Download Transcriptional mechanisms regulating skeletal muscle

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Transcriptional mechanisms regulating
skeletal muscle differentiation,
growth and homeostasis
Thomas Braun* and Mathias Gautel‡
Abstract | Skeletal muscle is the dominant organ system in locomotion and energy
metabolism. Postnatal muscle grows and adapts largely by remodelling pre-existing fibres,
whereas embryonic muscle grows by the proliferation of myogenic cells. Recently, the
genetic hierarchies of the myogenic transcription factors that control vertebrate muscle
development — by myoblast proliferation, migration, fusion and functional adaptation into
fast-twitch and slow-twitch fibres — have become clearer. The transcriptional mechanisms
controlling postnatal hypertrophic growth, remodelling and functional differentiation
redeploy myogenic factors in concert with serum response factor (SRF), JUNB and forkhead
box protein O3A (FOXO3A). It has also emerged that there is extensive post-transcriptional
regulation by microRNAs in development and postnatal remodelling.
Myofibril
The structural unit of striated
muscle fibres, which is formed
from longitudinally joined
sarcomeres. Several myofibrils
form each fibre.
Myoblasts
Embryonic cells that will
become a muscle cell or
part of a muscle cell.
*Max-Planck-Institute for
Heart and Lung Research,
Department for Cardiac
Development and
Remodelling,
Benekestrasse 2,
61231 Bad Nauheim,
Germany.
‡
King’s College London,
Randall Division for Cell
and Molecular Biophysics
and Cardiovascular Division,
Muscle Signalling and
Development Section,
New Hunt’s House,
Guy’s Campus,
London SE1 1UL, UK.
e-mails: thomas.braun@
mpi-bn.mpg.de;
[email protected]
doi:10.1038/nrm3118
Movement is a defining feature of all animals, even if it
is sometimes restricted to specific developmental stages.
Larval migration, feeding, flight, reproduction and blood
circulation all rely on the directional and coordinated
movement that is afforded by muscles. The evolutionary
advantages of efficient locomotion led to several solutions
for the construction of the motile organs (the muscles)
in all animal phyla. In all muscle cells, myosin II motor
proteins and actin filaments generate force and move­
ment. In the striated muscles that are used for locomotion,
acto­myosin contraction is amplified in serial and parallel
arrangements of numerous contractile units, called sarco­
meres. These are made up of actin and myosin filaments
arranged in highly ordered, almost crystalline arrange­
ments, as well as hundreds of regulatory proteins such
as the troponin–tropomyosin complex, and scaffolding
and cytoskeletal crosslinking proteins such as α‑actinin,
myomesi­n and the kinase titin1,2 (FIG. 1).
In vertebrates, striated muscle cells are found in two
tissues: skeletal and heart muscle (FIG. 1). Although they
both have highly ordered myofibril structures, they have
distinct embryonic origins and are tailored for particular
purposes by different genetic programmes. Furthermore,
in vertebrates specialized skeletal muscles with different
contractile (slow-twitch or fast-twitch) and metabolic
properties coexist3,4.
Such distinctions are determined by the activity of spe­
cific transcription factors, the myogenic regulatory factors
(MRFs), which act together with pleiotropic transcription
factors and epigenetic regulatory mechanisms to control
muscle development and postnatal remodelling. During
embryonic development, these genetic programmes
determine the primary differentiation and also the
future metabolic and contractile properties of tissues in
specific anatomical compartments, thus controlling the
commit­ment of future slow-twitch or fast-twitch mus­
cles. Postnatally, the degree of muscle use and the specifi­c
innervation patterns determine the composition and
turnover rates of muscle contractile proteins, as well as
of the supporting metabolic enzymes, ion channels and
signal transduction proteins. This remodelling of differ­
entiated muscle determines the contractile properties and
the preferred sources of energy. It is governed by signal­
ling cascades (modulated by growth factors and cytokines,
steroid hormones and mechanical activity) that control
the transcriptional activit­y of muscle-specific and pleio­
tropic transcription factors. These genetic programmes
also regulate changes in muscle mass. During embryonic
development, muscle mass increases predominantly by
proliferative growth of myoblasts. Postnatally, the contri­
bution of cell proliferation decreases, and hypertrophic
growth and remodelling of pre-existing muscle fibres
dominates; the resident stem cells (the satellite cells) are
then mostly engaged in damage repair 5–7.
Recent research has shed important light on the
basic mechanisms that lead to the commitment of pre­
cursor cells to the muscle lineage (myogenesis). As a
result, we now have a better understanding of how the
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Figure 1 | Striated muscle structure. The contractile machinery of skeletal muscle
syncytial myotubes (left) and single cardiomyocytes
(right) is formed from long arrays
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of sarcomere units, which are joined into myofibrils. The sarcomere (bottom) is
constructed from interdigitating, antiparallel filaments of actin and myosin, the elastic
titin filaments and the crosslinker proteins for actin — α‑actinin, myosin and myomesin.
Sarcomeres contain many other accessory components, including proteins involved in
transcriptional regulation and turnover control. The transcription factor CLOCK, the
transcriptional cofactors muscle LIM protein (MLP), muscle ankyrin-repeat proteins
(MARPs) and LIM domain-binding protein 3 (LDB3) are found at the Z‑disk and/or
the I‑band. Multifunctional components of the protein turnover machinery include
sequestosome 1 (SQSTM1), NBR1 and the muscle-upregulated RING finger proteins
(MURFs). MYOZs, myozenins.
functional diversity of different striated muscles evolves
on the develop­mental level, and how physiological stimuli
are translated into changes in muscle gene expression
patterns, metabolic flow and protein turnover. This
recent research has also revealed how postnatal muscle
Paraxial mesoderm
The mesodermal areas that
form directly lateral to the
neural tube.
Rostrocaudal axis
A description of anatomical
location in animals. Rostral
(from the latin rostrum
meaning beak) refers to the
anterior (‘nose-end’) of
the animal and caudal
(from the latin caudum
meaning tail) refers to the
posterior (‘tail or feet end’).
Somites
Mesodermal structures found
on either side of the neural
tube in vertebrate embryos
that eventually give rise to
muscle, skin and vertebrae.
Delamination
A process in embryology in
which cells from a single layer
separate to form two different
layers, or laminae.
remodelling uses myogenic transcription factors that are
also active during early development. In this Review, we
outline how tight developmental regulation, mechanical
functions and the homeostasis of the organism are inter­
linked during development and postnatal muscle adapta­
tion. We focus on recent insights into the transcriptional
and post-transcriptiona­l control that commits myogenic
cells to proliferation or differentiation, determines their
lineage fate as slow-twitch or fast-twitch striated mus­
cles and regulates the postnatal adaptive remodelling
of committed muscle cells. Where appropriate, we con­
sider relevant similarities between skeletal and cardiac
striated muscles.
Transcriptional control of myogenesis
Skeletal muscle cells of higher vertebrates arise during
midgestation (in mice between embryonic day 9 (E9)
and E12) from three different locations within the middle
layer of cells in the primitive embryo: the segmented som­
itic paraxial mesoderm, the unsegmented cranial paraxial
mesoderm and the prechordal mesoderm; these repre­
sent different parts of the mesoderm along the rostrocauda­l
axis (reviewed in REF. 8). The skeletal muscles of the trunk
and limbs are derived from cells of the segmented paraxial
mesoderm (known as somites), which form on either side
of the neural tube in vertebrate embryos. Somites further
differentiate into a dorsal part, called the dermomyo­
tome (which retains an epithelial structure) and a ventra­l
part, called the sclerotome (which breaks up into the
mesenchyme that contributes to the axial skeleto­n of the
embryo). The first muscle cells are formed in the myotome,
which is located directly underneath the dermomyotome
and is formed by cells that separate by delamination from
the dermomyotome (reviewed in REF. 9) (BOX 1).
In all the anatomical sites where skeletal muscle forms,
determination and terminal differentiation of muscle
cells are governed by a network of four MRFs: myogenic
Box 1 | Skeletal muscle cell specification and differentiation during embryogenesis
Formation of trunk muscles
The morphogenetic events that are involved in the formation of the myotome have been extensively studied. Cell tracing
methods have shown that a first wave of myogenic factor 5 (MYF5)-expressing muscle progenitor cells, which
progressively withdraw from the cell cycle, delaminate and migrate towards the rostral somite, making up the medial
region of epithelial somites. Myofibres differentiate in rostrocaudal (head to tail) and mediolateral (centre outwards)
directions (reviewed in REF. 133). This initial wave of myogenesis is followed by a second wave of myoblasts. These cells
come from all four lips of the dermomyotome, which is the epithelial cell layer that comprises all of the mesodermal
somites (but not the sclerotome) and that gives rise to the axial skeleton. The dorso–medial lip is initially the sole
contributor to myogenesis, followed by the posterior lip, until eventually the anterior and lateral borders begin to
contribute to myogenesis134. It should be emphasized that the myotome that is formed by the epithelial borders of
the dermomyotome is composed of postmitotic cells, which do not contribute to further muscle growth. Only the
continuous addition of proliferating muscle progenitor cells allows the primary muscle compartment to expand.
Formation of head muscles
The formation of head muscles differs significantly from the formation of their counterparts in the trunk and limbs.
The head musculature originates from the cranial paraxial mesoderm (CPM), which is located anterior to the somites;
additional input to head muscles comes from the lateral splanchnic mesoderm (SpM). Cells from the CPM migrate to
the proximal part of the central region of the branchial arches, and SpM-derived cells contribute to the distal region.
Later in development, CPM-derived myogenic cells of the first branchial arch will form the masseter muscle, whereas
SpM- derived myogenic cells generate the lower jaw muscles (reviewed in REF. 135). Formation of head muscles is also
strongly influenced by cranial neural crest cells136. Deletion of neural crest cells leads to severe reduction of jaw muscles
after the onset of muscle specification and differentiation135,137, and this is probably due to interactions between
muscles and neural crest-derived tendons138.
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factor 5 (MYF5), muscle-specific regulatory factor 4
(MRF4; also known as MYF6), myoblast determination
protein (MYOD) and myogenin. MRFs are transcription
factors that activate many downstream genes to initiate
muscle cell differentiation. MYOD and MYF5 are musclespecific transcription factors and constitute a crossregulator­y transcriptional network that is at the core of
muscle cell determination and differentiation (FIG. 2);
disruption of this network completely abrogates skel­
etal muscle formation. MYF5 and MYOD are generally
thought to act as determination genes, whereas myogenin
is essential for the terminal differentiation of committed
myoblasts (reviewed in REF. 10). MRF4 seems to have a
dual role: it is thought to be a differentiation gene acting
in postmitotic maturating cells, but it is also expressed by
undifferentiated proliferating cells in which it might act as
a determination gene11. The upstream signals (transcrip­
tion factors and extracellular signals (BOX 2)) that activate
MRFs differ significantly at various anatomical locations
(FIG. 2), although some molecular cues are shared.
Neural crest
A group of embryonic cells that
separate from the embryonic
neural plate and migrate,
giving rise to the spinal and
autonomic ganglia, peripheral
glia, chromaffin cells,
melanocytes and some
haematopoietic cells.
Hypaxial muscles
Muscles that usually lie ventral
to the vertebrae and are
innervated by the ventral
ramus of the spinal nerves.
Epaxial muscles
Muscles that usually lie dorsal
to the vertebrae (in fish and
amphibiae they lie dorsal to
the septum). They are
innervated by the dorsal
ramus of the spinal nerves.
Regulation of trunk skeletal muscle formation. Numerous
transcription factors, such as paired box, homeobox and
T-box proteins, have been identified as binding upstream
of MRF genes. None of these factors is exclusively
expressed in the muscle progenitor cells that give rise
to differentiated muscl­e cells, which suggests that these
trans­cription factor­s prepare the stage for additional actors
(such as MRFs) to initiate myogenesis. Alternatively, they
may act together with other transcription factors to allow
the activation of MRFs. In both cases, instructive or per­
missive inductional cues are necessary to achieve stable
self-sustained activation of MRFs in muscle progenitor
cells and muscle cell formation.
The differentiation of hypaxial muscles and epaxial
muscles seems to differ with regard to the transcriptional
network that activates MRFs. Indeed, the target genes of
paired box protein 3 (PAX3), which is upstream of the
MRFs, differ in hypaxial and epaxial muscles.
In mouse epaxial muscles, PAX3 seems to activate
MYF5 by controlling the expression of Dmrt2, which in
turn activates the epaxial enhancer of Myf5 in somites12.
Furthermore, analysis of mice depleted of PAX3 and
MYF5 showed almost complete loss of trunk muscles
and a loss of MyoD expression; this indicates that MyoD
expression depends on either PAX3 or MYF5 (REF. 13).
At present, it is not known precisely how PAX3 activates
MyoD expression, although experiments in presomitic
explant cultures suggest that WNT signalling regulates
MyoD expression in a PAX3‑dependent manner 14. Taken
together, current evidence suggests that PAX3 medi­
ates the activation of MYOD and MYF5 in a tight inter­
play with muscle-inducing signals. Early experiments
showed that ectopic expression of PAX3 is sufficient to
induce the expression of MYOD, MYF5 and myogenin
in the absence of inducing tissues (that is, neural tube
and paraxial surface epithelium) in both the paraxial and
latera­l plate mesoderm of chicken embryo, and in the
neural tube15,16; however, directed expression of PAX3
in vivo did not induce a stable myogenic fate. Similarly,
expression of LBX1, a PAX3 target gene, resulted in the
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Figure 2 | Different ways to activate the genetic
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programme of muscle
differentiation. All muscle cells
express a core set of myogenic factors (for example,
myogenic factor 5 (MYF5), muscle-specific regulatory
factor 4 (MRF4), myoblast determination protein (MYOD)
and myogenin), which are required for myogenic
differentiation. Other transcription factors reflect
lineage-specific differences and are necessary for the
activation of myogenic factors and/or proliferation and
survival of muscle progenitor cells. Muscle groups from the
head, including extraocular, tongue and laryngeal muscles,
and branchial arches, are derived from occipital somites,
cranial paraxial mesoderm, splanchnic mesoderm and
prechordal mesoderm. In these cells, pituitary homeobox 2
(PITX2) predominantly controls the myogenic hierarchy,
leading to the activation of MYF5 and MYOD and
eventually terminal differentiation induced by myogenin.
By contrast, muscles from the limbs and the trunk are all
derived from trunk somites. In limb muscles, sine oculis
homeobox homologue (SIX) and eyes absent (EYA) proteins
regulate paired box protein 3 (PAX3), which in turn controls
the proliferative myogenic cell pool, the differentiation
of which is induced by a cascade involving MYF5, MRF4,
MYOD and myogenin. In trunk muscles, MYF5 or MRF4 can
show parallel activation of MYOD and myogenin, whereas
PAX3 acts upstream of MYOD. Solid lines represent direct
control and dashed lines represent indirect control.
TBX1, T-box transcription factor. Figure modified, with
permission, from REF. 150 © (2009) Elsevier.
activation of myogenesis in vitro and transient induc­
tion of myogenic determination genes in vivo in chicken
embryos13,16. Interestingly, mutants of the Pax3 paralogue
Pax7 do not show overt muscle defects during mouse
development17, and the combined loss of PAX3 and PAX7
yields a phenotype that is similar to PAX3 mutants, with
the initial formation of skeletal muscle occurring in the
myotome until E10.5. This indicates that the initial form­
ation of skeletal muscle cells in the myotome occurring
up to E10.5 is not directly under the control of PAX3 and
PAX7. As compound PAX3 and PAX7 mutants show a
severe disruption of muscle development at later stages,
with very few differentiated muscle cells18, it is possible
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Box 2 | Extracellular signals directing muscle development
The activation of the network of transcription factors that controls skeletal muscle
development depends on paracrine factors that are released by adjacent tissues, such as
the neural tube, notochord, surface ectoderm and lateral mesoderm. Several secreted
factors have been identified that determine the spatial and temporal onset of
myogenesis. Surprisingly, no consensus has been reached as to whether these molecules
instruct naive cells (instructive induction), amplify a pool of committed progenitors
and/‌or enable a default differentiation pathway (permissive induction) or primarily
prevent programmed cell death of muscle progenitor cells (reviewed in REF. 139).
From numerous studies, it is clear that sonic hedgehog (SHH) and WNT signalling have
pivotal roles in the induction of myogenesis. Moreover, other signalling molecules, such
as Noggin and bone morphogenetic proteins (BMPs) — which inactivate and activate
receptors of the transforming growth factor-β (TGFβ) superfamily, respectively — play
an important part in orchestrating the activation of myogenesis140.
A study in mice showed that the canonical β‑catenin-mediated WNT signalling
pathway acts co-operatively with SHH, its receptor Patched and its downstream
target, GLI (a zinc-finger transcription factor), to regulate the expression of myogenic
factor 5 (Myf5). According to this model, the SHH–Patched–GLI pathway is not
sufficient to induce myogenesis if the local concentration of β‑catenin does not
suffice to support full transcriptional activity of lymphoid enhancer-bracking factor 1
LEF1 or other transcription factors, which facilitate the localization of β‑catenin to
specific cis-regulatory elements. The transcription of the first myogenic transcription
factor, MYF5, is activated in the epaxial domain only when both SHH and canonical
WNT signalling pathways are activated at the onset of somitogenesis141. Despite
compelling evidence for the decisive role of canonical β‑catenin-mediated WNT
signalling in the induction of myogenesis, other authors reported that adenylyl
cyclase signalling through protein kinase A (PKA) and its target transcription factor,
cAMP-responsive element-binding protein (CREB), are required for WNT-directed
myogenic gene expression142. It is possible, however, that the WNT signalling pathway
that involves PKA and CREB acts in parallel to canonical β‑catenin-mediated WNT
signalling and becomes limiting only under specific conditions.
that PAX3 and/or PAX7 are responsible for the enlarge­
ment of muscle precursor cell populations. This would
increase the bias towards myogenic differentiation and
thus enable myogenic cells to respond to environmental
cues16. This view is further supported by the observation
that PAX3 directly regulates components of the fibroblast
growth factor (FGF) signalling pathway, which have a role
in the expansion of the myogenic progenitor cell pool19.
In hypaxial muscle cells, PAX3 seems to directly activate
MYF5 but not MYOD. Similarly, a study in muscle stem
cell-derived myoblasts showed that expression of MYF5
is regulated by PAX7 (REF. 20). However, despite having a
role in the activation of MYF5 and the morpho­genesis
of the hypaxial dermomyotome, PAX3 does not seem to
be essential for the development of hypaxial muscles, as
the lateral halves of somites from E9.25 PAX3‑mutant
embryos transplanted into the limb bud of chicken host
embryos differentiated normally 21. Furthermore, recent
studies using a PAX3–engrailed fusion protein, which acts
as a transcriptional repressor, suggested that PAX3 directly
regulates Myf5 in the hypaxial somite and its limb mus­
cle derivatives. In these mice, the expression of Myf5, but
not of MyoD, was compromised in hypaxial muscles. The
authors postulated that the regulation of Myf5 by PAX3
in hypaxial muscles was achieved by a PAX3 consensus
site located in a 145 bp regulatory element –57.5 kb from
Myf5. Mutation of the PAX3‑binding site in the context
of the 145 bp element abolished all expression in trans­
genic mouse embryos22. Similarly, sequences have been
described that contain a predicted homeobox adjacent to
a putative paired domain-binding site within the –58 to
–56 kb distal Myf5 enhancer, which directs Myf5 expres­
sion in myogenic progenitor cells in limbs. Both PAX3
and mesenchyme homeobox gene 2 (MEOX2) transcrip­
tion factors could bind these consensus sites in vitro, so
they are potential regulators23. However, there is no valid
genetic evidence showing that MEOX2 has a signifi­
cant effect on Myf5 expression in the limb, despite some
early reports claiming a reduced expression of MYF5 in
MEOX2‑mutant limbs. It seems more likely that sine oculis
homeobox homologue 1 (SIX1) and SIX4 (see below)
occupy the homeobox-binding site in the 145 bp element.
In fact, it was recently shown that SIX1 and SIX4 regulate
the transcription of MYF5 in the limb together with PAX3,
by binding to the 145 bp element –57.5 kb from Myf5. An
additional regulatory element seems to contribute to the
expression of Myf5 in the hindlimbs, and this element
might be responsible for some of the remaining expressio­n
of Myf5 in the hindlimb buds of SIX1 mutants24.
SIX proteins are also involved in the regulation of
myogenesis in the limb. SIX proteins act, at least in part,
upstream of myogenic regulatory factors and PAX pro­
teins (FIG. 2). The human and mouse genome contains
6 SIX genes (SIX1–SIX6), 3 of which (SIX1, SIX4 and
SIX5) are expressed from E8 in overlapping expression
patterns in somites, limb buds, dorsal root ganglia and
branchial arches. SIX proteins form complexes with their
transcriptional co-activators, eyes absent (EYA) proteins,
to stimulate transcription synergistically 25. SIX1 mutants
die at birth and show selective muscle hypoplasia in the
diaphragm, forelimb, distal ventral hindlimb and abdo­
men26. The muscle phenotype is aggravated in compound
mutants of SIX1 and SIX4, which do not have myogenic
progenitor cells in their limb buds, resulting in legs with
no muscles27. Furthermore, neither double SIX1 and
SIX4 mutants nor double EYA1 and EYA mutants express
Pax3 in the hypaxial dermomyotome, indicating that Six
and Eya lie upstream of Pax3 in the genetic hierarchy
of hypaxial myogenesis. SIX1 and SIX4 double mutants
also show a reduced expression of MyoD, myogenin and
other myotomal markers, although the early activation
of MYF5 in the epaxial somite is un­affected27. However,
later in development the expression of SIX genes seem
to depend on MRFs, indicating a dual role for SIX pro­
teins both upstream and downstream of MRFs during
myogenesis28. The role of SIX proteins in the differentia­
tion of muscle fibres intro slow-twitch or fast-twitch is
discussed below.
MRFs are also assisted by many other factors such
as PBX and MEIS proteins29, which function as hetero­
dimers and act as cofactors with basic helix–loop–helix
(bHLH) proteins such as MRFs and various homeobox
transcription factors. PBX and MEIS proteins participate
in the feedforward mechanisms that enable MRFs to acti­
vate ‘early’ genes of the muscle differentiation programme
immediately, whereas genes activated later on in the dif­
ferentiation programme need both MRFs and the addi­
tional participation of one of the earlier MRF targets29
(reviewed in REF.10). Myocyte enhancer factor 2 (MEF2)
proteins are also important members of this regulatory
circuit. MEF2 proteins interact directly with MYOD
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in vitro and synergistically activate reporter genes that
contain E-boxes and MEF2‑binding sites30. E-boxes and
MEF2‑binding sites are often located close to one another
within the promoters and enhancers of muscle-specific
genes, suggesting a model in which MRFs and MEF2 bind
DNA and activate transcription in a cooperative manner.
Three of the four MEF2 proteins (MEF2A, MEF2C and
MEF2D) are expressed in skeletal muscle, and expression
and splicing of these isoforms is altered in response to
MYOD, which further supports a role for MYOD in the
feedforward mechanism that is driven by MRFs29.
The transcriptional regulation of head muscles. Recent
evidence suggests that head muscle development follows
a distinct programme that does not require the action of
PAX3 and PAX7. Initial specification of mouse massete­r
muscles depends on the bHLH genes MyoR and capsu­
lin31. Additional players are pituitary homeobox 2 (Pitx2)
and T-box transcription factor 1 (Tbx1), which collabo­
rate with the core myogenic programme to gener­ate head
muscles. Both Pitx2 and Tbx1 are expressed widely in
the developing mouse embryo and play an important
part in the formation of head muscles (FIG. 2). Indeed,
PITX2 mutants do not properly develop the muscles that
are derived from the first branchial arch32 and they lack
extraocular muscles33. TBX1 mutants also suffer from
impaired myogenesis in the first branchial arch and lack
the muscles that normally originate from other arches
owing to severe malformations of these structures34,35.
It has recently been shown that TBX1 and MYF5 act
syner­gistically to govern myogenesis in pharyngeal muscle
progenitor­s, thereby acting as a complementary pathway
to that involving PAX3 and MYF5 in the body36. It is poss­
ible that PITX2 and TBX1 regulate the quiescence and
self-renewal of muscle progenitors in the head similarl­y
to PAX3 and PAX7 in trunk skeletal muscles.
Masseter muscles
A specific subset of
branchiomeric muscles that
are derived from the first
branchial arch and are
involved in mastication.
Pharyngeal muscle
A subgroup of head muscles
acting on the pharynx to
control swallowing.
Post-transcriptional control by microRNAs
The human genome contains thousands of non-coding
RNAs, the best-studied class of which are microRNAs
(miRNAs) (reviewed in REF. 37), which regulate gene
expression at the transcriptional and post-transcriptiona­l
levels. miRNAs suppress gene expression through
their complementarity to the sequence of one or more
mRNAs, usually at a site in the 3′ untranslated region.
The formation of an miRNA–target complex results
either in inhibition of protein translation or in degrada­
tion of the mRNA transcript through a process similar to
RNA interference38. There is no doubt that the formation,
maintenance, and physiological and pathophysiological
responses of skeletal muscles, with all their complex regu­
latory circuits, are subject to regulation by non-coding
RNAs. In fact, the increase of complexity provided by
the extent of genomic non-coding sequences provides a
satisfying explanation for the intricate layers of regulation
found in skeletal muscle cells.
Many miRNAs are expressed in skeletal and cardiac
muscle. Some of them are found specifically in skeletal
and/or cardiac muscle, or at least are highly enriched
in these tissues, suggesting specific roles in myogen­
esis39. The expression of the muscle-specific miRNAs
miR‑1, miR‑133, miR‑206 and miR‑208 seems to be
under the control of a core muscle transcriptional net­
work, which involves the pleiotropic serum response
factor (SRF), MYOD, and the bHLH transcription factor
Twist in co­operation with MEF2 (REFS 40–43). Chromatin
immuno­precipitation followed by microarray (ChIP–
chip) analysis indicated that MYOD and myogenin bind
sequences upstream of miR‑1 and miR‑133 (REF. 41).
Furthermore, MEF2 has a crucial role in the control of an
intragenic enhancer located in the miR‑1–2 locus (which
contains miR‑1 and miR‑2)44. Recently, analysis of mice
deficient in MYF5 and MYOD revealed a surprisingly
specific requirement of MYF5 for miR‑1 and miR‑206
expression. At early developmental stages, the expression
of both miR‑1 and miR‑206 was almost entirely absent in
the somites of MYF5‑mutant embryos, whereas mouse
embryos lacking MYOD showed an apparently normal
expression of both miRNAs45. Whereas miR‑133 and
miR‑206 are expressed as independent transcriptional
units, miR‑208 is encoded by an intron of its host gene,
α‑myosin heavy chain (αMHC)46. Both miR‑208 and
αMHC are heart specific and concurrently expressed
during development, suggesting that their expression is
controlled by a common regulatory element46.
Some experiments suggest that miRNAs act as modu­
lators of myogenic differentiation, in particular because
some miRNAs, such as miR‑1 and miR‑133 (FIG. 3a), are
absent from undifferentiated myoblasts and are strongly
upregulated upon differentiation47. An increase of miR‑1
expression in tissue culture accelerates myoblast dif­
ferentiation by downregulating histone deacetylase 4
(HDAC4), a repressor of muscle differentiation, and
knockdown of miR‑1 impedes myogenic differentia­
tion40. Although a similar role for miR‑1 in facilitating
differentiation has been shown in the developing mouse
heart, where a tissue-specific overexpression of miR‑1
induces premature differentiation of cardiomyocytes43,
conclusive evidence for an indispensable role of miR‑1 in
the regulation of myogenic differentiation is still missing.
Recent studies, however, suggest that re-expression of
miR‑206 in human rhabdomyosarcoma cells promotes
myogenic differentiation and blocks tumour growth
in xenografted mice by switching the global mRNA
expression profile to one that resembles mature mus­
cle48. This finding supports an important modulatory
role for miR‑206 in the control of myogenesis (FIG. 3b),
although miR‑206‑knockout mice show no major arrest
of myogeni­c differentiation49.
The modulatory role of miRNAs on muscle dif­
ferentiation is underscored by several observations on
the role of miR‑1, miR‑27, miR‑206 and miR‑486 in the
regulation of PAX3 and PAX7 (FIG. 3c). miR-1, miR‑27
and miR‑206 were found to inhibit PAX3 and to thereby
release the inhibitory effect of PAX3 on terminal muscle
differentiation50,51; an analogous role was described for
the regu­lation of PAX7, which is repressed by miR–1 and
miR‑206 (REFS 52,53). Therefore, the feedback loop ampli­
fying term­inal MYOD-induced muscle differentiation
involves the transcriptional switch towards late factors
such as MEF2C, but also the epigenetic downregulation
of muscle stem cell factors such as PAX3 and PAX7.
NATURE REVIEWS | MOLECULAR CELL BIOLOGY
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© 2011 Macmillan Publishers Limited. All rights reserved
REVIEWS
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Figure 3 | Regulation of major myogenic pathways
by microRNAs. a | Regulation of
0CVWTG4GXKGYU^/QNGEWNCT%GNN$KQNQI[
myogenic differentiation by miR‑1 and miR‑133. miR‑1 and miR‑133 are downstream
of the myogenic transcription factors myogenic factor 5 (MYF5) and myocyte enhancer
factor 2C (MEF2C) and control the expression of differentiation genes: first, through
the pleiotropic transcription factor serum response factor (SRF); second, by inhibiting the
assembly of splicing-regulatory protein complexes by neural polypyrimidine tract-binding
protein 2 (nPTB2), which controls the splicing of many mRNAs in muscles; and third, by
inhibiting histone deacetylase 4 (HDAC4), which blocks myogenic differentiation.
b | miR‑206 is downstream of the myogenic master transcription factor myoblast
determination protein (MYOD) and feeds back on MyoD expression by inhibiting
follistatin and bone morphogenetic proteins (BMPs), ultimately determining the
expression of differentiation markers such as utrophin. c | Regulation of proliferation and
differentiation by the myoblast transcription factors paired box protein 3 (PAX3) and PAX7
is controlled upstream by miR‑1, miR‑27, miR‑206 and miR‑486, which repress PAX3 and
PAX7 and thus promote the terminal differentiation of myoblasts downstream of MYOD.
Antagomirs
Synthetic or genetically
engineered oligonucleotides
used to silence endogenous
microRNAs.
Soleus muscle
A muscle in the calf of the leg
that flexes the ankle; in rodents
it is predominantly composed
of slow-twitch fibres.
The regulation of myogenesis by miR‑1 and miR‑133
is unclear, as the human and mouse genomes each con­
tain two bicistronic loci, which both express miR‑1 and
miR‑133. It has been reported that the selective disruption
of the miR‑1–2 locus in mice results in a severe cardiac
phenotype, including 50% embryonic lethality with fre­
quent ventricular septum defects54, which is surprising
given that the sequences of miR‑1 and miR‑133 are identi­
cal and that both loci are expressed in skeletal and cardiac
muscle. Combined knockouts of the miR‑1‑2–miR‑133a1
and miR‑1‑1–miR‑133a2 clusters are still pending and will
provide more definitive answers. Nevertheless, it seems
likely that miR‑1 and miR‑133 are required to delineate
the phenotype of differentiated muscl­e cells, which is
consistent with the conclusions drawn from the miR‑1
overexpression experiment55.
Currently views on miRNA functions remain dispa­
rate and are complicated by some underlying technical
problems. For example, massive overexpression might
not necessarily reflect a physiological function of an indi­
vidual miRNA. Instead, the non-physiological presence
of large amounts of an exogenous miRNA might cause
numerous off-target effects, as also demonstrated for small
interfering RNAs, and could lead to wrong conclusions56.
Similarly, knockdown approaches using antagomir­s,
although highly appealing because of their relative ease
of use and wide applicability, might hold some pitfalls that
need to be explored57.
Postnatal control of muscle phenotype
Skeletal muscle has a remarkable ability to rapidly adjust
to changes in physiological requirements, by changes
in excitability, contractile characteristics, metabolism,
and fibre size and mass (see below). These sweeping
changes involve the concerted control of transcription
regulation, protein synthesis, protein degradation and
metabolic flow, the details of which have only recently
emerged. During postnatal adaptation of muscle, or
muscle remodelling, aspects of early developmental
programmes can be reactivated and can cooperate with
specific factors to determine fibre-type characteristics58;
MEF2, nuclear factor of activated T cells (NFAT), myo­
genin, JUNB and SRF have prominent roles in these
processes59–64. Post-translational modifiers of histones
(histone acetylases and HDACs)64,65 and of transcription
factors (protein kinases and phosphatases, and ubiquitin
and SUMO transferases) cooperate in the complex inter­
actions between chromatin structure and transcriptional
machinery. Many of these responses are modulated
by muscle activity, either by sensing neuronal activity
though intra­cellular Ca2+ levels or directly by mechani­
cal strain. In addition, various hormonal and cytokine
signalling mechanisms feed into the same pathways.
The adult skeletal musculature contains highly dif­
ferentiated fibre types: one slow-twitch, oxidative fibre
type (type I) and three fast-twitch, glycolytic fibre types
(type 2)4. Fast-twitch fibre development follows defined
patterns, involving control by SIX1 and SIX4 (see
above)66, which also play an important part postnatally
in the adaptation to the fast muscle phenotype25. By con­
trast, slow-twitch fibres are determined by the transcrip­
tional repressor BLIMP1 (REF. 67), which suppresses the
transcription factor SOX6 (REFS 68,69). The distinct origin
of fast-twitch and slow-twitch fibres is also highlighted
by the observation that the developmental lineages
and the regeneration of slow- and fast-twitch muscles
stems from the recruitment of intrinsically different
myogenic precursors70 or satellite cells71. Postnatally,
SIX1 co­operates with EYA1 in fibre type differentiation.
Indeed, forced co-expression of SIX1 and EYA1 in the
slow-twitch fibres of soleus muscle induced a transition
to a fast-twitch fibre type, with the replacement of the
slower myosin heavy chain isoforms I and IIA by the
faster IIB and/or IIX, accompanied by the activation
of other fast-twitch fibre-specific genes and a switch to
glycolyti­c metabolism.
For physiological, activity-dependent adaptation of
existing fibres, the type of neuronal activity acting on
a fibre is probably the most important factor determin­
ing fibre type. The firing pattern of neurons innervating
fast-twitch and slow-twitch muscle fibres has markedly
different frequencies and temporal patterns, leading
to different membrane potentials and ultimately to net
differences in intracellular Ca2+ levels. These changes
can be sensed by the Ca2+-activated Ser phosphatase
calcineurin (also known as PP2B), which dephosphoryl­
ates and thereby activates NFAT. This can then act as a
calcineurin-dependent sensor and can translate nerve
activity in skeletal muscle into altered fibre-type-specific
gene expression programmes (FIG. 4). The nuclear version
354 | JUNE 2011 | VOLUME 12
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REVIEWS
Tibialis anterior muscle
A muscle in the front muscle
compartment of the lower leg
that helps to extend the ankle;
in rodents it is predominantly
composed of fast-twitch fibres.
Microgravity
Gravity below the 1Gal
experienced on Earth; for
example, during space flight.
Ubiquitin–proteasome
system
A system of selective,
ATP-dependent protein
degradation, in which target
proteins that have been
conjugated by ubiquitin
are degraded by the 26S
proteasome. The ubiquitin
conjugation step requires
the activity of highly specific
ubiquitin ligases.
Autophagy
A catabolic process involving
the engulfment of (usually
damaged) organelles and
long-lived proteins or
protein aggregates by
double-membrane vesicles
(autophagosomes) that fuse
with lysosomes to form
autolysosomes, in which their
contents are degraded by
acidic lysosomal hydrolases.
of NFAT (NFATC1) cooperates with numerous musclespecific cofactors such as MYF5 (REF. 72) to modulate
muscle-specific gene expression 59, thus controlling
the expression of fast and slow myosin isoforms 73.
The activit­y-dependent nuclear import and export of
NFATC1 is a rapid event, occurring within minutes
of the appropriate slow-type stimulation pattern74. Its
activity is counteracted by the calcineurin inhibitor
CAIN (also known as CABIN1), which promotes nuclear
export of NFATC1 both in soleus and stimulated tibialis
anterior muscle.
Further regulation of calcineurin activity is mediated
by a family of sarcomere-associated proteins known as
myozenins (MOYZs; also known as FATZs and cal­
sarcins), which are encoded by the three MYOZ genes.
MYOZ2 is expressed in adult cardiac and slow-twitch
skeletal muscle, whereas MYOZ1 is restricted to fasttwitch skeletal muscle. MYOZs interact with calcineurin
and the PDZ domain and LIM domain-containing
protein LIM domain-binding protein 3 (LDB3; also
known as ZASP, cypher and oracle) (reviewed in REF. 2).
MYOZ2‑knockout mice show increased calcineurin
signalling in striated muscles, suggesting that MYOZ2
acts as a brake on calcineurin activity 75. As a result of
the abnormal calcineurin activation, MYOZ2‑knockout
mice have an excess of slow-twitch skeletal muscle fibres,
which is consistent with the established roles of calci­
neurin and NFAT in activity-dependent regulation of
fibre types59. Similarly, MYOZ2‑knockout mice activate
a cardiac hypertrophic gene programme in the absence
of hypertrophy and show enhanced cardiac growth in
response to pressure overload75.
Owing to the probable function of the sarcomere
Z‑disk (FIG. 1) as a sensor for muscle mechanical strain
(reviewed in REF. 76), calcineurin activity might also be
mechanically modulated by the MYOZs in the Z‑disk.
The interplay of transcriptional regulation by calci­
neurin, SIX1 and EYA, and MRFs, which seems possible
from these observations, and the control of SIX1 activity
in fibre-type transition is currently unclear.
Postnatal control of muscle mass
The adaptive changes of muscle in response to changes
in activity not only determine the contractile phenotype
but also importantly affect muscle protein turnover and
thus muscle mass. Muscle mass increases by hypertrophy
(increased cellular protein content) and is controlled by
anabolic and catabolic mechanisms, which regulate the
increased synthesis of muscle proteins or their degra­
dation, respectively. The atrophic loss of muscle mass
often responds to inverse stimuli and can be triggered
by disuse, microgravity, catabolic steroids such as gluco­
corticoids, cytokines such as tumour necrosis factor
(TNF), genetic factors, acidosis and catabolic nutritional
states. Hypertrophy and atrophy are associated with
changes in sarcomeric protein composition (BOX 3), meta­
bolic enzymes and contractile phenotype, and atroph­y
especially can be regarded as an extreme end-stage of
activity-regulated adaptation. In fact, disuse­d muscle loss
is associated with a shift towards a fast-twitch pheno­
type and the reactivation of developmental transcription
factors4. The coordinated changes of transcriptional and
splice mechanisms, protein turnover and cell fate inte­
grates signalling pathways from hormone and cytokine
receptors, as well as the sarcomere itself.
Satellite cells play important parts in skeletal muscle
repair 7,77,78 and thus, in the long-term, homeostasis of
muscl­e tissue. They are also thought to be responsible for
early postnatal muscle growth. By contrast, their role in
eliciting hypertrophic growth is disputed, as hypertrophic
growth in adults seems to occur without satellite cell acti­
vation79. Owing to the large number of recent studies on
satellite cells, which would require a separate review, and
their unclear role in regulation of postnatal muscle mass,
we do not discuss satellite cells further.
Transcriptional control of muscle protein turnover.
Recently, the expression of catabolic genes was found
to be regulated by the transcription factor forkhead box
O1 (FOXO1) and FOXO3A80,81. The activity of FOXO
famil­y proteins is predominantly regulated by phospho­
rylation: dephosphorylated FOXOs translocate to the
nucleus and are transcriptionally active, whereas phos­
phorylated FOXOs are sequestered in the cytoplasm.
Signalling through several receptor Tyr kinases activates
the phospho­inositide 3‑kinase (PI3K)–AKT pathway,
a key regulator of muscle mass and metabolism that
directly stimulates protein synthesis by activating mam­
malian target of rapamycin (mTOR) and its downstream
targets82 (FIG. 4). Apart from its anabolic effects on protein
synthesis and carbohydrate metabolism, AKT-mediated
phosphoryl­ation also blocks FOXO1 and FOXO3A
nuclear import and transcriptional activity.
Genes depending on FOXO proteins for their expres­
sion in muscle include a range of atrophy-related genes
known as atrogenes, such as atrogin 1 (also known as
FBXO32 and MAFBX1) and the muscle-upregulated RING
finger (MURF) ubiquitin ligases82. The protein products
of atrogenes target myofibrillar, metabolic and transcrip­
tional proteins for degradation by both the ubiquiti­n–
proteasome system and the autophagy–lysosomal system83.
FOXO3A, which seems to be the dominant FOXO protein
in muscles, also controls the transcription of ubiquitous
autophagy-related genes (which can also be regarded as
atrogenes), such as those genes encodin­g microtubuleassociated protein 1 light chain 3 (LC3), sequestosome 1
(SQSTM1) and probably the related protein NBR1, and
BNIP3 — BNIP3 seems to mediate the effect of FOXO3
on autophagy 84,85. Therefore, repressing the transcription
of atrogenes augments the AKT-mediated increase of pro­
tein translation by shutting down protein breakdown, thus
increasing muscle mass.
Apart from AKT signalling, many pathways with input
from both catabolic and anabolic signals affect FOXOmediated transcription and therefore atrogene expression
(FIG. 4). A direct amplification seems to be provided by
atrogin 1, which triggers the degradation of calcineurin86,
thus inhibiting the anabolic transcriptional signals medi­
ated by the calcineurin–NFAT pathway (see above).
Furthermore, although the transfer of Lys48‑linked
polyubiquitin chains generally targets proteins for protea­
somal degradation, atrogin 1 in cardiac muscle mediates
NATURE REVIEWS | MOLECULAR CELL BIOLOGY
VOLUME 12 | JUNE 2011 | 355
© 2011 Macmillan Publishers Limited. All rights reserved
REVIEWS
Lys63‑linked polyubiquitylation of FOXO1 and FOXO3A,
which leads to their activation independently of AKT
and provides an atrogin 1-dependen­t positive feedback
mechanism for atrophic gene expression87. However,
C
+)(
although these mechanisms have been linked mainly to
muscle atrophy, it is now emerging that autophagy has
an important role not only under the extreme condi­
tions of atrophic muscle remodelling but also in muscle
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Figure 4 | Control of postnatal muscle transcription and protein turnover. a | Protein synthesis is transcriptionally
0CVWTG4GXKGYU^/QNGEWNCT%GNN$KQNQI[
regulated by anabolic transcription factors (shown in green), including myogenic regulatory
factors, transcription factors
such as serum response factor (SRF), JUNB, CLOCK and nuclear factor of activated T cells (NFAT), and transcriptional
cofactors and modifiers such as muscle LIM protein (MLP) and the muscle ankyrin repeat proteins (MARPs). Several of these
are multi-compartment proteins shuttling to the nucleus from the sarcomere, where they mostly associate with the
Z‑disk or I‑band and can thus respond to mechanical strain-induced conformational changes in the sarcomere. Protein
degradation is controlled by a set of catabolic transcription factors (shown in magenta) that include forkhead box protein
O1 (FOXO1), FOXO3A, myogenin (MYOG), KLF14, nuclear factor-κB (NF‑κB), SMAD2 and SMAD3. FOXO proteins and
NF‑κB regulate the transcription of atrogenes (shown in orange): atrogin 1, muscle-upregulated RING finger (MURF)
ubiquitin ligases, and components of the autophagy pathway such as microtubule-associated protein 1 light chain 3 (LC3),
BNIP3, sequestosome 1 (SQSTM1) and possibly NBR1. In addition, the atrogenes MURF1, MURF2 and SQSTM1 translocate
to the nucleus and repress SRF activity, presumably by MURF ubiquitin ligase activity. The anabolic regulation of protein
synthesis by the phosphoinositide 3‑kinase (PI3K)–AKT cascade represses the degradation signals mediated by FOXO
proteins. The postnatal pro-atrophic action of myogenin is downstream and redundant with histone deacetylase 4 (HDAC4)
and HDAC5. Several amplifying feedback mechanisms exist that either repress or augment anabolic or catabolic signals,
including SRF-mediated expression of miR‑486, which represses FOXO proteins in hypertrophy, and FOXO proteinmediated expression of miR‑1, which represses insulin-like growth factor 1 (IGF1) in atrophy. Additional inputs exist
through the activity of hormones and cytokines, especially anabolic signalling through IGF1, leptin and androgen receptor
(AR), and catabolic signals through tumour necrosis factor receptor (TNFR), myostatin and glucocorticoid receptor (GR).
b | Fibre-type remodelling depends largely on the sensing of Ca2+ levels by calcineurin and the resultant modulation of
NFAT and myogenic factor 5 (MYF5) activity; this is antagonized by the myozenins (MYOZs), which are sarcomeric
calcineurin inhibitors. c | Ca2+, the activator of muscle contraction, also indirectly regulates the mechanical feedback
systems embedded in the sarcomere. Titin has emerged as a sarcomeric strain-sensor at the Z‑disk, I‑band and M‑band
that feeds into the activity and localization of transcription factors and atrogenes. Solid lines represent direct actions
and dashed lines represent indirect actions. Bold dashed arrows represent receptor-linked signals. CBFβ, core-binding
factor β; IKK, inhibitor of NF‑κB kinase; MEF2C, myocyte enhancer factor 2C; mTOR, mammalian target of rapamycin;
PTEN, phosphatase and tensin homologue.
356 | JUNE 2011 | VOLUME 12
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Box 3 | Transcriptional control of sarcomere assembly
The concerted interplay of transcription factors and the modulation of their action on
the transcript level by microRNAs orchestrate the final functional differentiation of
muscle cells: the formation of contractile structures in sarcomeres and myofibrils
(FIG. 1). The assembly (and disassembly) of these multiprotein complexes (sarcomere
assembly or sarcomerogenesis) follows ordered pathways143, regulated on the
transcriptional, translational and post-translational levels. Furthermore, myofibril
assembly involves the participation of transient scaffolds and adaptors, notably the
microtubule network, which seems to be necessary both for the directional dispersion
of protein intermediates and for that of mRNAs (reviewed in REF. 1). The highly ordered
assembly of sarcomeres also requires the giant molecular ruler protein titin, which has
several functions: it forms a giant scaffold that interacts with almost all other
sarcomeric proteins; it acts as an elastic spring; and it is a signal transduction protein
acting through its carboxy-terminal protein kinase domain94,144. Therefore, it is
unsurprising that titin depletion or truncation disrupts sarcomere assembly
(reviewed in REF. 145).
The appearance of sarcomeric proteins, notably the contractile proteins actin and
myosin and the structural proteins of the Z‑disk and M‑line (myomesin and α‑actinin,
respectively) follows a strictly ordered programme of sequential transcription,
translation and incorporation into nascent cytoskeletal structures, with many
components only transiently associating with intermediates of the assembly process.
The actin filament system and actin-associated cytoskeletal proteins — such as
α‑actinin and the Z‑disk portion of titin — assemble first, followed by the formation
of an M‑line scaffold of titin and myomesin and the integration of myosin filaments
(reviewed in REF. 1). This ordered sequence of the appearance and incorporation
of sarcomeric proteins seems to be due to both transcriptional and translational
mechanisms (which regulate the sequential appearance of sarcomeric proteins) as well
as post-translational control mechanisms that coordinate their ordered incorporation
into assembling sarcomeres. Recently, it emerged that the expression of genes for
different parts of the sarcomere is regulated by different transcription factors.
Although early myogenic differentiation requires, among other factors, myocyte
enhancer factor 2A (MEF2A), the expression of myosin filament and M‑line proteins
is under the control of the late factor MEF2C146,147. The actin filament proteins are
regulated independently and require serum response factor (SRF), which is necessary
for de novo sarcomere assembly in the embryo because it mediates the transcription
of other of key sarcomeric proteins such as troponin C148. SRF also has a pivotal role in
postnatal hypertrophic growth of differentiated cardiac and skeletal muscle61,62,149.
maintenance, by removing defective organelles and con­
tributing to energy homeostasis88. The expression of
some atrogenes is also regulated by nuclear factor‑κB
(NF‑κB) (reviewed in REF. 89), thereby linking catabolic
signalling and cytokine signalling (for example, through
TNF) (FIG. 4). How FOXO proteins and NF‑κB signal­
ling are coordinated is currently unclear — the deletion
of the upstream activating kinase of NF‑κB, inhibitor of
NF‑κΒ kinase B (IKKB), protects muscle from atroph­y 90,
whereas constitutively active FOXO protein­s are sufficien­t
to induce atrophy 80,81.
Myogenin, which as discussed above is a key regu­lator
of muscle development, also has a key role in muscle atro­
phy by directly regulating the transcription of MURF1
(also known as TRIM63) and atrogin 1 (REFS 64,91).
Myogenin is rapidly upregulated in denervated muscle,
and its trans­criptional actions on atrogenes are appar­
ently controlled by HDAC4 and HDAC5 (REF. 64). This
provides a bridge between early muscle development
and postnatal adaptation of muscle. The catabolic pro­
grammes controlled by FOXO proteins, myogenin and
NF‑κB are in balance with a network of muscle-specific
and pleiotropic transcription factors that cooperate in
only partly understood ways to control the maintenance
of the differentiated pheno­typ­­e (FIG. 4). Of the pleiotropic
factors, SRF is required for postnatal hypertrophy, and
JUNB promotes muscle growth partly by directly inter­
acting with and repressing FOXO proteins 63. Given the
extensive involvement of miRNAs in feedback regulation
during development, it is not surprising that miRNAs
also have roles in muscle maintenance and remodellin­g.
In fact, miR‑486, which induces myoblast differentiation
by inhibiting PAX7 (see above), promotes PI3K–AKT
signalling by downregulating phosphatase and tensin
homologue (PTEN; which inhibits PI3K) and directly
inhibits FOXO1A itself 92. miR‑1 also amplifies the
atrophic response; it is trans­criptionally regu­lated by
FOXO proteins and suppresses insulin growth factor 1
(IGF1) expression, thus inhibitin­g AKT signalling 93
(FIG. 4). These pathways are extensively modulated by
mechanical stress and cytokine and hormona­l signals
(see below).
Mechanical control of protein turnover. Slow- and fasttwitch fibres show distinct mechanical behaviour, but the
mechanisms regulating their differentiation and adaption
are also partly sensitive to mechanical stress. The adap­
tive changes of muscle to changes in mechanical load and
activity not only affect the contractile phenotype but also
have an impact on muscle mass.
Recent progress shows that hypertrophic and atrophic
signalling pathways communicate with several ‘hubs’
within the sarcomere. Mechanical forces seem to have
important roles in modulating the conformation and thus
the activity of protein complexes94,95. Several transcription
factors and transcriptional modifiers communicate with
mechanosensors embedded in the sarcomere. The Z‑disk
coordinates several direct links to mechanically modulated
transcriptional regulation. MOYZ2, a negative regulator of
calcineurin activity, seems to be mechanically sensitive75.
The transcriptional co-regulator muscle LIM protein
(MLP; also known as CRP3) weakly associates with the
sarcomeric cytoskeleton, but translocates to the nucleus
in response to mechanical strain, suggesting that it acts
as a transducer of mechanical strain, although whether
it is a direct mechanosensor is still controversial96–98.
CLOCK, another transcription factor that is also located
at the Z‑disk, has also been implicated in the mechanically
modulated interplay of circadian rhythm, mechanical
activity and energy demand regulation99 (FIG. 4). Similarly
to MLP, CLOCK seems to shuttle between Z‑disks and the
nucleus in response to mechanical strain and to contribute
to MYOD-dependent gene expression in the daily cycling
maintenance of adult skeletal muscle100. This activity may
be important to coordinate the actual disuse atrophy
programmes with circadian rhythms, to prevent muscle
atrophy initiation during sleep. The recent identification
of core binding factor-β (CBFβ), a key element of JUNBmediated gene expression, at the Z‑disk101 emphasizes the
fact that links between cytoskeletal mechanical stress and
anabolic gene expression seem to be a prominent feature
in postnatal muscle growth and maintenance control.
Thus, several mechanically responsive direct transcrip­
tional links exist between the sarcomeric Z‑disk and
the nucleus.
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E3 ubiquitin ligases
Mechanosensitive signal complexes are also found at
the I‑band (FIG 1). Ankyrin-repeat domain 2 (ANKRD2),
cardiac ankyrin repeat protein (CARP) and diabetesassociated ankyrin repeat protein (DARP) — collectively
known as the muscle ankyrin repeat proteins (MARPs)
— can form a complex with the I‑band proteins titin,
myopalladin and calpain 3 (REF. 102). This complex is
sensitive to stretch and muscle injury and links to the
transcriptional pathways that control cell survival and
muscle gene expression103,104 (FIG. 4). Owing to the very
elastic nature of the I‑band region of titin, it is perhaps
not surprising to find mechanically modulated transcrip­
tional links in this region, which may predominantl­y
sense the passive strain on the sarcomere.
At the M‑band, the mechanically modulated kinase
domain of titin interacts with a complex of the protein
products of the atrogenes NBR1, SQSTM1 and the
MURFs105,106. This complex dissociates under mechanica­l
arrest, and MURF1 and MURF2 (MURF3 has not yet
been analysed) translocate to the cytoplasm and the
nucleus105,107. SQSTM1 was also recently shown to trans­
locate together with MURF1 and MURF2 to the nucleus
in mechanically arrested skeletal muscle­s107. One of the
probable nuclear targets of MURFs is SRF105,108,109, suggest­
ing that the MURF-induced nuclear export and transcrip­
tional repression of SRF may contribute to amplifying the
transcriptional atrophy programme105,107,110; for example,
by suppressing SRF-dependent miR‑486 expression
(FIG. 4). The multiple sarcomeric and nuclear localiza­
tions of MURFs suggest their intracellular localization
may also be important in coordinating their diverse
functions in protein turnover regulation. Titin might
act as an activity-dependent brake on protein degrada­
tion by suppressing the expression or stability of atro­
genes (FIG. 4). Both NBR1 and SQSTM1 promote the
autophagy of poly­ubiquitylated proteins by interacting
with LC3 (REFS 111–114), but the substrates they recruit
for degradation remain to be identified. In addition, these
proteins interact with several protein kinases apart from
titin and seem to have major signalling functions115,116.
MURF1 also seems to be required for TNF-induced
reduction in skeletal muscle force development 117, sug­
gesting that MURF signalling might be involved in the
NF‑κB-mediated atrophy response89 that is a hallmark of
cachexia118. Direct links to hormone and cytokine recep­
tors, such as those between the atrogene SQSTM1 and
TNF receptor signalling, suggest a complex interplay
of mechanical and cell surface receptor-linked signal­
ling 116. Thus, the differentiated sarcomere serves as a
complex feedback device for the adaptive remodelling
of the contractile machinery and its supporting energy
metabolism in response to mechanical load, and several
mechanically active regions feed back to distinct trans­
criptional programmes. It seems likely that further such
mechano–transcriptional links will emerge.
A group of proteins that
mediate the transfer of
ubiquitin, often by linking the
catalytic activity of an E2
transferase to recognition
and binding of the specific
substrates.
Hormonal control of muscle growth. Muscle mass can
also be regulated by hormones. One of the most promi­
nent known muscle growth factors is IGF1, which
is secreted by myocytes in an autocrine manner in
response to mechanical strain as a muscle-specific splice
Cachexia
A syndrome of muscle loss that
is usually caused by increased
catabolic metabolism.
variant 119,120, or by the liver. Signalling through IGF1
receptors activates the PI3K–AKT signalling pathway
(FIG. 4) and thus represses FOXO protein activity, pro­
moting muscle growth. The activity of IGF1 on the AKT
pathway is antagonized by myostatin (also known as
GDF8) (reviewed in REF. 121) primarily in differentiated
fibres122–124. The importance of myostatin as a negative
regulator of muscle mass is highlighted by its increased
serum levels in patients with heart failure, which leads
to skeletal muscle cachexia that is completely blocked
by genetic deletion of the gene encoding myostatin in
heart tissue125,126. The myostatin-activated transcrip­
tion factors SMAD2 and SMAD3 relay downstream
myostatin signalling into an atrophy programme that
also depends on FOXO proteins 123 and that is amplified
by the inhibition of AKT and mTOR signalling 122. These
observations suggest that the myostatin and AKT path­
ways crosstalk at the cell signalling and transcriptional
levels (FIG. 4). Intriguingly, FOXO protein-dependent
atrophy through SMAD2 and SMAD3 is independent
of MURF1 and atrogin 1, but also leads to a decrease
in the levels of these atrogenes122,123. Whether atrophy
is then promoted by the activation of other E3 ubiquitin
ligases or by the repression of pro-hypertrophic pathway­s
remains unclear.
Another hormone that affects muscle mass is leptin,
a major regulator of energy intake and expenditure.
Leptin was shown to positively regulate muscle mass
by suppressing the activity of FOXO3A, further dem­
onstrating how muscle and fat tissue metabolism are
interlinked127 (FIG. 4).
A further hormonal input feeding into the AKT–
FOXO protein nexus occurs by signalling through the
cytoplasmic steroid receptors, the anabolic androgen
receptor and the catabolic glucocorticoid receptor. The
muscle anabolic effects of androgens have long been
known; however, it has only recently been shown, using
muscle-specific androgen receptor‑knockout mice,
that the myocytic androgen receptor is required for
the production of the androgen-induced IGF1 isoform
IGF1EA128 and can thus regulate autocrine or paracrine
activation of the muscle AKT signalling pathway (FIG. 4).
Conversely, the glucocorticoid receptor acts upstream
of FOXO proteins and is itself regulated in a feedback
loop by the IGF1–AKT-activated mTOR129, thus pro­
viding a crosstalk loop between anabolic and catabolic
hormonal signals. This feedback loop also involves the
transcription factor KLF15, which is implicated in sev­
eral skeletal muscle metabolic processes. Exploiting such
cross-connection­s to suppress the unwanted atrophic
effects of corticoid treatment might be an interesting
practical application of this observation.
Conclusions and perspectives
The versatility and plasticity of striated muscles is due
to finely tuned networks of transcription factors and
their regulation by extracellular and intracellular cyto­
skeletal signals. The embryonic origin and the functional
remodelling of slow-twitch oxidative or fast-twitch
glyco­lytic muscle fibres has become mechanistically
clear. Similarly, the control of muscle mass in response
358 | JUNE 2011 | VOLUME 12
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© 2011 Macmillan Publishers Limited. All rights reserved
REVIEWS
Sarcopenia
The degenerative loss of
skeletal muscle mass and
strength associated with ageing
and pathological processes.
to activity has become clearer by the identification of
the major pathways controlling atrogene expression
and the cytokine and mechanical stimuli that modu­
late their activity on the gene and protein levels. Model
organisms from nematodes to vertebrates have proved
invaluable in outlining these molecular mechanisms. It
will now be important to identify how the mechanisms
specifying embryonic fibre fates are postnatally modu­
lated by activity-regulate­d pathways. The redeployment
of embryonic factors in postnatal muscle remodelling
is an exciting recent development, both for our mecha­
nistic understanding of postnatal adaptation and for
the identification of pathways amenable to pharmaco­
logical intervention. It is still an open question how
muscl­e stem cells are specified, and how this popula­
tion is maintained to ensure proper muscle homeostasis
under different physio­logical conditions. Understanding
these mechanisms in more detail may lead to the iden­
tification of molecular targets that could be exploited to
direct cellular therapies for muscular dystrophies, which
are partly manifested by depletion of the satellite cell
pool. Similarly, preventing dysfunction or decline of the
number of satellite cells during ageing could ameliorate
sarcopenia, a major cause of age-related disability.
Many of the pathways controlling postnatal muscle
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molecular analysis of cardiac atrophy and remodelling
to a new level. However, we still need to understan­d the
complex interplay of hypertrophic and atrophic factors,
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Acknowledgements
We acknowledge the contributions that have also advanced
the field by those authors whose work we could not cite owing
to space constraints. Work in the authors’ laboratories was
funded by: the UK Medical Research Council; the British
Heart Foundation; The Excellence Cluster Cardio-Pulmonary
System (ECCPS) of the Justus-Liebig-University, Giessen,
Germany, the Goethe University, Frankfurt, Germany and the
Max-Planck-Institute for Heart and Lung Research (DFG)
Germany; and the University of Giessen and the University of
Marburg Lung Center (UGMLC), funded by the government
of the state of Hessen, Germany.
Competing interests statement
The authors declare no competing financial interests.
FURTHER
INFORMATION
Thomas Braun’s homepage:
http://www.mpg.de/369653/herz_lungenforschung_wissM10
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http://www.kcl.ac.uk/schools/biohealth/research/randall/
res-sections/musclesig/gautel/mgautel
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