphatase 466!
2a (PP2A) (Feng et al., 2013). This reduction in PP2A activity caused a reduction in 467!
extracellular signal regulated kinase 1/2 (ERK1/2) phosphorylation and increased C2C12 468!
myoblast differentiation. These diametrically opposing results (increased or reduced myoblast 469!
proliferation and increased or suppressed differentiation) indicate that miR-133 targets a 470!
range of components in the differentiation and proliferation of muscle cells with effects on 471!
both the rate of cell proliferation and on differentiation. The precise function of the miRNA 472!
! 20!
will therefore depend on the relative expression and importance of each targeted component 473!
in the cell system under study. Knockout of both miR-133a isoforms causes cardiac defects 474!
but does not appear to alter skeletal muscle morphology, possibly due to redundancy with 475!
miR-133b (Liu et al., 2008). Similarly knockout of the locus containing miR-133b does not 476!
produce any phenotype, probably as a result of redundancy with miR-133a loci (Boettger et 477!
al., 2014). In vivo over-expression of miR-133a did not alter muscle development (Deng, 478!
Chen & Wang, 2011) 479!
miR-499 is expressed from an intron in the gene encoding MyH7b and appears to maintain 480!
the slow fibre phenotype (van Rooij et al., 2009). miR-499 targets components of the fast 481!
muscle program, including the transcription factor SRY-box 6 (Sox6), suppressing the type II 482!
fibre transcriptional program and enabling a feedback loop maintaining the slow fibre 483!
phenotype. In COPD patients, there is an increase in the proportion of hybrid fibres, raising 484!
the possibility of a reduction in the ability to maintain fibre type (Natanek et al., 2013a). A 485!
reduction in miR-499 could promote this fibre shift by enabling the fast transcription 486!
program. In addition to contributing to the maintenance of the slow fibre type, miR-499 may 487!
contribute to the maintenance of muscle mass as in silico analysis predicts that it targets 488!
myostatin signalling, but these effects remain to be demonstrated experimentally. Consistent 489!
with a role in regulating muscle mass, miR-499 expression is suppressed in muscle from 490!
patients with established ICUAW, as well as being associated with fat-free mass index 491!
(FFMI) in patients with COPD (Lewis et al., 2012; Bloch et al., 2015). However, it is also 492!
possible that this association is a result of the restriction of miR-499 expression to type I 493!
fibres combined with the reduction in the proportion of type I fibres (through another 494!
mechanism) that is seen in both conditions. 495!
496!
(3) A cluster of atromiRs!497!
! 21!
Several signalling systems contribute to muscle wasting by reducing net protein synthesis and 498!
increasing protein breakdown via the Akt/mTOR/FOXO pathways (reviewed in Sandri, 499!
2008). If disease is a major contributor to mobilisation of amino acids from muscle tissue, 500!
these processes and the expression of their signalling intermediates and ultimate target genes 501!
would be expected to associate with the severity of disease. For example, in COPD it would 502!
be expected that there would be a robust negative association of MuRF and atrogin with 503!
markers of residual lung function [e.g. forced expiratory volume in 1 second (FEV1) or 504!
transfer capacity of the lung for carbon monoxide (TLCO)] and/or a positive association with 505!
phospho-Akt or other markers of the protein synthetic signalling pathway. However, as 506!
indicated above, increased levels of MuRF and atrogin are not always demonstrated. 507!
Furthermore, the phosphorylation components of the Akt pathway were not increased in the 508!
quadriceps of COPD patients (Plant et al., 2010) compared to controls so cannot be correlated 509!
with disease severity. In part the small differences required to cause muscle wasting are likely 510!
to contribute to this lack of significant change but other factors may also contribute to the 511!
regulation of muscle protein turnover in disease. 512!
Recently, the expression of a cluster of miRNAs on the X-chromosome was found to be 513!
increased in the quadriceps of patients with COPD (Farre-Garros et al., 2015, 2017; Connolly 514!
et al., 2017). The miRNAs in this locus include miR-542-3p, -542-5p, -424-5p, -424-3p, 515!
miR-503, -450a and -450b. Interestingly the expression of at least three of these miRNAs in 516!
the quadriceps is associated with markers of primary disease severity (Connolly et al., 2017; 517!
Farre Garros et al., 2017). For example, in COPD their expression is associated with lung 518!
function, whereas in patients about to undergo aortic surgery their expression is associated 519!
with cardiac function and Euroscore (a clinical marker of risk associated with cardiac 520!
surgery) but not with lung function. Similarly, in patients with established ICUAW, their 521!
expression was associated with hospital length of stay and SOFA score (a clinical marker of 522!
! 22!
critical illness severity) at the time of biopsy. Hence, quadriceps expression of these miRNAs 523!
appears to correlate with extent of disease in a remote organ(s), suggesting that they may 524!
regulate the response necessary to produce amino acids for tissue repair elsewhere. 525!
Interestingly, quadriceps expression of these miRNAs in patients about to undergo aortic 526!
surgery is proportional to the amount of muscle that they lose over the subsequent 7 days, 527!
indicating a role in determining the ability of patients to respond to the insult of surgery. 528!
Analysis of the targets of these miRNAs indicates that they have a direct role in establishing 529!
the atrophic milieu of the muscle, targeting the production of ribosomes in both the 530!
cytoplasm and mitochondria as well as regulating the TGF-β signalling pathway. 531!
Furthermore, over-expression of these miRNAs in mice causes rapid muscle wasting 532!
(Connolly et al., 2017; Farre Garros et al., 2017). 533!
Functionally, miR-542-3p targets the synthesis of a range of cytoplasmic and mitochondrial 534!
ribosomal proteins (Wang et al., 2014; Farre Garros et al., 2017). The rate of translational 535!
initiation is determined by the number and relative activity of small ribosomal subunits. A 536!
reduction in the number of ribosomal subunits may well show little detectable effect on basal 537!
or fasted rates of protein synthesis but is likely to limit maximal protein synthetic capacity. 538!
Consequently, as the amount of protein synthesised over the course of a day will result from 539!
periods of low and of stimulated protein synthesis, a reduction in ribosomal complement is 540!
likely to reduce the average rate of protein synthesis. As indicated above, the required change 541!
in protein synthesis is relatively small to generate the loss of muscle seen in myopenic 542!
patients. 543!
An inability to produce complete small ribosomal subunits (Fig. 3) also leads to ribosome 544!
stress (Wang et al., 2014), in which the amount of rRNA is reduced with a preferential 545!
reduction in the small ribosomal subunit rRNA [18S (Wang et al., 2014) and 12S rRNA for 546!
the cytoplasmic and mitochondrial ribosomes, respectively (Miller et al., 2004; Saada et al., 547!
! 23!
2007)]. In extremis in mammals, ribosomal stress is associated with activation of p53 and the 548!
apoptotic pathway due to an excess of free ribosomal protein L11 (RPL11), a protein of the 549!
large ribosomal subunit a which can displace mouse double-minute 2 (mdm2) from p53, 550!
stabilising p53 (Miller et al., 2004; Saada et al., 2007). 551!
Consistent with mitochondrial ribosomal stress, levels of 12S rRNA were reduced compared 552!
to the levels of 16S rRNA in response to miR-542-3p in cells in culture and in response to the 553!
over-expression of miR-542 in the muscle of mice (Farre Garros et al., 2017). Furthermore, 554!
there was a reduction in 12S rRNA compared to 16S rRNA in patients with COPD and 555!
ICUAW, indicating mitochondrial ribosomal stress in these patients compared to their 556!
respective controls. A consequence of mitochondrial ribosomal stress will be reduced 557!
expression of proteins encoded by the mitochondrial genome. These proteins include 558!
components of complexes I, III, IV and V of oxidative phosphorylation. In cells transfected 559!
with miR-542-3p, there was a reduction in the expression of cytochrome b (cytb), a 560!
component of complex III, and a reduction in the membrane potential. Similarly, in mice 561!
over-expressing miR-542 there was a reduction in complex I activity and in membrane 562!
potential. Consistent with Wang et al. (2014), we also found a reduction in the expression of 563!
ribosomal protein S22 (RPS22), a cytoplasmic ribosomal protein, and an association of miR-564!
542-3p expression in the muscle of COPD patients with 18S rRNA expression (R. Farre-565!
Garros and PK, unpublished data). 566!
miR-424 is also likely to contribute to muscle wasting by reducing the number of ribosomes, 567!
but not by specifically targeting ribosomal proteins. In pull-down assays, the most highly 568!
enriched target for miR-424 was RNA polymerase IA (PolR1A) and the transcription factor 569!
UBTF (Connolly et al., 2017), which are required for the synthesis of the rRNAs (Fig. 3). 570!
The expression of both PolR1A and UBTF was suppressed by miR-424-5p. Furthermore, 571!
using β2 microglobulin and hypoxanthine phosphoribosyl transferase (HPRT) as reference 572!
! 24!
genes, the expression of 18S and 28S and the precursor rRNA 47S, was suppressed in cells 573!
transfected with miR-424-5p. Consistent with these observations, in patients with ICUAW 574!
(those with the largest increase in miR-424-5p expression) there was a reduction in the 575!
expression of UBTF (Connolly et al., 2017). miR-424 is also predicted to target many 576!
components of the IGF-1 signalling system and inhibits the expression of both IGF-1 577!
(Connolly et al., 2018) and insulin-like growth factor receptor 1 (IGF-1R) (Llobet-Navas et 578!
al., 2014) suggesting that it contributes to dampening a major hypertrophic signal as well as 579!
the capacity to respond to such a signal. 580!
In addition to causing mitochondrial dysfunction by inhibiting mitochondrial ribosome 581!
synthesis, miR-542 contributes directly to muscle wasting by acting as an intracellular 582!
regulator of SMAD signalling, the pathway used by myostatin. Transfection of cells with 583!
miR-542-5p increases basal activity of the SMAD2/3-dependent CAGA12 promoter (Farre 584!
Garros et al., 2017) and nuclear phospho-SMAD2/3. To achieve this increase, miR-542 585!
targets inhibitors of the SMAD signalling system including SMAD7 and the SMAD specific 586!
ubiquitin ligase SMURF1, inhibitors of the receptor complex, and phosphatases that 587!
inactivate phospho-SMAD. For example, transfection of myoblasts with miR-542 reduces the 588!
expression of the PP2A catalytic subunit alpha, PPP2CA (Farre Garros et al., 2017), which 589!
dephosphorylates the receptor and SMAD3 (Heikkinen et al., 2010), and is predicted to target 590!
the protein phosphatases PPM1a, CTDSP1 and CTDSP2, the C-terminal SMAD 591!
phosphatases that limit pathway activity. Consistent with targeting these proteins, the 592!
expression of SMAD7, SMURF1 and PPP2CA is suppressed in the muscle of patients with 593!
ICUAW who have the highest levels of miR-542-5p, as well increased TGF-β signalling 594!
(Farre Garros et al., 2017). miR-424 has also been shown to target the expression of SMAD7 595!
(Fig. 3) and increase the sensitivity of cells to TGF-β (Wang et al., 2016). 596!
! 25!
The combined effects of the miR-542/424 locus – increased SMAD signalling and sensitivity 597!
to TGF-β ligands, reduced IGF-1 and IGF-1R together with suppressed rRNA synthesis and 598!
ribosomal stress – would contribute to increased atrophy, reduced protein synthesis and 599!
mitochondrial dysfunction, all of which contribute to muscle wasting. Their association with 600!
severity of disease in the diseased organ also suggests a role for these miRNAs in wasting in 601!
the presence of disease. Consistent with this suggestion, the expression of miR-424 and miR-602!
542 in the muscle prior to cardiac surgery is proportional to the amount of muscle that will be 603!
lost in the subsequent 7 days (Connolly et al., 2017; Farre Garros et al., 2017). 604!
605!
(4) miR-422a and miR-378!606!
miR-422a is only present in primates and is expressed from an intragenic region. Circulating 607!
levels of miR-422a were originally inversely associated with muscle strength and size in 608!
patients with COPD (Paul et al., 2018). In the muscle, expression of miR-422a was higher in 609!
patients than in controls but surprisingly miR-422a expression was directly proportional to 610!
muscle size in COPD patients but not in normal healthy elderly individuals. This observation 611!
raised the possibility that increased expression of miR-422a was a protective response that 612!
limited wasting. Consistent with this suggestion, SMAD4 was identified as a target of miR-613!
422a and transfection of human skeletal myoblasts with miR-422a inhibited TGF-β 614!
signalling. As canonical TGF-β signalling is dependent on SMAD4, these data suggest that at 615!
least one mechanism by which miR-422 provides resistance to muscle wasting is through the 616!
inhibition of SMAD-dependent wasting (Paul et al., 2018). These data suggest a microRNA-617!
dependent mechanism that regulates the TGF-β signalling pathway in skeletal muscle as 618!
shown in Fig. 3. 619!
Another potentially important target of miR-422a is mutL homologue 1 (MLH1), a 620!
transcriptional co-activator of p53 (Mao et al., 2012). Consequently, increased miR-422a is 621!
! 26!
likely to reduce p53 activity. In addition to targeting these mRNAs, miR-422a was able to 622!
bind to the mRNAs for several transcription regulators that contribute to skeletal muscle 623!
phenotype, including myogenic factor 6 (myf6), inhibitor of DNA binding 2 (ID2) and 624!
myocyte enhancer factor 2D (MEF2D) (Paul et al., 2018). Whether and in what direction it 625!
regulates the expression of these proteins remains to be determined. 626!
The mature sequence of miR-422a is only two nucleotides different from that of miR-378a-627!
3p, a microRNA encoded by intron 1 of the PGC-1β gene. As these miRNAs share a seed 628!
sequence, they are likely to target many of the same mRNAs. Transgenic mice over-629!
expressing miR-378a have increased energy usage and are more resistant to a high-fat diet 630!
(Zhang et al., 2016b). The transgenic mice are smaller than their wild-type counterparts and 631!
have less skeletal muscle. Furthermore, their muscles contain fewer fibres which also tend to 632!
be smaller than those of their wild-type counterparts. Further analysis indicates that the miR-633!
378 transgenic mice have reduced satellite cell differentiation in response to injury. The 634!
authors suggest that this is a result of reduced IGF-1R expression but would also be 635!
consistent with reduced SMAD4 expression, as BMPs are required for muscle differentiation. 636!
These data raise the question of why, if they target the same mRNAs, miR-422a associates 637!
with strength in muscle yet miR-378a over-expression leads to smaller mice. One suggestion 638!
is that the timing of miRNA expression is critical. In the transgenic mouse model, the 639!
miRNA is expressed at all stages of muscle development but normally PGC-1β expression 640!
(and thereby, probably, miR-378a expression) increases during myoblast differentiation 641!
(Shintaku et al., 2016). Early expression of the miRNA would reduce myoblast proliferation 642!
by inhibiting the response to IGF-1 (Knezevic et al., 2012) and reduce SMAD4 inhibiting 643!
BMP signalling, whereas during normal development this does not occur. Consistent with 644!
this suggestion, tail-vein injection of miR-378 is reported to promote muscle hypertrophy 645!
! 27!
with injection of the antagonist to this miR (antagomiR) reducing muscle mass and strength 646!
(McCormick et al., 2017). 647!
648!
(5) miR-128!649!
Increased miR-128 has been reported in the skeletal muscle of patients with chronic kidney 650!
disease (Watson et al., 2016). miR-128 expression increases with muscle differentiation in 651!
cells in culture. However, this expression appears to limit the hypertrophy of myofibres by 652!
targeting the expression of insulin receptor substrate 1 (IRS-1) and down-regulating Akt 653!
phosphorylation. Consistent with this function, inhibition of miR-128 in vitro and in vivo 654!
promotes myotube hypertrophy (Motohashi et al., 2013). Interestingly miR-128 was 655!
identified by a screen of miRNAs dysregulated in the muscle of COPD patients (Farre Garros 656!
et al., 2017) and is associated with FFMI in these patients (R. Farre-Garros and PK 657!
unpublished data). 658!
659!
(6) Imprinting-associated miRNAs!660!
Genomic imprinting is an epigenetic phenomenon that causes genes to be expressed in a 661!
parent-of-origin-specific manner. In a screen of miRNAs that were different between COPD 662!
patients with a low FFMI and those with a normal FFMI, patients with a low FFMI had 663!
reduced expression of miRNAs from an imprinted miRNA cluster on chromosome 19 664!
(C19MC) (Lewis et al., 2016) that is expressed from the paternally inherited chromosome. 665!
Conversely, the same patients had higher expression of the maternally expressed miR-675. 666!
Furthermore, the expression of miR-675 and its host gene (H19) was inversely proportional 667!
to strength. That there was no association of the expression of these miRNAs with FFMI or 668!
strength (Lewis et al., 2016) in healthy controls and no difference in their expression between 669!
patients and controls, indicates that their expression does not change significantly in response 670!
! 28!
to disease, but that patients at one end of the expression profile are more likely to undergo 671!
muscle wasting in the presence of disease. 672!
These miRNAs contribute to the control of cell differentiation and proliferation. The 673!
paternally expressed miRNAs from the C19MC promote the pluripotent stem cell phenotype 674!
(Lin et al., 2010; Sun et al., 2011), whereas miR-675 inhibits myoblast proliferation and 675!
promotes terminal myoblast differentiation at least in part by inhibiting BMP signalling 676!
(Keniry et al., 2012; Dey et al., 2014). Interestingly, like miR-378-3p the effects of miR-675 677!
are also likely to be timing dependent. For example, in human patients the expression of miR-678!
675 was inversely proportional to muscle mass and strength, but experiments in mice showed 679!
that increasing miR-675 promoted myoblast differentiation and increased muscle mass. 680!
However, in the latter studies miR-675 was increased after the myoblasts had proliferated 681!
(Dey et al., 2014). Had miR-675 been increased at the time of injury, it seems likely that it 682!
would have inhibited cell proliferation and thereby reduced regeneration. 683!
These imprinted miRNAs are likely to control muscle mass in patients with chronic diseases 684!
via the recruitment of new nuclei (Fig. 4). As indicated above, myonuclear recruitment is a 685!
key component of the maintenance of muscle mass and individuals who can stimulate this 686!
process when required, through having a sufficient number of satellite cells that go through 687!
more rounds of cell division before terminally differentiating, will be able to maintain muscle 688!
better than those whose myoblasts differentiate and fuse earlier. 689!
Further evidence for differential expression of miRNAs from imprinted genomic regions 690!
contributing to a susceptibility to wasting comes from the observation that circulating levels 691!
of miR-485-3p, a maternally expressed miRNA from a cluster on chromosome 14, are 692!
associated with muscle strength in COPD patients (Lee et al., 2017). These data raise the 693!
possibility that the measurement of imprinted miRNAs in the circulation will allow the 694!
identification of individuals likely to waste following a catabolic signal. 695!
! 29!
696!
V. A UNIFIED MODEL!697!
The data described above indicate that the maintenance of muscle mass is the consequence of 698!
a delicately balanced set of processes, each with multiple inputs and controls that are 699!
regulated by environmental, genetic and epigenetic factors. One important set of controls are 700!
miRNAs that modulate the activity of fundamental biological processes, thereby contributing 701!
to the flexibility of the system (Fig. 5). 702!
A key environmental factor is the presence of a disease that, via one or more mechanisms, 703!
promotes muscle loss. The response to this environmental challenge will depend on the 704!
sensitivity of the individual to signals from the diseased tissue (e.g. inflammatory cytokines, 705!
myostatin or GDF-15) and their ability to regenerate. The data suggest that the expression of 706!
miRNAs from the miR-424/542 cluster is elevated by systemic disease (Connolly et al., 707!
2017; Farre Garros et al., 2017). These miRNAs alter the expression of a range of proteins 708!
associated with protein synthesis, reducing both the sensitivity of the muscle to IGF-1 and the 709!
capacity of the muscle to synthesise protein. Furthermore, by increasing the sensitivity of the 710!
muscle to TGF-β signalling, they will increase protein degradation. The effects of these 711!
miRNAs on TGF-β signalling are likely to be amplified by the reduction in miR-1 and miR-712!
181 that has been observed (Lewis et al., 2012; Bloch et al., 2015). Both miRNAs target 713!
activin A receptor-like kinase (Alk5), a TGF-βR1 involved in myostatin signalling, and a 714!
reduction in these miRNAs would increase Alk5 expression. Consequently, by changing the 715!
expression of these miRNAs, SMAD signalling can increase in the absence of any change in 716!
TGF-β or a similar ligand. The sensitivity with which the protein turnover is balanced means 717!
that absolute changes in the expression of genes that need to occur in an individual to 718!
promote muscle loss are smaller than the natural variation in the population, making 719!
quantifying important changes in individuals difficult. 720!
! 30!
The variation in response to the environmental insult (which drives the difference in response 721!
across the population) appears to come from at least two sources. The first is the sensitivity of 722!
individuals to each pathway. The TGF-β signalling system provides a good example of the 723!
regulation of signalling by miRNAs. Increased miR-422a reduces SMAD4 expression, 724!
thereby reducing the ability of the muscle to respond to myostatin and contributing to 725!
increased strength (Paul et al., 2018). The second area of variability is myonuclear 726!
recruitment, which, in muscle, will depend on the balance of satellite cell/myoblast 727!
proliferation and differentiation. The data suggest that higher levels of miRNAs that promote 728!
the pluripotent-stem-cell phenotype favour maintenance of muscle mass, whereas higher 729!
levels of miRNAs that promote withdrawal from the cell cycle and differentiation favour 730!
wasting (Lewis et al., 2016). These observations imply that delaying myocyte differentiation 731!
and extending the proliferation period increases the ability of the muscle to regenerate, thus 732!
leading to a slower loss of muscle mass in response to injury or insult. It is interesting that the 733!
miRNAs that contribute to this process come from imprinted loci, indicating a role for DNA 734!
methylation in the control of the muscle response to disease. The expression of these 735!
miRNAs is not affected by disease suggesting that individuals respond differently to the 736!
extent of disease as a function of miRNA expression. 737!
The miRNA pattern we have observed is consistent in patients with chronic disease and those 738!
on the critical care unit. It is easy to understand how the miRNA pattern contributes to 739!
muscle loss in the chronic conditions as the change in balance of processes required to 740!
produce muscle loss over months is so small. The question is why does this pattern also 741!
predict the much more rapid muscle loss in patients following cardiac surgery? The simplest 742!
answer to this is that muscle loss in these circumstances occurs in response to a large signal, 743!
in this case marked elevation of inflammatory cytokines and GDF-15. Individual variation in 744!
the sensitivity to these signals will be governed by the expression of the components of the 745!
! 31!
signalling complexes, from the receptor through to the transcription factors that activate or 746!
inhibit the expression of target genes. It is at this level that the miRNAs will contribute and 747!
individuals that are more sensitive to the signals will waste more rapidly. This mechanism is 748!
exemplified by the ability of miR-422a and miR-542-5p to control the sensitivity of cell 749!
SMAD signalling as described above (Farre Garros et al., 2017; Paul et al., 2018). However, 750!
these are unlikely to be the only mechanisms by which sensitivity is regulated. For example, 751!
the lethal 7 (let-7) family of miRNAs regulate toll-like receptors and STAT signalling so 752!
individual variation in these miRNAs is likely to contribute to the sensitivity of individuals to 753!
cytokines (Dissanayake & Inoue, 2016). Counterintuitively therefore, let-7f is elevated in 754!
patients with COPD who have lost fat-free mass but not in those that have retained their fat-755!
free mass (Lewis et al., 2016), suggesting that if let-7f contributes to muscle wasting it is 756!
through an alternative mechanism. 757!
Not only are these miRNAs likely to contribute to the variability in the wasting response of 758!
individuals, but some are likely to contribute to the variability in response to therapy. For 759!
example, anti-myostatin therapies may not work as well in individuals with reduced SMAD4 760!
as the contribution of myostatin signalling to wasting will not be as large. Similarly, their 761!
effect may be markedly suppressed in patients with high levels of miR-542-5p, if the pathway 762!
has been activated intracellularly. In this latter case, the effectiveness of the therapy will be 763!
less in individuals with more severe disease. These are just two examples affecting one 764!
pathway but raise the possibility of miRNAs regulating steroid responses by targeting the 765!
androgen receptor or other similar effects. 766!
The effect of the changes in expression of a miRNA on protein levels in a tissue will be 767!
dependent on several factors. These include the relative expression of the miRNA compared 768!
to its target mRNAs, the affinity of the miRNA for its target, and the relative expression and 769!
affinity of an individual target compared to the total pool of target sequences. miRNAs that 770!
! 32!
bind many or very abundant mRNAs will effectively be buffered within the cell, raising the 771!
likelihood that their effects on an individual protein may be minor. For example, patients with 772!
ICUAW show a 50-fold increase in the miRNAs but a much smaller reduction in UBTF 773!
(Connolly et al., 2017). Consequently, where there are smaller changes in miRNA level the 774!
effects on absolute target protein expression is likely to be close to undetectable within the 775!
variation present within the population. However, given the size of the change in pathway 776!
activity needed to cause significant wasting over time, the difference may be sufficient and 777!
account for associations of the miRNA with phenotype. This is also indicative of the subtlety 778!
of the response system consistent with an ongoing mechanism that continuously recalibrates 779!
itself, rather than responding only to intermittent major challenges. 780!
There are a multiple limitations to this model. Firstly, it can only be a partial story as it is 781!
based on the analysis of a subset of miRNAs in a limited number of diseases and on the 782!
analysis of a subset of the targets of those miRNAs. Consequently, we acknowledge that 783!
other miRNAs will contribute to susceptibility to wasting both positively and negatively. 784!
Secondly, miRNAs target multiple mRNAs and the degree to which any one target will be 785!
affected is determined not only by the relative concentrations of the miRNA and the 786!
individual target mRNA, but also by the relative concentrations of competitor species 787!
including other target mRNAs and long non-coding RNAs (lncRNAs) acting as miR-sponges 788!
(Cesana et al., 2011). Thirdly, the data are based on correlation analysis and the 789!
interventional experiments are transfection and over-expression. Quantification of transfected 790!
miRNAs is imprecise because a significant proportion of the transfected miRNA is 791!
sequestered in vesicles and so not functional (Thomson et al., 2013) and over-expression can 792!
increase miRNA levels above physiologically relevant concentrations. Consequently, all 793!
miRNA-expression experiments give is an indication of what the miRNA can do within a 794!
given context, not what it does in any other context. However, in defence of the model, it is 795!
! 33!
consistent across a range of human diseases and is accompanied by relevant changes in the 796!
expression of identified targets within those disease conditions. 797!
798!
VI. FUTURE QUESTIONS!799!
The model described in Section IV gives potential mechanisms for both disease-associated 800!
muscle wasting and variability in the response of individuals with the same disease severity. 801!
As such, it provides a framework to explain why patients with chronic disease waste, and 802!
why some are more likely to waste than others. However, there are many questions left 803!
unanswered as to the generalisability of the model. The data reviewed come, predominantly, 804!
from patients with cardio-respiratory diseases, and cannot therefore be uncritically extended 805!
to other groups of patients; thus more data from patients with other forms of cachexia, such 806!
as renal disease is urgently needed. Similarly understanding patterns of atrophy and 807!
hypertrophy in response to evolutionarily relevant stimuli such as starvation, exercise and 808!
hibernation would be relevant. It is also obviously not a complete picture. There are many 809!
miRNAs that change in the muscle of patients with COPD, the majority of which are 810!
suppressed compared to controls. Many of these changes are also likely to contribute to the 811!
overall phenotype. Some miRNAs will not change in response to the disease process but will 812!
control the susceptibility of patients to muscle wasting. For example, in addition to 813!
associations of miRNAs from imprinted regions of the genome with muscle loss, the data 814!
also identified a number of other miRNAs that were suppressed in patients who retained 815!
muscle mass compared with those who did not (Lewis et al., 2016). Many of these miRNAs 816!
are predicted to target inhibitors of the cell cycle or have been shown to contribute to a 817!
proliferative cell phenotype. These observations are consistent with the overall picture but 818!
represent increased resolution. 819!
! 34!
Of particular therapeutic interest are the mechanisms that elevate the expression of the 820!
miRNAs in the miR-424/542 cluster, because we predict that inhibiting this increase may 821!
reduce the ‘drive to wasting’ that is seen in COPD and aortic-surgery patients. The 822!
expression of this cluster appears to be co-ordinated, implying a single promoter, but as yet 823!
we have been unable to detect a single transcript containing both miR-424 and miR-542. 824!
Determining whether this is the case and whether inhibition of this cluster suppresses muscle 825!
wasting is therefore a key goal of future research. 826!
827!
VII. CONCLUSIONS!828!
(1) Muscle wasting, when severe, is a debilitating condition that reduces quality of life. This 829!
wasting results from small changes in the relative rates of protein synthesis and degradation 830!
that arise from a complex interaction of environment, genetics and epigenetics against a 831!
background of continuous protein turnover with a half-life in man of approximately 20 days 832!
(Poortmans et al., 2012). !833!
(2) The rate of protein turnover in muscle means that the degree of imbalance in protein 834!
synthesis and degradation that can lead to significant muscle loss over an extended period is 835!
small, suggesting that the system is capable of very fine tuning of protein regulation. 836!
Classical mechanisms seem implausible for this degree of control.!837!
(3) Recent data suggest that changes in the miRNA profile in muscle make an important 838!
contribution to the loss of muscle mass in patients with chronic diseases. These profiles 839!
contribute to muscle loss, both in response to ongoing chronic muscle wasting signals as well 840!
as to single bigger insults. The miRNAs regulate fundamental biochemical pathways and 841!
affect the capacity of muscle to make protein as well as the sensitivity of the muscle to 842!
anabolic and catabolic signals.!843!
! 35!
(4) Some of the individual variation in the susceptibility to muscle wasting arises from 844!
differences in the ability to regenerate muscle, probably because the rate of recruitment of 845!
new myonuclei represents a balance between proliferation to make sufficient nuclei and 846!
differentiation into the muscle fibre. Relative expression of miRNAs again contributes to this 847!
balance and may well be set during early development or as a component of imprinting. !848!
(5) Taken together, we suggest the complexity of the mechanism implies that its primary 849!
purpose is not regulation of muscle mass but rather that muscle loss is the clinical observation 850!
that derives from skeletal muscle’s role as an energy and amino acid bank for repair in other 851!
organs.!852!
853!
VIII. REFERENCES 854!
ABNEY, M., MCPEEK, M. S. & OBER, C. (2001). Broad and narrow heritabilities of 855!
quantitative traits in a founder population. American Journal of Human Genetics 68(5), 856!
1302–1307. 857!
ABRIGO, J., ELORZA, A. A., RIEDEL, C. A., VILOS, C., SIMON, F., CABRERA, D., ESTRADA, L. & 858!
CABELLO-VERRUGIO, C. (2018). Role of Oxidative Stress as Key Regulator of Muscle 859!
Wasting during Cachexia. Oxidative Medicine and Cellular Longevity 2018, 2063179. 860!
AGUSTI, A., CALVERLEY, P. M., CELLI, B., COXSON, H. O., EDWARDS, L. D., LOMAS, D. A., 861!
MACNEE, W., MILLER, B. E., RENNARD, S., SILVERMAN, E. K., TAL-SINGER, R., WOUTERS, E., 862!
YATES, J. C., VESTBO, J. & THE EVALUATION OF COPD LONGITUDINALLY TO IDENTIFY 863!
PREDICTIVE SURROGATE ENPOINTS (ECLIPSE) INVESTIGATORS (2010). Characterisation of 864!
COPD heterogeneity in the ECLIPSE cohort. Respiratory Research 11: 122. 865!
ALEXEYEV, M. F., LEDOUX, S. P. & WILSON, G. L. (2004). Mitochondrial DNA and aging. 866!
Clinical Science (London) 107(4), 355–364. 867!
! 36!
ANASTASI, S., GIORDANO, S., STHANDIER, O., GAMBAROTTA, G., MAIONE, R., COMOGLIO, P. 868!
& AMATI, P. (1997). A natural hepatocyte growth factor/scatter factor autocrine loop in 869!
myoblast cells and the effect of the constitutive Met kinase activation on myogenic 870!
differentiation. Journal of Cell Biology 137(5), 1057–1068. 871!
ANKER, S. D., PONIKOWSKI, P., VARNEY, S., CHUA, T. P., CLARK, A. L., WEBB-PEPLOE, K. M., 872!
HARRINGTON, D., KOX, W. J., POOLE-WILSON, P. A. & COATS, A. J. (1997). Wasting as 873!
independent risk factor for mortality in chronic heart failure. Lancet 349(9058), 1050–1053. 874!
ANKER, S. D. & SHARMA, R. (2002). The syndrome of cardiac cachexia. International 875!
Journal of Cardiology 85(1), 51–66. 876!
ARGILES, J. M., CAMPOS, N., LOPEZ-PEDROSA, J. M., RUEDA, R. & RODRIGUEZ-MANAS, L. 877!
(2016). Skeletal Muscle Regulates Metabolism via Interorgan Crosstalk: Roles in Health and 878!
Disease. Journal of the American Medical Directors Association 17(9), 789–796. 879!
BAERTLING, F., HAACK, T. B., RODENBURG, R. J., SCHAPER, J., SEIBT, A., STROM, T. M., 880!
MEITINGER, T., MAYATEPEK, E., HADZIK, B., SELCAN, G., PROKISCH, H. & DISTELMAIER, F. 881!
(2015). MRPS22 mutation causes fatal neonatal lactic acidosis with brain and heart 882!
abnormalities. Neurogenetics 16(3), 237–240. 883!
BEISEL, W. R. (1984). Metabolic effects of infection. Progress in Food and Nutrition 884!
Sciences 8(1–2), 43–75. 885!
BHAT, A. H., DAR, K. B., ANEES, S., ZARGAR, M. A., MASOOD, A., SOFI, M. A. & GANIE, S. A. 886!
(2015). Oxidative stress, mitochondrial dysfunction and neurodegenerative diseases; a 887!
mechanistic insight. Biomedical Pharmacotherapeutics 74, 101–110. 888!
BIOLO, G., FLEMING, R. Y., MAGGI, S. P., NGUYEN, T. T., HERNDON, D. N. & WOLFE, R. R. 889!
(2000). Inhibition of muscle glutamine formation in hypercatabolic patients. Clinical Science 890!
(London) 99(3), 189–194. 891!
! 37!
BISTRIAN, B. (2007). Systemic response to inflammation. Nutrition Reviews 65(12 Pt 2), 892!
S170–172. 893!
BLOCH, S. A., LEE, J. Y., SYBURRA, T., ROSENDAHL, U., GRIFFITHS, M. J., KEMP, P. R. & 894!
POLKEY, M. I. (2015). Increased expression of GDF-15 may mediate ICU-acquired weakness 895!
by down-regulating muscle microRNAs. Thorax 70(3), 219–228. 896!
BLOCH, S., LEE, J. Y., WORT, S. J., POLKEY, M. I., KEMP, P. R. & GRIFFITHS, M. (2013). 897!
Sustained elevation of circulating GDF-15 and a dynamic imbalance in mediators of muscle 898!
homeostasis are associated with the development of acute muscle wasting following cardiac 899!
surgery. Critical Care Medicine 41, 982–989. 900!
BLOCH, S., POLKEY, M. I., GRIFFITHS, M. & KEMP, P. (2012). Molecular mechanisms of 901!
intensive care unit-acquired weakness. European Respiratory Journal 39(4), 1000–1011. 902!
BODINE, S. C., LATRES, E., BAUMHUETER, S., LAI, V. K., NUNEZ, L., CLARKE, B. A., 903!
POUEYMIROU, W. T., PANARO, F. J., NA, E., DHARMARAJAN, K., PAN, Z. Q., VALENZUELA, D. 904!
M., DECHIARA, T. M., STITT, T. N., YANCOPOULOS, G. D. & D. J. GLASS. (2001). 905!
Identification of ubiquitin ligases required for skeletal muscle atrophy. Science 294(5547), 906!
1704–1708. 907!
BOETTGER, T., WUST, S., NOLTE, H. & BRAUN, T. (2014). The miR-206/133b cluster is 908!
dispensable for development, survival and regeneration of skeletal muscle. Skeletal Muscle 909!
4(1), 23. 910!
BOISVERT, F. M., VAN KONINGSBRUGGEN, S., NAVASCUES, J. & LAMOND, A. I. (2007). The 911!
multifunctional nucleolus. Nature Reviews Molecular Cellular Biology l 8(7), 574–585. 912!
BREEN, L., STOKES, K. A., CHURCHWARD-VENNE, T. A., MOORE, D. R., BAKER, S. K., SMITH, 913!
K., ATHERTON, P. J. & PHILLIPS, S. M. (2013). Two weeks of reduced activity decreases leg 914!
lean mass and induces "anabolic resistance" of myofibrillar protein synthesis in healthy 915!
elderly. Journal of Clinical Endocrinology and Metabolism 98(6), 2604–2612. 916!
! 38!
BREITBART, A., AUGER-MESSIER, M., MOLKENTIN, J. D. & HEINEKE, J. (2011). Myostatin 917!
from the heart: local and systemic actions in cardiac failure and muscle wasting. American 918!
Journal of Physiology Heart and Circulatory Physiology 300(6), H1973–1982. 919!
BRENNECKE, J., STARK, A., RUSSELL, R. B. & COHEN, S. M. (2005). Principles of microRNA-920!
target recognition. PLOS Biology 3(3), e85. 921!
BUCCI, L., YANI, S. L., FABBRI, C., BIJLSMA, A. Y., MAIER, A. B., MESKERS, C. G., NARICI, M. 922!
V., JONES, D. A., MCPHEE, J. S., SEPPET, E., GAPEYEVA, H., PAASUKE, M., SIPILA, S., 923!
KOVANEN, V., STENROTH, L., et al. (2013). Circulating levels of adipokines and IGF-1 are 924!
associated with skeletal muscle strength of young and old healthy subjects. Biogerontology 925!
14(3), 261–272. 926!
CACCHIARELLI, D., LEGNINI, I., MARTONE, J., CAZZELLA, V., D'AMICO, A., BERTINI, E. & 927!
BOZZONI, I. (2011). miRNAs as serum biomarkers for Duchenne muscular dystrophy. EMBO 928!
Molecular Medicine 3(5), 258–265. 929!
CASO, G., GARLICK, P. J., BALLOU, L. M., VOSSWINKEL, J. A., GELATO, M. C. & MCNURLAN, 930!
M. A. (2006). The increase in human muscle protein synthesis induced by food intake is 931!
similar when assessed with the constant infusion and flooding techniques. Journal of 932!
Nutrition 136(6), 1504–1510. 933!
CASTETS, P., BERTRAND, A. T., BEUVIN, M., FERRY, A., LE GRAND, F., CASTETS, M., CHAZOT, 934!
G., REDERSTORFF, M., KROL, A., LESCURE, A., ROMERO, N. B., GUICHENEY, P. & ALLAMAND, 935!
V. (2011). Satellite cell loss and impaired muscle regeneration in selenoprotein N deficiency. 936!
Human Molecular Genetetics 20(4), 694–704. 937!
CENTNER, T., YANO, J., KIMURA, E., MCELHINNY, A. S., PELIN, K., WITT, C. C., BANG, M. L., 938!
TROMBITAS, K., GRANZIER, H., GREGORIO, C. C., SORIMACHI, H. & LABEIT, S. (2001). 939!
Identification of muscle specific ring finger proteins as potential regulators of the titin kinase 940!
domain. Journal of Molecular Biology 306(4), 717–726. 941!
! 39!
CESANA, M., CACCHIARELLI, D., LEGNINI, I., SANTINI, T., STHANDIER, O., CHINAPPI, M., 942!
TRAMONTANO, A. & BOZZONI, I. (2011). A long noncoding RNA controls muscle 943!
differentiation by functioning as a competing endogenous RNA. Cell 147(2), 358–369. 944!
CHAN, E. K., IMAI, H., HAMEL, J. C. & TAN, E. M. (1991). Human autoantibody to RNA 945!
polymerase I transcription factor hUBF. Molecular identity of nucleolus organizer region 946!
autoantigen NOR-90 and ribosomal RNA transcription upstream binding factor. Journal of 947!
Experimental Medicine 174(5), 1239–1244. 948!
CHARBONNEAU, D. E., HANSON, E. D., LUDLOW, A. T., DELMONICO, M. J., HURLEY, B. F. & 949!
ROTH, S. M. (2008). ACE genotype and the muscle hypertrophic and strength responses to 950!
strength training. Medicine and Science in Sports and Exercise 40(4), 677–683. 951!
CHEN, J. F., CALLIS, T. E. & WANG, D. Z. (2009). microRNAs and muscle disorders. Journal 952!
of Cell Science 122(Pt 1), 13–20. 953!
CHEN, J. F., MANDEL, E. M., THOMSON, J. M., WU, Q., CALLIS, T. E., HAMMOND, S. M., 954!
CONLON, F. L. & WANG, D. Z. (2006). The role of microRNA-1 and microRNA-133 in 955!
skeletal muscle proliferation and differentiation. Nature Genetics 38(2), 228–233. 956!
CMARKO, D., SMIGOVA, J., MINICHOVA, L. & POPOV, A. (2008). Nucleolus: the ribosome 957!
factory. Histology and Histopathology 23(10), 1291–1298. 958!
COHEN, S., BRAULT, J. J., GYGI, S. P., GLASS, D. J., VALENZUELA, D. M., GARTNER, C., 959!
LATRES, E. & GOLDBERG, A. L. (2009). During muscle atrophy, thick, but not thin, filament 960!
components are degraded by MuRF1-dependent ubiquitylation. Journal of Cell Biology 961!
185(6), 1083–1095. 962!
COLLAMATI, A., MARZETTI, E., CALVANI, R., TOSATO, M., D'ANGELO, E., SISTO, A. N. & 963!
LANDI, F. (2016). Sarcopenia in heart failure: mechanisms and therapeutic strategies. Journal 964!
of Geriatric Cardiology 13(7), 615–624. 965!
! 40!
COMAI, L., TANESE, N. & TJIAN, R. (1992). The TATA-binding protein and associated factors 966!
are integral components of the RNA polymerase I transcription factor, SL1. Cell 68(5), 965–967!
976. 968!
CONNOLLY, M., GARFIELD, B. E., CROSBY, A., MORRELL, N. W., WORT, S. J. & KEMP, P. R. 969!
(2018). miR-322-5p targets IGF-1 and is suppressed in the heart of rats with pulmonary 970!
hypertension. FEBS Open Bio 8, 339–348. 971!
CONNOLLY, M., FARRE GARROS, R., PAUL, R., NATANEK, S. A., BLOCH, S., SAYER, A. A., 972!
PATEL, H. P., COOPER, C., GRIFFITHS, M. J., POLKEY, M. I. & KEMP, P. R. (2017). miR-424-5p 973!
reduces rRNA and protein synthesis in muscle wasting Journal of Sarcopenia Cachexia and 974!
Muscle, doi: 10.1002/JCSM12266. 975!
COOK, S. J. & MORLEY, S. J. (2007). Nutrient-responsive mTOR signalling grows on Sterile 976!
ground. Biochemical Journal 403(1), e1–3. 977!
DAI, Y., WANG, Y. M., ZHANG, W. R., LIU, X. F., LI, X., DING, X. B. & GUO, H. (2016). The 978!
role of microRNA-1 and microRNA-206 in the proliferation and differentiation of bovine 979!
skeletal muscle satellite cells. In Vitro Cellular Developmental Biology - Animal 52(1), 27–980!
34. 981!
DE BOER, M. D., SELBY, A., ATHERTON, P., SMITH, K., SEYNNES, O. R., MAGANARIS, C. N., 982!
MAFFULLI, N., MOVIN, T., NARICI, M. V. & RENNIE, M. J. (2007). The temporal responses of 983!
protein synthesis, gene expression and cell signalling in human quadriceps muscle and 984!
patellar tendon to disuse. Journal of Physiology 585(Pt 1), 241–251. 985!
DEHOUX, M., GOBIER, C., LAUSE, P., BERTRAND, L., KETELSLEGERS, J. M. & THISSEN, J. P. 986!
(2007). IGF-I does not prevent myotube atrophy caused by proinflammatory cytokines 987!
despite activation of Akt/Foxo and GSK-3beta pathways and inhibition of atrogin-1 mRNA. 988!
American Journal of Physiology Endocrinology and Metabolism 292(1), E145–150. 989!
! 41!
DENG, Z., CHEN, J. F. & WANG, D. Z. (2011). Transgenic overexpression of miR-133a in 990!
skeletal muscle. BMC Musculoskeletal Disorders 12, 115. 991!
DEY, B. K., PFEIFER, K. & DUTTA, A. (2014). The H19 long noncoding RNA gives rise to 992!
microRNAs miR-675-3p and miR-675-5p to promote skeletal muscle differentiation and 993!
regeneration. Genes and Development 28(5), 491–501. 994!
DIDERIKSEN, K., BOESEN, A. P., KRISTIANSEN, J. F., MAGNUSSON, S. P., SCHJERLING, P., 995!
HOLM, L. & KJAER, M. (2016). Skeletal muscle adaptation to immobilization and subsequent 996!
retraining in elderly men: No effect of anti-inflammatory medication. Experimental 997!
Gerontology 82, 8–18. 998!
DISSANAYAKE, E. & INOUE, Y. (2016). MicroRNAs in Allergic Disease. Current Allergy and 999!
Asthma Reports 16(9), 67. 1000!
DONALDSON, A., NATANEK, S. A., LEWIS, A., MAN, W. D., HOPKINSON, N. S., POLKEY, M. I. 1001!
& KEMP, P. R. (2013). Increased skeletal muscle-specific microRNA in the blood of patients 1002!
with COPD. Thorax 68, 1140–1149. 1003!
DOUCET, M., RUSSELL, A. P., LEGER, B., DEBIGARE, R., JOANISSE, D. R., CARON, M. A., 1004!
LEBLANC, P. & MALTAIS, F. (2007). Muscle atrophy and hypertrophy signaling in patients 1005!
with chronic obstructive pulmonary disease. American Journal of Respiratory and Critical 1006!
Care Medicine 176(3), 261–269. 1007!
DRUMMOND, M. J., MCCARTHY, J. J., FRY, C. S., ESSER, K. A. & RASMUSSEN, B. B. (2008). 1008!
Aging differentially affects human skeletal muscle microRNA expression at rest and after an 1009!
anabolic stimulus of resistance exercise and essential amino acids. American Journal of 1010!
Physiology Endocrinology and Metabolism 295(6), E1333–1340. 1011!
EGERMAN, M. A., CADENA, S. M., GILBERT, J. A., MEYER, A., NELSON, H. N., SWALLEY, S. E., 1012!
MALLOZZI, C., JACOBI, C., JENNINGS, L. L., CLAY, I., LAURENT, G., MA, S., BRACHAT, S., 1013!
! 42!
LACH-TRIFILIEFF, E., SHAVLAKADZE, T., et al. (2015). GDF11 Increases with Age and 1014!
Inhibits Skeletal Muscle Regeneration. Cell Metabolism 22(1), 164–174. 1015!
EMMERSON, P. J., WANG, F., DU, Y., LIU, Q., PICKARD, R. T., GONCIARZ, M. D., COSKUN, T., 1016!
HAMANG, M. J., SINDELAR, D. K., BALLMAN, K. K., FOLTZ, L. A., MUPPIDI, A., ALSINA-1017!
FERNANDEZ, J., BARNARD, G. C., TANG, J. X., et al. (2017). The metabolic effects of GDF15 1018!
are mediated by the orphan receptor GFRAL. Nature Medicine 23(10), 1215–1219. 1019!
ESSER, K. (2008). Regulation of mTOR signaling in skeletal muscle hypertrophy. Journal of 1020!
Musculoskeletal and Neuronal Interactions 8(4), 338–339. 1021!
EVANS, D. J., MURRAY, R. & KISSEBAH, A. H. (1984). Relationship between skeletal muscle 1022!
insulin resistance, insulin-mediated glucose disposal, and insulin binding. Effects of obesity 1023!
and body fat topography. Journal of Clinical Investigations 74(4), 1515–1525. 1024!
FARRE GARROS, R., PAUL, R., CONNOLLY, M., LEWIS, A., GARFIELD, B. E., NATANEK, S. A., 1025!
BLOCH, S., MOULY, V., GRIFFITHS, M. J., POLKEY, M. I. & KEMP, P. R. (2017). miR-542 1026!
Promotes Mitochondrial Dysfunction and SMAD Activity and is Raised in ICU Acquired 1027!
Weakness. American Journal of Respiratory and Critical Care Medicine 196, 1422–1433. 1028!
FARRE-GARROS, R., NATANEK, S. A., BLOCH, S., POLKEY, M. I. & KEMP, P. R. (2015). miR-1029!
542, a novel regulator of muscle mass and function. J Muscle Res Cell Motil 36, 593–594. 1030!
FEARON, K., EVANS, W. J. & ANKER, S. D. (2011). Myopenia-a new universal term for muscle 1031!
wasting. Journal of Cachexia Sarcopenia Muscle 2(1), 1–3. 1032!
FENG, Y., NIU, L. L., WEI, W., ZHANG, W. Y., LI, X. Y., CAO, J. H. & ZHAO, S. H. (2013). A 1033!
feedback circuit between miR-133 and the ERK1/2 pathway involving an exquisite 1034!
mechanism for regulating myoblast proliferation and differentiation. Cell Death and Disease 1035!
4, e934. 1036!
FIGUEIREDO, V. C., CALDOW, M. K., MASSIE, V., MARKWORTH, J. F., CAMERON-SMITH, D. & 1037!
BLAZEVICH, A. J. (2015). Ribosome biogenesis adaptation in resistance training-induced 1038!
! 43!
human skeletal muscle hypertrophy. American Journal of Physiology Endocrinology and 1039!
Metabolism 309(1), E72–83. 1040!
FORMAN, D. E., DANIELS, K. M., CAHALIN, L. P., ZAVIN, A., ALLSUP, K., CAO, P., 1041!
SANTHANAM, M., JOSEPH, J., ARENA, R., LAZZARI, A., SCHULZE, P. C. & LECKER, S. H. 1042!
(2014). Analysis of skeletal muscle gene expression patterns and the impact of functional 1043!
capacity in patients with systolic heart failure. Journal of Cardiac Failure 20(6), 422–430. 1044!
FREEMAN, C. M., MARTINEZ, C. H., TODT, J. C., MARTINEZ, F. J., HAN, M. K., THOMPSON, D. 1045!
L., MCCLOSKEY, L. & CURTIS, J. L. (2015). Acute exacerbations of chronic obstructive 1046!
pulmonary disease are associated with decreased CD4+ & CD8+ T cells and increased 1047!
growth & differentiation factor-15 (GDF-15) in peripheral blood. Respiratory Research 16, 1048!
94. 1049!
FRIEDRICHS, M., WIRSDOERFER, F., FLOHE, S. B., SCHNEIDER, S., WUELLING, M. & 1050!
VORTKAMP, A. (2011). BMP signaling balances proliferation and differentiation of muscle 1051!
satellite cell descendants. BMC Cell Biology 12, 26. 1052!
FRIMAN, G. & WESSLEN, L. (2000). Special feature for the Olympics: effects of exercise on 1053!
the immune system: infections and exercise in high-performance athletes. Immunology and 1054!
Cell Biology 78(5), 510–522. 1055!
FULSTER, S., TACKE, M., SANDEK, A., EBNER, N., TSCHOPE, C., DOEHNER, W., ANKER, S. D. & 1056!
VON HAEHLING, S. (2013). Muscle wasting in patients with chronic heart failure: results from 1057!
the studies investigating co-morbidities aggravating heart failure (SICA-HF). European 1058!
Heart Journal 34(7), 512–519. 1059!
GAO, S., DURSTINE, J. L., KOH, H. J., CARVER, W. E., FRIZZELL, N. & CARSON, J. A. (2017). 1060!
Acute myotube protein synthesis regulation by IL-6-related cytokines. American Journal of 1061!
Physiology Cell Physiology 313(5), C487–C500. 1062!
! 44!
GEORGE, M., JENA, A., SRIVATSAN, V., MUTHUKUMAR, R. & DHANDAPANI, V. E. (2016). GDF 1063!
15--A Novel Biomarker in the Offing for Heart Failure. Current Cardiology Reviews 12(1), 1064!
37–46. 1065!
GIELEN, S., SANDRI, M., KOZAREZ, I., KRATZSCH, J., TEUPSER, D., THIERY, J., ERBS, S., 1066!
MANGNER, N., LENK, K., HAMBRECHT, R., SCHULER, G. & ADAMS, V. (2012). Exercise 1067!
training attenuates MuRF-1 expression in the skeletal muscle of patients with chronic heart 1068!
failure independent of age: the randomized Leipzig Exercise Intervention in Chronic Heart 1069!
Failure and Aging catabolism study. Circulation 125(22), 2716–2727. 1070!
GLASS, D. J. (2010). PI3 kinase regulation of skeletal muscle hypertrophy and atrophy. 1071!
Current Topics in Microbiology and Immunology 346, 267–278. 1072!
GLASS, D. J. (2005). Skeletal muscle hypertrophy and atrophy signaling pathways. 1073!
International Journal of Biochemistry and Cell Biology 37(10), 1974–1984. 1074!
GOBET, C. & NAEF, F. (2017). Ribosome profiling and dynamic regulation of translation in 1075!
mammals. Current Opinion Genetics and Development 43, 120–127. 1076!
GORE, D. C. & JAHOOR, F. (2000). Deficiency in peripheral glutamine production in pediatric 1077!
patients with burns. Journal of Burn Care Rehabilitation 21(2), 171; discussion 172–177. 1078!
GROBET, L., MARTIN, L. J., PONCELET, D., PIROTTIN, D., BROUWERS, B., RIQUET, J., 1079!
SCHOEBERLEIN, A., DUNNER, S., MENISSIER, F., MASSABANDA, J., FRIES, R., HANSET, R. & 1080!
GEORGES, M. (1997). A deletion in the bovine myostatin gene causes the double-muscled 1081!
phenotype in cattle. Nature Genetics 17(1), 71–74. 1082!
GULLER, I. & RUSSELL, A. P. (2010). MicroRNAs in skeletal muscle: their role and regulation 1083!
in development, disease and function. Journal of Physiology 588(Pt 21), 4075–4087. 1084!
HADDAD, F., ZALDIVAR, F., COOPER, D. M. & ADAMS, G. R. (2005). IL-6-induced skeletal 1085!
muscle atrophy. Journal of Applied Physiology (1985) 98(3), 911–917. 1086!
! 45!
HAMMERS, D. W., MERSCHAM-BANDA, M., HSIAO, J. Y., ENGST, S., HARTMAN, J. J. & 1087!
SWEENEY, H. L. (2017). Supraphysiological levels of GDF11 induce striated muscle atrophy. 1088!
EMBO Molecular Medicine 9(4), 531–544. 1089!
HARPER, S. C., BRACK, A., MACDONNELL, S., FRANTI, M., OLWIN, B. B., BAILEY, B. A., 1090!
RUDNICKI, M. A. & HOUSER, S. R. (2016). Is Growth Differentiation Factor 11 a Realistic 1091!
Therapeutic for Aging-Dependent Muscle Defects? Circulation Research 118(7), 1143–1150; 1092!
discussion 1150. 1093!
HARRIDGE, S. D. (2007). Plasticity of human skeletal muscle: gene expression to in vivo 1094!
function. Experimental Physiology 92(5), 783–797. 1095!
HE, L. & HANNON, G. J. (2004). MicroRNAs: small RNAs with a big role in gene regulation. 1096!
Nature Reviews Genetics 5(7), 522–531. 1097!
HEIKKINEN, P. T., NUMMELA, M., LEIVONEN, S. K., WESTERMARCK, J., HILL, C. S., KAHARI, 1098!
V. M. & JAAKKOLA, P. M. (2010). Hypoxia-activated Smad3-specific dephosphorylation by 1099!
PP2A. Journal of Biological Chemistry 285(6), 3740–3749. 1100!
HOFFER, L. J. & BISTRIAN, B. R. (2016). Nutrition in critical illness: a current conundrum. 1101!
F1000Research 5, 2531. 1102!
HOPKINSON, N. S., NICKOL, A. H., PAYNE, J., HAWE, E., MAN, W. D., MOXHAM, J., 1103!
MONTGOMERY, H. & POLKEY, M. I. (2004). Angiotensin converting enzyme genotype and 1104!
strength in chronic obstructive pulmonary disease. American Journal of Respiratory and 1105!
Critical Care Medicine 170(4), 395–399. 1106!
HOYO, C., DALTVEIT, A. K., IVERSEN, E., BENJAMIN-NEELON, S. E., FUEMMELER, B., 1107!
SCHILDKRAUT, J., MURTHA, A. P., OVERCASH, F., VIDAL, A. C., WANG, F., HUANG, Z., 1108!
KURTZBERG, J., SEEWALDT, V., FORMAN, M., JIRTLE, R. L., et al. (2014). Erythrocyte folate 1109!
concentrations, CpG methylation at genomically imprinted domains, and birth weight in a 1110!
multiethnic newborn cohort. Epigenetics 9(8), 1120–1130. 1111!
! 46!
IADEVAIA, V., LIU, R. & PROUD, C. G. (2014). mTORC1 signaling controls multiple steps in 1112!
ribosome biogenesis. Seminars in Cell and Developmental Biology 36, 113–120. 1113!
JACKSON, R. J., HELLEN, C. U. & PESTOVA, T. V. (2010). The mechanism of eukaryotic 1114!
translation initiation and principles of its regulation. Nature Reviews Molecular and Cellular 1115!
Biology 11(2), 113–127. 1116!
JEANPLONG, F., BASS, J. J., SMITH, H. K., KIRK, S. P., KAMBADUR, R., SHARMA, M. & 1117!
OLDHAM, J. M. (2003). Prolonged underfeeding of sheep increases myostatin and myogenic 1118!
regulatory factor Myf-5 in skeletal muscle while IGF-I and myogenin are repressed. Journal 1119!
of Endocrinology 176(3), 425–437. 1120!
JESPERSEN, J. G., NEDERGAARD, A., ANDERSEN, L. L., SCHJERLING, P. & ANDERSEN, J. L. 1121!
(2011a). Myostatin expression during human muscle hypertrophy and subsequent atrophy: 1122!
increased myostatin with detraining. Scandinavian Journal of Medicine and Science in Sports 1123!
21(2), 215–223. 1124!
JESPERSEN, J. G., NEDERGAARD, A., REITELSEDER, S., MIKKELSEN, U. R., DIDERIKSEN, K. J., 1125!
AGERGAARD, J., KREINER, F., POTT, F. C., SCHJERLING, P. & KJAER, M. (2011b). Activated 1126!
protein synthesis and suppressed protein breakdown signaling in skeletal muscle of critically 1127!
ill patients. PLOS One 6(3), e18090. 1128!
JOHNEN, H., LIN, S., KUFFNER, T., BROWN, D. A., TSAI, V. W., BAUSKIN, A. R., WU, L., 1129!
PANKHURST, G., JIANG, L., JUNANKAR, S., HUNTER, M., FAIRLIE, W. D., LEE, N. J., ENRIQUEZ, 1130!
R. F., BALDOCK, P. A., et al. (2007). Tumor-induced anorexia and weight loss are mediated 1131!
by the TGF-beta superfamily cytokine MIC-1. Nature Medicine 13(11), 1333–1340. 1132!
KAYSEN, G. A. (2004). Inflammation: cause of vascular disease and malnutrition in dialysis 1133!
patients. Seminars in Nephrology 24(5), 431–436. 1134!
KEMP, P. & NATANEK, A. (2017). Epigenetics and Susceptibility to Muscle Wasting in 1135!
COPD. Arch Bronconeumol 53(7), 364–365. 1136!
! 47!
KENIRY, A., OXLEY, D., MONNIER, P., KYBA, M., DANDOLO, L., SMITS, G. & REIK, W. (2012). 1137!
The H19 lincRNA is a developmental reservoir of miR-675 that suppresses growth and Igf1r. 1138!
Nature Cell Biology 14(7), 659–665. 1139!
KIM, H. K., LEE, Y. S., SIVAPRASAD, U., MALHOTRA, A. & DUTTA, A. (2006). Muscle-specific 1140!
microRNA miR-206 promotes muscle differentiation. Journal of Cell Biology 174(5), 677–1141!
687. 1142!
KNEZEVIC, I., PATEL, A., SUNDARESAN, N. R., GUPTA, M. P., SOLARO, R. J., NAGALINGAM, R. 1143!
S. & GUPTA, M. (2012). A novel cardiomyocyte-enriched microRNA, miR-378, targets 1144!
insulin-like growth factor 1 receptor: implications in postnatal cardiac remodeling and cell 1145!
survival. Journal of Biological Chemistry 287(16), 12913–12926. 1146!
KYTHREOTIS, P., KOKKINI, A., AVGEROPOULOU, S., HADJIOANNOU, A., ANASTASAKOU, E., 1147!
RASIDAKIS, A. & BAKAKOS, P. (2009). Plasma leptin and insulin-like growth factor I levels 1148!
during acute exacerbations of chronic obstructive pulmonary disease. BMC Pulmonary 1149!
Medicine 9, 11. 1150!
LATRES, E., AMINI, A. R., AMINI, A. A., GRIFFITHS, J., MARTIN, F. J., WEI, Y., LIN, H. C., 1151!
YANCOPOULOS, G. D. & GLASS, D. J. (2005). Insulin-like growth factor-1 (IGF-1) inversely 1152!
regulates atrophy-induced genes via the phosphatidylinositol 3-kinase/Akt/mammalian target 1153!
of rapamycin (PI3K/Akt/mTOR) pathway. Journal of Biological Chemistry 280(4), 2737–1154!
2744. 1155!
LEE, J. Y., DONALDSON, A. V., LEWIS, A., NATANEK, S. A., POLKEY, M. I. & KEMP, P. R. 1156!
(2017). Circulating miRNAs from imprinted genomic regions are associated with peripheral 1157!
muscle strength in COPD patients. European Respiratory Journal 49(4). 1158!
LEE, J. Y., LORI, D., WELLS, D. J. & KEMP, P. R. (2015). FHL1 activates myostatin signalling 1159!
in skeletal muscle and promotes atrophy. FEBS Open Bio 5, 753–762. 1160!
! 48!
LEE, J. Y., HOPKINSON, N. S. & KEMP, P. R. (2011). Myostatin induces autophagy in skeletal 1161!
muscle in vitro. Biochemical and Biophysical Research Communications 415(4), 632–636. 1162!
LEFAUCHEUR, J. P. & SEBILLE, A. (1995). Muscle regeneration following injury can be 1163!
modified in vivo by immune neutralization of basic fibroblast growth factor, transforming 1164!
growth factor beta 1 or insulin-like growth factor I. Journal of Neuroimmunology 57(1–2), 1165!
85–91. 1166!
LEMIRE, B. B., DEBIGARE, R., DUBE, A., THERIAULT, M. E., COTE, C. H. & MALTAIS, F. 1167!
(2012). MAPK signaling in the quadriceps of patients with chronic obstructive pulmonary 1168!
disease. Journal of Applied Physiology (1985) 113(1), 159–166. 1169!
LEWIS, A., LEE, J. Y., DONALDSON, A. V., NATANEK, S. A., VAIDYANATHAN, S., MAN, W. D., 1170!
HOPKINSON, N. S., SAYER, A. A., PATEL, H. P., COOPER, C., SYDDALL, H., POLKEY, M. I. & 1171!
KEMP, P. R. (2016). Increased expression of H19/miR-675 is associated with a low fat free 1172!
mass index in patients with COPD Jorunal of Cachexia Sarcopenia and Muscle 7, 330–344. 1173!
LEWIS, A., RIDDOCH-CONTRERAS, J., NATANEK, S. A., DONALDSON, A., MAN, W. D., 1174!
MOXHAM, J., HOPKINSON, N. S., POLKEY, M. I. & KEMP, P. R. (2012). Downregulation of the 1175!
serum response factor/miR-1 axis in the quadriceps of patients with COPD. Thorax 67(1), 1176!
26–34. 1177!
LIGHTFOOT, A., MCARDLE, A. & GRIFFITHS, R. D. (2009). Muscle in defense. Critical Care 1178!
Medicine 37(10 Suppl), S384–390. 1179!
LIN, S., CHEUNG, W. K., CHEN, S., LU, G., WANG, Z., XIE, D., LI, K., LIN, M. C. & KUNG, H. F. 1180!
(2010). Computational identification and characterization of primate-specific microRNAs in 1181!
human genome. Computational Biological Chemistry 34(4), 232–241. 1182!
LIU, N., BEZPROZVANNAYA, S., WILLIAMS, A. H., QI, X., RICHARDSON, J. A., BASSEL-DUBY, 1183!
R. & OLSON, E. N. (2008). microRNA-133a regulates cardiomyocyte proliferation and 1184!
! 49!
suppresses smooth muscle gene expression in the heart. Genes and Development 22(23), 1185!
3242–3254. 1186!
LLOBET-NAVAS, D., RODRIGUEZ-BARRUECO, R., CASTRO, V., UGALDE, A. P., SUMAZIN, P., 1187!
JACOB-SENDLER, D., DEMIRCAN, B., CASTILLO-MARTIN, M., PUTCHA, P., MARSHALL, N., 1188!
VILLAGRASA, P., CHAN, J., SANCHEZ-GARCIA, F., PE'ER, D., RABADAN, R., et al. (2014). The 1189!
miR-424(322)/503 cluster orchestrates remodeling of the epithelium in the involuting 1190!
mammary gland. Genes and Development 28(7), 765–782. 1191!
LOFFREDO, F. S., STEINHAUSER, M. L., JAY, S. M., GANNON, J., PANCOAST, J. R., 1192!
YALAMANCHI, P., SINHA, M., DALL'OSSO, C., KHONG, D., SHADRACH, J. L., MILLER, C. M., 1193!
SINGER, B. S., STEWART, A., PSYCHOGIOS, N., GERSZTEN, R. E., et al. (2013). Growth 1194!
differentiation factor 11 is a circulating factor that reverses age-related cardiac hypertrophy. 1195!
Cell 153(4), 828–839. 1196!
LOKIREDDY, S., WIJESOMA, I. W., SZE, S. K., MCFARLANE, C., KAMBADUR, R. & SHARMA, M. 1197!
(2012). Identification of atrogin-1-targeted proteins during the myostatin-induced skeletal 1198!
muscle wasting. American Journal of Physiology Cell Physiology 303(5), C512–529. 1199!
MA, G., WANG, Y., LI, Y., CUI, L., ZHAO, Y., ZHAO, B. & LI, K. (2015). MiR-206, a key 1200!
modulator of skeletal muscle development and disease. International Journal of Biological 1201!
Sciences 11(3), 345–352. 1202!
MAHMOUDABADY, M., MATHIEU, M., DEWACHTER, L., HADAD, I., RAY, L., JESPERS, P., 1203!
BRIMIOULLE, S., NAEIJE, R. & MCENTEE, K. (2008). Activin-A, transforming growth factor-1204!
beta, and myostatin signaling pathway in experimental dilated cardiomyopathy. Journal of 1205!
Cardiac Failure 14(8), 703–709. 1206!
MAN, W. D., NATANEK, S. A., RIDDOCH-CONTRERAS, J., LEWIS, A., MARSH, G. S., KEMP, P. R. 1207!
& POLKEY, M. I. (2010). Quadriceps myostatin expression in COPD. European Respiratory 1208!
Journal 36(3), 686–688. 1209!
! 50!
MAO, G., LEE, S., ORTEGA, J., GU, L. & LI, G. M. (2012). Modulation of microRNA 1210!
processing by mismatch repair protein MutLalpha. Cell Research 22(6), 973–985. 1211!
MASIERO, E. & SANDRI, M. (2010). Autophagy inhibition induces atrophy and myopathy in 1212!
adult skeletal muscles. Autophagy 6(2), 307–309. 1213!
MCCARTHY, J. J. (2008). MicroRNA-206: the skeletal muscle-specific myomiR. Biochimica 1214!
et Biophysica Acta 1779(11), 682–691. 1215!
MCCORMICK, R., CHINDER, C., MCARDLE, A. & GOLJANEK-WHYSALL, K. (2017). The role of 1216!
miR-378 in sarcopenia. Journal of Muscle Research and Cell Motility 38, 79. 1217!
MCCROSKERY, S., THOMAS, M., MAXWELL, L., SHARMA, M. & KAMBADUR, R. (2003). 1218!
Myostatin negatively regulates satellite cell activation and self-renewal. Journal of Cell 1219!
Biology 162(6), 1135–1147. 1220!
MELOV, S., TARNOPOLSKY, M. A., BECKMAN, K., FELKEY, K. & HUBBARD, A. (2007). 1221!
Resistance exercise reverses aging in human skeletal muscle. PLOS One 2(5), e465. 1222!
MILLER, C., SAADA, A., SHAUL, N., SHABTAI, N., BEN-SHALOM, E., SHAAG, A., 1223!
HERSHKOVITZ, E. & ELPELEG, O. (2004). Defective mitochondrial translation caused by a 1224!
ribosomal protein (MRPS16) mutation. Annals of Neurology 56(5), 734–738. 1225!
MITTENDORFER, B., ANDERSEN, J. L., PLOMGAARD, P., SALTIN, B., BABRAJ, J. A., SMITH, K. & 1226!
RENNIE, M. J. (2005). Protein synthesis rates in human muscles: neither anatomical location 1227!
nor fibre-type composition are major determinants. Journal of Physiology 563(Pt 1), 203–1228!
211. 1229!
MO, K., RAZAK, Z., RAO, P., YU, Z., ADACHI, H., KATSUNO, M., SOBUE, G., LIEBERMAN, A. P., 1230!
WESTWOOD, J. T. & MONKS, D. A. (2010). Microarray analysis of gene expression by skeletal 1231!
muscle of three mouse models of Kennedy disease/spinal bulbar muscular atrophy. PLOS 1232!
One 5(9), e12922. 1233!
! 51!
MOSHER, D. S., QUIGNON, P., BUSTAMANTE, C. D., SUTTER, N. B., MELLERSH, C. S., PARKER, 1234!
H. G. & OSTRANDER, E. A. (2007). A mutation in the myostatin gene increases muscle mass 1235!
and enhances racing performance in heterozygote dogs. PLOS Genetics 3(5), e79. 1236!
MOTOHASHI, N., ALEXANDER, M. S., SHIMIZU-MOTOHASHI, Y., MYERS, J. A., KAWAHARA, G. 1237!
& KUNKEL, L. M. (2013). Regulation of IRS1/Akt insulin signaling by microRNA-128a 1238!
during myogenesis. Journal of Cell Science 126(Pt 12), 2678–2691. 1239!
MOYLAN, J. S. & REID, M. B. (2007). Oxidative stress, chronic disease, and muscle wasting. 1240!
Muscle and Nerve 35(4), 411–429. 1241!
MUSARO, A., MCCULLAGH, K., PAUL, A., HOUGHTON, L., DOBROWOLNY, G., MOLINARO, M., 1242!
BARTON, E. R., SWEENEY, H. L. & ROSENTHAL, N. (2001). Localized Igf-1 transgene 1243!
expression sustains hypertrophy and regeneration in senescent skeletal muscle. Nature 1244!
Genetics 27(2), 195–200. 1245!
NATANEK, S. A., GOSKER, H. R., SLOT, I. G., MARSH, G. S., HOPKINSON, N. S., MAN, W. D., 1246!
TAL-SINGER, R., MOXHAM, J., KEMP, P. R., SCHOLS, A. M. & POLKEY, M. I. (2013a). 1247!
Heterogeneity of quadriceps muscle phenotype in chronic obstructive pulmonary disease 1248!
(Copd); implications for stratified medicine? Muscle and Nerve 48(4), 488–497. 1249!
NATANEK, S. A., RIDDOCH-CONTRERAS, J., MARSH, G. S., HOPKINSON, N. S., MOXHAM, J., 1250!
MAN, W. D., KEMP, P. R. & POLKEY, M. I. (2013b). MuRF-1 and atrogin-1 protein expression 1251!
and quadriceps fiber size and muscle mass in stable patients with COPD. Journal of Chronic 1252!
Obstructive Pulmonary Disease 10(5), 618–624. 1253!
NEDELEC, B., DE OLIVEIRA, A. & GARREL, D. R. (2003). Acute phase modulation of systemic 1254!
insulin-like growth factor-1 and its binding proteins after major burn injuries. Critical Care 1255!
Medicine 31(6), 1794–1801. 1256!
! 52!
NIELSEN, S., SCHEELE, C., YFANTI, C., AKERSTROM, T., NIELSEN, A. R., PEDERSEN, B. K. & 1257!
LAYE, M. J. (2010). Muscle specific microRNAs are regulated by endurance exercise in 1258!
human skeletal muscle. Journal of Physiology 588(Pt 20), 4029–4037. 1259!
PARK, K. H. (2015). Mechanisms of Muscle Denervation in Aging: Insights from a Mouse 1260!
Model of Amyotrophic Lateral Sclerosis. Aging Diseases 6(5), 380–389. 1261!
PATEL, M. S., LEE, J. Y., BAZ, M., WELLS, C. E., BLOCH, S. A. A., LEWIS, A., DONALDSON, A. 1262!
V., GARFIELD, B. E., HOPKINSON, N. S., NATANEK, S. A., MAN, W. D.-C., WELLS, D. J., 1263!
BAKER, E. H., POLKEY, M. I. & KEMP, P. R. (2016). GDF-15 is associated with muscle mass 1264!
in COPD and promotes muscle wasting in vivo. Journal of Cachexia Sarcopenia and Muscl 1265!
7, 436–448. 1266!
PAUL, R., LEE, J., DONALDSON, A. V., CONNOLLY, M., SHARIF, M., NATANEK, S. A., 1267!
ROSENDAHL, U., POLKEY, M. I., GRIFFITHS, M. & KEMP, P. R. (2018). miR-422a suppresses 1268!
SMAD4 protein expression and promotes resistance to muscle loss. Journal of Cachexia 1269!
Sarcopenia Muscle 9(1), 119–128. 1270!
PAVO, N., WURM, R., NEUHOLD, S., ADLBRECHT, C., VILA, G., STRUNK, G., CLODI, M., RESL, 1271!
M., BRATH, H., PRAGER, R., LUGER, A., PACHER, R. & HULSMANN, M. (2016). GDF-15 Is 1272!
Associated with Cancer Incidence in Patients with Type 2 Diabetes. Clinical Chemistry 1273!
62(12), 1612–1620. 1274!
PERALTA, S., WANG, X. & MORAES, C. T. (2012). Mitochondrial transcription: lessons from 1275!
mouse models. Biochimica et Biophysica Acta 1819(9–10), 961–969. 1276!
PERRY, R. P. (2007). Balanced production of ribosomal proteins. Gene 401(1–2), 1–3. 1277!
PHILIPPOU, A., PAPAGEORGIOU, E., BOGDANIS, G., HALAPAS, A., SOURLA, A., MARIDAKI, M., 1278!
PISSIMISSIS, N. & KOUTSILIERIS, M. (2009). Expression of IGF-1 isoforms after exercise-1279!
induced muscle damage in humans: characterization of the MGF E peptide actions in vitro. In 1280!
Vivo 23(4), 567–575. 1281!
! 53!
PHILLIPS, S. M., GLOVER, E. I. & RENNIE, M. J. (2009). Alterations of protein turnover 1282!
underlying disuse atrophy in human skeletal muscle. Journal of Applied Physiology (1985) 1283!
107(3), 645–654. 1284!
PLANT, P. J., BROOKS, D., FAUGHNAN, M., BAYLEY, T., BAIN, J., SINGER, L., CORREA, J., 1285!
PEARCE, D., BINNIE, M. & BATT, J. (2010). Cellular markers of muscle atrophy in chronic 1286!
obstructive pulmonary disease. American Journal of Respiratory Cell and Molecular Biology 1287!
42(4), 461–471. 1288!
POCOCK, S. J., MCMURRAY, J. J., DOBSON, J., YUSUF, S., GRANGER, C. B., MICHELSON, E. L., 1289!
OSTERGREN, J., PFEFFER, M. A., SOLOMON, S. D., ANKER, S. D. & SWEDBERG, K. B. (2008). 1290!
Weight loss and mortality risk in patients with chronic heart failure in the candesartan in 1291!
heart failure: assessment of reduction in mortality and morbidity (CHARM) programme. 1292!
European Heart Journal 29(21), 2641–2650. 1293!
POLKEY, M. I., PRAESTGAARD, J., BERWICK, A., FRANSSEN, F. M. E., SINGH, D., STEINER, M. 1294!
C., CASABURI, R., TILLMANN, H. C., LACH-TRIFILIEFF, E., ROUBENOFF, R. & ROOKS, D. S. 1295!
(2018). Activin Type II Receptor Blockade for Treatment of Muscle Depletion in COPD: A 1296!
Randomized Trial. American Journal of Respiratory and Critical Care Medicine, DOI: 1297!
10.1164/rccm.20180210286OC. 1298!
POORTMANS, J. R., CARPENTIER, A., PEREIRA-LANCHA, L. O. & LANCHA, A., JR. (2012). 1299!
Protein turnover, amino acid requirements and recommendations for athletes and active 1300!
populations. Brazilian Journal of Medical and Biolical Research 45(10), 875–890. 1301!
POWERS, S. K., SMUDER, A. J. & JUDGE, A. R. (2012). Oxidative stress and disuse muscle 1302!
atrophy: cause or consequence? Current Opinion in Clinical Nutrition and Metababolic Care 1303!
15(3), 240–245. 1304!
PREISER, J. C., ICHAI, C., ORBAN, J. C. & GROENEVELD, A. B. (2014). Metabolic response to 1305!
the stress of critical illness. British Journal of Anaesthesia 113(6), 945–954. 1306!
! 54!
PUENTE-MAESTU, L., PEREZ-PARRA, J., GODOY, R., MORENO, N., TEJEDOR, A., GONZALEZ-1307!
ARAGONESES, F., BRAVO, J. L., ALVAREZ, F. V., CAMANO, S. & AGUSTI, A. (2009). Abnormal 1308!
mitochondrial function in locomotor and respiratory muscles of COPD patients. European 1309!
Respiratory Journal 33(5), 1045–1052. 1310!
PUIG-VILANOVA, E., MARTINEZ-LLORENS, J., AUSIN, P., ROCA, J., GEA, J. & BARREIRO, E. 1311!
(2015). Quadriceps muscle weakness and atrophy are associated with a differential epigenetic 1312!
profile in advanced COPD. Clinical Science (London) 128(12), 905–921. 1313!
PUIG-VILANOVA, E., AUSIN, P., MARTINEZ-LLORENS, J., GEA, J. & BARREIRO, E. (2014). Do 1314!
epigenetic events take place in the vastus lateralis of patients with mild chronic obstructive 1315!
pulmonary disease? PLOS One 9(7), e102296. 1316!
QUEVEDO, C., ALCAZAR, A. & SALINAS, M. (2000). Two different signal transduction 1317!
pathways are implicated in the regulation of initiation factor 2B activity in insulin-like 1318!
growth factor-1-stimulated neuronal cells. Journal of Biological Chemistry 275(25), 19192–1319!
19197. 1320!
REEDS, P. J., FJELD, C. R. & JAHOOR, F. (1994). Do the differences between the amino acid 1321!
compositions of acute-phase and muscle proteins have a bearing on nitrogen loss in traumatic 1322!
states? Journal of Nutrition 124(6), 906–910. 1323!
REISZ-PORSZASZ, S., BHASIN, S., ARTAZA, J. N., SHEN, R., SINHA-HIKIM, I., HOGUE, A., 1324!
FIELDER, T. J. & GONZALEZ-CADAVID, N. F. (2003). Lower skeletal muscle mass in male 1325!
transgenic mice with muscle-specific overexpression of myostatin. American Journal of 1326!
Physiology Endocrinology and Metabolism 285(4), E876–888. 1327!
REMELS, A. H., GOSKER, H. R., LANGEN, R. C. & SCHOLS, A. M. (2013). The mechanisms of 1328!
cachexia underlying muscle dysfunction in COPD. Journal of Applied Physiology (1985) 1329!
114(9), 1253–1262. 1330!
! 55!
RHODES, C. J., WHARTON, J. & WILKINS, M. R. (2013). Pulmonary hypertension: biomarkers. 1331!
Handbook of Experimental Pharmacology 218, 77–103. 1332!
RIDDOCH-CONTRERAS, J., GEORGE, T., NATANEK, S. A., MARSH, G. S., HOPKINSON, N. S., 1333!
TAL-SINGER, R., KEMP, P. & POLKEY, M. I. (2012). p38 mitogen-activated protein kinase is 1334!
not activated in the quadriceps of patients with stable chronic obstructive pulmonary disease. 1335!
Journal of Chronic Obstructive Pulmonary Disease 9(2), 142–150. 1336!
RISTOW, M., ZARSE, K., OBERBACH, A., KLOTING, N., BIRRINGER, M., KIEHNTOPF, M., 1337!
STUMVOLL, M., KAHN, C. R. & BLUHER, M. (2009). Antioxidants prevent health-promoting 1338!
effects of physical exercise in humans. Proceedings of the National Academy of Sciences U S 1339!
A 106(21), 8665–8670. 1340!
ROMANELLO, V. & SANDRI, M. (2013). Mitochondrial biogenesis and fragmentation as 1341!
regulators of protein degradation in striated muscles. Journal of Molecular and Cellular 1342!
Cardiology 55, 64–72. 1343!
ROMANELLO, V. & SANDRI, M. (2010). Mitochondrial biogenesis and fragmentation as 1344!
regulators of muscle protein degradation. Current Hypertension Reports 12(6), 433–439. 1345!
ROMMEL, C., BODINE, S. C., CLARKE, B. A., ROSSMAN, R., NUNEZ, L., STITT, T. N., 1346!
YANCOPOULOS, G. D. & GLASS, D. J. (2001). Mediation of IGF-1-induced skeletal myotube 1347!
hypertrophy by PI(3)K/Akt/mTOR and PI(3)K/Akt/GSK3 pathways. Nature Cell Biology 1348!
3(11), 1009–1013. 1349!
ROTH, E. (2007). Immune and cell modulation by amino acids. Clin Nutr 26(5), 535–544. 1350!
ROWLAND, L. A., BAL, N. C. & PERIASAMY, M. (2015). The role of skeletal-muscle-based 1351!
thermogenic mechanisms in vertebrate endothermy. Biological Reviews Cambridge 1352!
Philosophical Society 90(4), 1279–1297. 1353!
! 56!
SAADA, A., SHAAG, A., ARNON, S., DOLFIN, T., MILLER, C., FUCHS-TELEM, D., LOMBES, A. & 1354!
ELPELEG, O. (2007). Antenatal mitochondrial disease caused by mitochondrial ribosomal 1355!
protein (MRPS22) mutation. Journal of Medical Genetics 44(12), 784–786. 1356!
SAEKI, H., HAMADA, M. & HIWADA, K. (2002). Circulating levels of insulin-like growth 1357!
factor-1 and its binding proteins in patients with hypertrophic cardiomyopathy. Circulation 1358!
Journal 66(7), 639–644. 1359!
SANDRI, M. (2008). Signaling in muscle atrophy and hypertrophy. Physiology (Bethesda) 23, 1360!
160–170. 1361!
SCHERTZER, J. D. & LYNCH, G. S. (2006). Comparative evaluation of IGF-I gene transfer and 1362!
IGF-I protein administration for enhancing skeletal muscle regeneration after injury. Gene 1363!
Therapy 13(23), 1657–1664. 1364!
SCHIAFFINO, S., DYAR, K. A., CICILIOT, S., BLAAUW, B. & SANDRI, M. (2013). Mechanisms 1365!
regulating skeletal muscle growth and atrophy. FEBS Journal 280(17), 4294–4314. 1366!
SCHOLS, A. M., BROEKHUIZEN, R., WELING-SCHEEPERS, C. A. & WOUTERS, E. F. (2005). 1367!
Body composition and mortality in chronic obstructive pulmonary disease. American Journal 1368!
of Clinical Nutrition 82(1), 53–59. 1369!
SCHUELKE, M., WAGNER, K. R., STOLZ, L. E., HUBNER, C., RIEBEL, T., KOMEN, W., BRAUN, 1370!
T., TOBIN, J. F. & LEE, S. J. (2004). Myostatin mutation associated with gross muscle 1371!
hypertrophy in a child. New England Journal of Medicine 350(26), 2682–2688. 1372!
SCHWARZ, N. A., MCKINLEY-BARNARD, S. K., SPILLANE, M. B., ANDRE, T. L., GANN, J. J. & 1373!
WILLOUGHBY, D. S. (2016). Effect of resistance exercise intensity on the expression of PGC-1374!
1alpha isoforms and the anabolic and catabolic signaling mediators, IGF-1 and myostatin, in 1375!
human skeletal muscle. Applied Physiology, Nutrition and Metabolism 41(8), 856–863. 1376!
SEGERSTROM, S. C. (2007). Stress, Energy, and Immunity: An Ecological View. Current 1377!
Directions in Psychological Science 16(6), 326–330. 1378!
! 57!
SHINTAKU, J., PETERSON, J. M., TALBERT, E. E., GU, J. M., LADNER, K. J., WILLIAMS, D. R., 1379!
MOUSAVI, K., WANG, R., SARTORELLI, V. & GUTTRIDGE, D. C. (2016). MyoD Regulates 1380!
Skeletal Muscle Oxidative Metabolism Cooperatively with Alternative NF-kappaB. Cell 1381!
Reports 17(2), 514–526. 1382!
SHNAPER, S., DESBAILLETS, I., BROWN, D. A., MURAT, A., MIGLIAVACCA, E., SCHLUEP, M., 1383!
OSTERMANN, S., HAMOU, M. F., STUPP, R., BREIT, S. N., DE TRIBOLET, N. & HEGI, M. E. 1384!
(2009). Elevated levels of MIC-1/GDF15 in the cerebrospinal fluid of patients are associated 1385!
with glioblastoma and worse outcome. International Journal of Cancer 125(11), 2624–2630. 1386!
SLOT, I. G., VAN DEN BORST, B., HELLWIG, V. A., BARREIRO, E., SCHOLS, A. M. & GOSKER, H. 1387!
R. (2014). The muscle oxidative regulatory response to acute exercise is not impaired in less 1388!
advanced COPD despite a decreased oxidative phenotype. PLOS One 9(2), e90150. 1389!
SRIRAM, S., SUBRAMANIAN, S., SATHIAKUMAR, D., VENKATESH, R., SALERNO, M. S., 1390!
MCFARLANE, C. D., KAMBADUR, R. & SHARMA, M. (2011). Modulation of reactive oxygen 1391!
species in skeletal muscle by myostatin is mediated through NF-kappaB. Aging Cell 10(6), 1392!
931–948. 1393!
STEC, M. J., KELLY, N. A., MANY, G. M., WINDHAM, S. T., TUGGLE, S. C. & BAMMAN, M. M. 1394!
(2016). Ribosome biogenesis may augment resistance training-induced myofiber hypertrophy 1395!
and is required for myotube growth in vitro. American Journal of Physiology Endocrinology 1396!
and Metabolism 310(8), E652–E661. 1397!
STEIN, T. P. & BLANC, S. (2011). Does protein supplementation prevent muscle disuse 1398!
atrophy and loss of strength? Critical Reviews in Food Science and Nutrition 51(9), 828–834. 1399!
STEPHENS, N. A., GALLAGHER, I. J., ROOYACKERS, O., SKIPWORTH, R. J., TAN, B. H., 1400!
MARSTRAND, T., ROSS, J. A., GUTTRIDGE, D. C., LUNDELL, L., FEARON, K. C. & TIMMONS, J. 1401!
A. (2010). Using transcriptomics to identify and validate novel biomarkers of human skeletal 1402!
muscle cancer cachexia. Genome Medicine 2(1), 1. 1403!
! 58!
STITT, T. N., DRUJAN, D., CLARKE, B. A., PANARO, F., TIMOFEYVA, Y., KLINE, W. O., 1404!
GONZALEZ, M., YANCOPOULOS, G. D. & GLASS, D. J. (2004). The IGF-1/PI3K/Akt pathway 1405!
prevents expression of muscle atrophy-induced ubiquitin ligases by inhibiting FOXO 1406!
transcription factors. Molecular Cell 14(3), 395–403. 1407!
STRAUB, R. H. (2011). Concepts of evolutionary medicine and energy regulation contribute to 1408!
the etiology of systemic chronic inflammatory diseases. Brain Behaviour and Immunity 1409!
25(1), 1–5. 1410!
STRAUB, R. H., CUTOLO, M., BUTTGEREIT, F. & PONGRATZ, G. (2010). Energy regulation and 1411!
neuroendocrine-immune control in chronic inflammatory diseases. Journal of Internal 1412!
Medicine 267(6), 543–560. 1413!
SUBRAMANIAM, K., FALLON, K., RUUT, T., LANE, D., MCKAY, R., SHADBOLT, B., ANG, S., 1414!
COOK, M., PLATTEN, J., PAVLI, P. & TAUPIN, D. (2015). Infliximab reverses inflammatory 1415!
muscle wasting (sarcopenia) in Crohn's disease. Alimentary Pharmacology and Therapeutics 1416!
41(5), 419–428. 1417!
SUN, Z., WEI, Q., ZHANG, Y., HE, X., JI, W. & SU, B. (2011). MicroRNA profiling of rhesus 1418!
macaque embryonic stem cells. BMC Genomics 12(1), 276. 1419!
TATSUMI, R., ANDERSON, J. E., NEVORET, C. J., HALEVY, O. & ALLEN, R. E. (1998). HGF/SF 1420!
is present in normal adult skeletal muscle and is capable of activating satellite cells. 1421!
Developmental Biology 194(1), 114–128. 1422!
TERRUZZI, I., SENESI, P., MONTESANO, A., LA TORRE, A., ALBERTI, G., BENEDINI, S., CAUMO, 1423!
A., FERMO, I. & LUZI, L. (2011). Genetic polymorphisms of the enzymes involved in DNA 1424!
methylation and synthesis in elite athletes. Physiological Genomics 43(16), 965–973. 1425!
THOMSON, D. W., BRACKEN, C. P., SZUBERT, J. M. & GOODALL, G. J. (2013). On measuring 1426!
miRNAs after transient transfection of mimics or antisense inhibitors. PLOS One 8(1), 1427!
e55214. 1428!
! 59!
TIMMERMAN, K. L., LEE, J. L., DREYER, H. C., DHANANI, S., GLYNN, E. L., FRY, C. S., 1429!
DRUMMOND, M. J., SHEFFIELD-MOORE, M., RASMUSSEN, B. B. & VOLPI, E. (2010). Insulin 1430!
stimulates human skeletal muscle protein synthesis via an indirect mechanism involving 1431!
endothelial-dependent vasodilation and mammalian target of rapamycin complex 1 signaling. 1432!
Journal of Clinical Endocrinology and Metabolism 95(8), 3848–3857. 1433!
TOLKOVSKY, A. M. (2009). Mitophagy. Biochim Biophys Acta 1793(9), 1508–1515. 1434!
VAN ROOIJ, E., QUIAT, D., JOHNSON, B. A., SUTHERLAND, L. B., QI, X., RICHARDSON, J. A., 1435!
KELM, R. J., JR. & OLSON, E. N. (2009). A family of microRNAs encoded by myosin genes 1436!
governs myosin expression and muscle performance. Developmental Cell 17(5), 662–673. 1437!
VANDENBURGH, H. H., KARLISCH, P., SHANSKY, J. & FELDSTEIN, R. (1991). Insulin and IGF-I 1438!
induce pronounced hypertrophy of skeletal myofibers in tissue culture. American Journal of 1439!
Physiology Cell Physiol 260(3 Pt 1), C475–484. 1440!
VON WALDEN, F., CASAGRANDE, V., OSTLUND FARRANTS, A. K. & NADER, G. A. (2012). 1441!
Mechanical loading induces the expression of a Pol I regulon at the onset of skeletal muscle 1442!
hypertrophy. American Journal of Physiology Cell Physiol 302(10), C1523–1530. 1443!
WAGNER, P. D. (2006). Skeletal muscles in chronic obstructive pulmonary disease: 1444!
deconditioning, or myopathy? Respirology 11(6), 681–686. 1445!
WANG, F., WANG, J., YANG, X., CHEN, D. & WANG, L. (2016). MiR-424-5p participates in 1446!
esophageal squamous cell carcinoma invasion and metastasis via SMAD7 pathway mediated 1447!
EMT. Diagnostic Pathology 11(1), 88. 1448!
WANG, Y., HUANG, J. W., CASTELLA, M., HUNTSMAN, D. G. & TANIGUCHI, T. (2014). p53 is 1449!
positively regulated by miR-542-3p. Cancer Research 74(12), 3218–3227. 1450!
WATSON, E., SYLVIUS, N. B., VIANA, J. L. L., GREENING, N., BARRATT, J. & SMITH, A. (2016). 1451!
Differential MicroRNA Expression In Skeletal Muscle Of Human CKD Patients And Healthy 1452!
Controls. Nephrology Dialysis and Transplantation 31 (Supplement), i472–i478. 1453!
! 60!
WEST, D. W., BAEHR, L. M., MARCOTTE, G. R., CHASON, C. M., TOLENTO, L., GOMES, A. V., 1454!
BODINE, S. C. & BAAR, K. (2016). Acute resistance exercise activates rapamycin-sensitive 1455!
and -insensitive mechanisms that control translational activity and capacity in skeletal 1456!
muscle. Journal of Physiology 594(2), 453–468. 1457!
WILLIAMS, A. H., VALDEZ, G., MORESI, V., QI, X., MCANALLY, J., ELLIOTT, J. L., BASSEL-1458!
DUBY, R., SANES, J. R. & OLSON, E. N. (2009). MicroRNA-206 delays ALS progression and 1459!
promotes regeneration of neuromuscular synapses in mice. Science 326(5959), 1549–1554. 1460!
YANG, L., CHANG, C. C., SUN, Z., MADSEN, D., ZHU, H., PADKJAER, S. B., WU, X., HUANG, T., 1461!
HULTMAN, K., PAULSEN, S. J., WANG, J., BUGGE, A., FRANTZEN, J. B., NORGAARD, P., 1462!
JEPPESEN, J. F., et al. (2017). GFRAL is the receptor for GDF15 and is required for the anti-1463!
obesity effects of the ligand. Nature Medicine 23(10), 1158–1166. 1464!
YIN, H., PRICE, F. & RUDNICKI, M. A. (2013). Satellite cells and the muscle stem cell niche. 1465!
Physiological Reviews 93(1), 23–67. 1466!
YUE, F., BI, P., WANG, C., LI, J., LIU, X. & KUANG, S. (2016). Conditional Loss of Pten in 1467!
Myogenic Progenitors Leads to Postnatal Skeletal Muscle Hypertrophy but Age-Dependent 1468!
Exhaustion of Satellite Cells. Cell Reports 17(9), 2340–2353. 1469!
ZHANG, D., LI, X., CHEN, C., LI, Y., ZHAO, L., JING, Y., LIU, W., WANG, X., ZHANG, Y., XIA, 1470!
H., CHANG, Y., GAO, X., YAN, J. & YING, H. (2012). Attenuation of p38-mediated miR-1/133 1471!
expression facilitates myoblast proliferation during the early stage of muscle regeneration. 1472!
PLOS One 7(7), e41478. 1473!
ZHANG, Y., HUA, W., NIU, L. C., LI, S. M., WANG, Y. M., SHANG, L., ZHANG, C., LI, W. N., 1474!
WANG, R., CHEN, B. L., XIN, X. Y., ZHANG, Y. Q. & WANG, J. (2016a). Elevated growth 1475!
differentiation factor 15 expression predicts poor prognosis in epithelial ovarian cancer 1476!
patients. Tumour Biology 37(7), 9423–9431. 1477!
! 61!
ZHANG, Y., LI, C., LI, H., SONG, Y., ZHAO, Y., ZHAI, L., WANG, H., ZHONG, R., TANG, H. & 1478!
ZHU, D. (2016b). miR-378 Activates the Pyruvate-PEP Futile Cycle and Enhances Lipolysis 1479!
to Ameliorate Obesity in Mice. EBioMedicine 5, 93–104. 1480!
1481!
1482!
! 62!
Figure Legends 1483!
Fig. 1. Protein turnover changes in response to the requirements of anabolic and catabolic 1484!
stimuli. The continuous turnover of protein allows muscle synthesis in response to anabolic 1485!
stimuli and the provision of amino acids in response to inflammation or tissue damage. In 1486!
adults (where growth is no longer an issue) activity and increased nutrient intake stimulate 1487!
muscle protein synthesis and suppress muscle breakdown whereas inflammatory mediators 1488!
and the demand for repair promote muscle breakdown and inhibit muscle protein synthesis. 1489!
These effects are mediated by a range of anabolic and catabolic factors including those 1490!
shown. At any one time the net direction of this process (protein synthesis or amino acid 1491!
release) will be dependent on the sum of all anabolic and catabolic inputs at that time. 1492!
Insulin-like growth factor 1 (IGF-1), Interleukin 6 (IL6), interferon (IFN), growth and 1493!
differentiation factor 15 (GDF-15). 1494!
1495!
Fig. 2. Changes in either protein synthesis or protein degradation that could lead to 5% 1496!
muscle mass loss in 6 months. Using a fractional synthetic rate of 1% for the fasted state and 1497!
2.4% for the fed state, and assuming that only protein degradation or synthesis changes, it is 1498!
possible to calculate the required change in either process that would lead to a 5% loss of 1499!
muscle mass in 6 months. Using these assumptions, it can be seen that only very small 1500!
changes are required even if only one process changes. Consequently, as both synthesis and 1501!
degradation are altered the required change in either that could lead to muscle loss is minimal 1502!
and indeed is within the measured standard deviation in a population. 1503!
1504!
Fig. 3. Regulation of protein turnover by microRNAs (miRNAs). The miRNAs miR-542 and 1505!
424 promote transforming growth factor β (TGF-β) signalling (by targeting inhibitors of the 1506!
TGF-β signalling pathway) and inhibit the production of ribosomes thereby promoting the 1507!
! 63!
release of amino acids by increasing proteolytic activity and reducing protein synthetic 1508!
activity. miR-1, miR-181 and miR-422a inhibit activators of the TGF-β pathway, 1509!
consequently they reduce atrophic signalling and the release of amino acids from protein. 1510!
Activin A receptor like kinase 5 (Alk-5), small mothers against decapentaplegic 4 (SMAD4), 1511!
SMAD specific ubiquitin ligase (SMURF1), protein phosphatase 2A (PP2A), ribosomal RNA 1512!
(rRNA). 1513!
1514!
Fig. 4. Proposed effects of imprinted microRNAs (miRNAs). miRNAs from the cluster on 1515!
chromosome 19 (C19MC) reduce the rate of differentiation of myoblasts allowing longer for 1516!
myoblasts to proliferate and maintaining the satellite cell pool. Consequently, they are 1517!
associated with the maintenance of muscle in the context of disease. Conversely, miR-675 1518!
promotes the differentiation of myoblasts into skeletal muscle and inhibits myoblast 1519!
proliferation. Consequently miR-675 reduces muscle mass in the context of disease. 1520!
1521!
Fig. 5. Model for wasting in the presence of disease. In response to disease there is an 1522!
increase in the expression of microRNAs (miRNAs) that suppress cytoplasmic and 1523!
mitochondrial ribosomal synthesis and increase transforming growth factor β (TGF-β) 1524!
signalling leading to the release of amino acids. In individuals with a low susceptibility to 1525!
small mothers against decapentaplegic (SMAD) signalling or good regenerative capacity, 1526!
muscle wasting is minimal. In those who are susceptible to SMAD signalling and/or have a 1527!
poor regenerative capacity muscle wasting occurs. C19MC, miRNA cluster on chromosome 1528!
19. 1529!
1530! 1531!
