Myogenin and Class II HDACs
Control Neurogenic Muscle Atrophy
by Inducing E3 Ubiquitin Ligases
Viviana Moresi,1Andrew H. Williams,1Eric Meadows,4Jesse M. Flynn,4Matthew J. Potthoff,1John McAnally,1
John M. Shelton,2Johannes Backs,1,5William H. Klein,4James A. Richardson,1,3Rhonda Bassel-Duby,1
and Eric N. Olson1,*
1Department of Molecular Biology
2Department of Internal Medicine
3Department of Pathology
University of Texas Southwestern Medical Center, Dallas, TX 75390, USA
4Department of Biochemistry and Molecular Biology, University of Texas MD Anderson Cancer Center, Houston, TX 77030, USA
5Present address: Department of Cardiology, University of Heidelberg, 69117 Heidelberg, Germany
Maintenance of skeletal muscle structure and func-
tion requires innervation by motor neurons, such
that denervation causes muscle atrophy. We show
that myogenin, an essential regulator of muscle
development, controls neurogenic atrophy. Myoge-
nin is upregulated in skeletal muscle followingdener-
vation and regulates expression of the E3 ubiquitin
ligases MuRF1 and atrogin-1, which promote muscle
proteolysis and atrophy. Deletion of myogenin from
adult mice diminishes expression of MuRF1 and
atrogin-1 in denervated muscle and confers resis-
tance to atrophy. Mice lacking histone deacetylases
(HDACs) 4 and 5 in skeletal muscle fail to upregulate
myogenin and also preserve muscle mass following
myogenin in skeletal muscle of HDAC mutant mice
restores muscle atrophy following denervation.
Thus, myogenin plays a dual role as both a regulator
atrophy. These findings reveal a specific pathway for
muscle wasting and potential therapeutic targets for
Maintenance of muscle mass depends on a balance between
protein synthesis and degradation. Innervation of skeletal
muscle fibers by motor neurons is essential for maintenance of
muscle size, structure, and function. Numerous disorders,
including amyotrophic lateral sclerosis (ALS), Guillain-Barre ´
syndrome, polio, and polyneuropathy, disrupt the nerve supply
to muscle, causing debilitating loss of muscle mass (referred to
as neurogenic atrophy) and eventual paralysis.
Loss of the nerve supply to muscle fibers results in muscle
atrophy mainly through excessive ubiquitin-mediated proteo-
lysis via the proteasome pathway (Beehler et al., 2006). Other
pathologic states and systemic disorders, including cancer,
diabetes, fasting, sepsis, and disuse, also cause muscle atrophy
through ubiquitin-dependent proteolysis (Attaix et al., 2008;
Attaix et al., 2005; Medina et al., 1995; Tawa et al., 1997). The
muscle-specific E3 ubiquitin ligases MuRF1 (also called
Trim63) and atrogin-1 (also called MAFbx or Fbxo32) are
upregulated during muscle atrophy and appear to represent final
common mediators of this process (Bodine et al., 2001; Clarke
et al., 2007; Gomes et al., 2001; Kedar et al., 2004; Lecker
et al., 2004; Li et al., 2004; Li et al., 2007; Willis et al., 2009).
However, the precise molecular mechanisms and signaling
pathways that control the expression of these key regulators of
muscle protein turnover have not been fully defined and it
remains unclear whether all types of atrophic signals control
these E3 ubiquitin ligase genes through the same or different
mechanisms. Further understanding of the molecular pathways
that regulate muscle mass is a prerequisite for the development
of novel therapeutics to ameliorate muscle-wasting disorders.
Myogenin is a bHLH transcription factor essential for skeletal
muscle development (Hasty et al., 1993; Nabeshima et al.,
1993). After birth, myogenin expression is downregulated in
skeletal muscle but is reinduced in response to denervation
(Merlie et al., 1994; Tang et al., 2008; Williams et al., 2009).
Upregulation of myogenin in denervated skeletal muscle
promotes the expression of acetylcholine receptors and other
components of the neuromuscular synapse (Merlie et al., 1994;
Tang and Goldman, 2006; Williams et al., 2009). However, it
has not been possible to address the potential involvement of
myogenin in neurogenic atrophy because myogenin null mice
die at birth due to failure in skeletal muscle differentiation (Hasty
et al., 1993; Nabeshima et al., 1993).
Histone acetylation has been implicated in denervation-
dependent changes in skeletal muscle gene expression, and
histone deacetylase (HDAC) inhibitors block the expression of
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. 35
In this regard, the class IIa HDACs, HDAC4 and HDAC5, which
act as transcriptional repressors (Haberland et al., 2009; McKin-
sey et al., 2000; Potthoff et al., 2007), are upregulated in skeletal
muscle upon denervation and repress the expression of Dach2,
a negative regulator of myogenin (Cohen et al., 2007; Tang et al.,
To investigate the potential involvement of myogenin, HDAC4,
and HDAC5 in neurogenic atrophy, we performed denervation
experiments in mutant mice in which these transcriptional
regulators were deleted in adult skeletal muscle. We show
that adult mice lacking myogenin fail to upregulate the E3 ubiq-
uitin ligases MuRF1 and atrogin-1 following denervation and
are resistant to neurogenic atrophy. We demonstrate that myo-
genin binds and activates the promoter regions of the MuRF1
and atrogin-1 genes, in vitro and in vivo. Similar to adult mice
lacking myogenin, mice lacking Hdac4 and Hdac5 in skeletal
muscle do not upregulate myogenin following denervation and
are resistant to muscle atrophy. Conversely, overexpression of
myogenin in skeletal muscle is sufficient to upregulate the
expression of MuRF1 and atrogin-1 and promote neurogenic
atrophy in mice lacking Hdac4 and Hdac5. These findings reveal
a key role of myogenin and class IIa HDACs as mediators of
neurogenic atrophy and potential therapeutic targets to treat
Adult Mice Lacking Myogenin Are Resistant
to Muscle Atrophy upon Denervation
To bypass the requirement of myogenin for skeletal muscle
development and investigate its functions in muscle of adult
mice, we used a conditional myogenin null allele (Knapp et al.,
2006), which could be deleted in adult muscle with a tamox-
ifen-regulated Cre recombinase transgene (Hayashi and McMa-
hon, 2002; Knapp et al., 2006). Tamoxifen was administered to
mice at 2 months of age, and 89% deletion of the conditional
myogenin allele occurred as measured by PCR genotyping
from genomic DNA 1 week after tamoxifen injection (see
Figure S1 available online). Hereafter, we refer to these mice
with deletion of myogenin during adulthood as Myog?/?mice.
To examine the role of myogenin in denervated skeletal
muscle, the sciatic nerve was severed one month following
tamoxifen administration, and muscle atrophy was assessed
14 days later by weighing denervated and contralateral tibialis
anterior (TA) muscles. Wild-type (WT) denervated TA showed
approximately a 40% decrease in weight following denervation
in comparison to the contralateral TA (Figure 1A). In contrast,
denervated TA from Myog?/?mice showed a minimal decrease
in muscle weight (?20%) compared to the contralateral
TA (Figure 1A), suggesting that Myog?/?mice were partially
resistant to muscle atrophy. Because we deleted myogenin in
adult mice, muscle development and growth occurred normally
prior to tamoxifen administration. As expected, the muscle
weights of the nondenervated contralateral TA in Myog?/?and
WT mice were similar (WT TA = 37.82 ± 0.87 mg; Myog?/?
TA = 36.27 ± 0.54 mg; t test = 0.19). Comparable resistance to
atrophy was observed in the gastrocnemius and plantaris (GP)
weight of Myog?/?mice (Figure 1A).
Immunostaining for laminin of TA cross-sections clearly delin-
eated a decrease of muscle fiber size in the WT denervated TA in
comparison to the contralateral muscle, indicative of muscle
atrophy (Figure 1B). In contrast, the decrease in fiber size was
less evident in the Myog?/?denervated TA (Figure 1B). Morpho-
metric analysis of TA cross-sections highlighted a significant
difference in myofiber size between WT and Myog?/?muscles
following denervation, confirming the latter were resistant to
muscle atrophy (Figure 1C).
As expected, seven days after denervation, MuRF1 and
atrogin-1 expression was dramatically upregulated in the GP of
denervated WT mice (Figure 1D). Remarkably, this upregulation
suggesting that the lack of upregulation of MuRF1 and atrogin-1
in denervated Myog?/?muscles was responsible for resistance
to atrophy. Deletion of myogenin mRNA from adult Myog?/?
muscle was confirmed by real-time PCR (Figure 1D). Of note,
expression of MyoD (Myod1), another bHLH myogenic regula-
tory factor (Davis et al., 1987), was highly upregulated in both
the contralateral and denervated GP of the Myog?/?mice, seven
nin does not regulate Myod1 expression following denervation.
The dramatic upregulation of Myod1 following denervation of
Myog?/?mice, which are resistant to atrophy, also argues
against a major role of Myod1 in promoting neurogenic atrophy.
following denervation (Jason O’Rourke and E. Olson, unpub-
Denervation is known to affect skeletal myofiber composition
(Herbison et al., 1979; Midrio et al., 1992; Nwoye et al., 1982;
Patterson et al., 2006; Sandri et al., 2006; Sato et al., 2009).
To determine whether the resistance to muscle atrophy ob-
served in mice lacking myogenin was due to differences in
fiber type composition, we performed fiber type analysis of
soleus muscles 2 weeks after denervation. Our findings re-
vealed no difference in fiber type composition between WT
and Myog?/?mice (Figure S2). These findings suggest that
myogenin, which is upregulated following denervation, is
required for maximal induction of E3 ubiquitin ligase genes
and neurogenic atrophy.
We next tested whether myogenin was necessary for medi-
ating other forms of atrophy, such as occurs in response to
fasting. As shown in Figure 1E, the GP muscles of WT and
Myog?/?mice displayed comparable loss in mass following a
48 hr fast. We observed the upregulation of MuRF1 and
atrogin-1 upon fasting in both WT and Myog?/?mice and vali-
dated the deletion of myogenin in Myog?/?mice (Figure 1F).
These data clearly demonstrate that myogenin is not required
for starvation atrophy, but rather is a specific mediator of
Myogenin Activates MuRF1 and Atrogin-1 Transcription
Because upregulation of MuRF1 and atrogin-1 was impaired in
Myog?/?mice, we analyzed the promoter regions of the
MuRF1 and atrogin-1 genes for E boxes (CANNTG) that might
confersensitivity to myogenin.Indeed, threeEboxesarelocated
36 Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc.
in the promoter of the MuRF1 gene, E1 (?143 bp), E2 (?66 bp),
and E3 (?44 bp), and one conserved E box is located 79 bp
upstream of the atrogin-1 gene (Figure S3A). The E boxes
upstream of MuRF1 are contained in a genomic region near
the binding site for FoxO transcription factors (Waddell et al.,
2008), but several kilobases away from a region shown to be
regulated by NFkB (Cai et al., 2004). The E box upstream of atro-
gin-1 is embedded in a region containing multiple FoxO-binding
sites (Sandri et al., 2004).
promoters, we performed chromatin immunoprecipitation (ChIP)
assays using differentiated C2C12 myotubes, as Myogenin
Figure 1. Adult Mice Lacking Myogenin
Are Resistant to Muscle Atrophy upon
(A) Percentage of TA or GP muscle weight of
WT and Myog?/?mice 14 days after denervation,
*p < 0.05 versus WT. **p < 0.005 versus WT.
n = 4 for each sample. Data are represented as
mean ± standard error of the mean (SEM).
(B) Immunostaining for laminin of contralateral and
denervated TA of WT and Myog?/?mice, 14 days
after denervation. Scale bar = 20 microns.
(C) Morphometric analysis of contralateral and
denervated TA of WT and Myog?/?mice, 14 days
after denervation. Values indicate the mean of
cross-sectional area of denervated TA fibers as
a percentage of the contralateral fibers ± SEM.
**p < 0.005 versus WT. n = 3 cross-sections.
Myod1 in contralateral (?) and denervated (+) GP
of WT and Myog?/?mice, 7 days after denerva-
tion, detected by real-time PCR. The values are
normalized to WT contralateral GP. Data are rep-
resented as mean ± SEM. *p < 0.05; **p < 0.005
versus WT. n = 4 for each sample.
(E) Weight of GP muscle of WT and Myog?/?mice
fed (?) or fasted (+) for 48 hr. Data are represented
as mean ± SEM. **p < 0.005 versus fed GP.
NS = not significant. n = 6 for each sample.
(F) Expression of MuRF1, atrogin-1 and Myogenin
in fed (?) and 48 hr fasted (+) GP of WT and
Myog?/?mice, detected by real-time PCR. The
values are normalized to WT fed GP. Data are
represented as mean ± SEM.zp < 0.005 versus
WT. **p < 0.005 versus fed. NS = not significant.
n = 6 for each sample.
See also Figure S1 and Figure S2.
expression correlates with MuRF1 and
atrogin-1 expression during muscle cell
et al., 2000). After six days of differentia-
tion, chromatin from C2C12 myotubes
was immunoprecipitated with antibodies
against myogenin or immunoglobulin G
(IgG) as a control. Using primers flanking
the E boxes in the MuRF1 and atrogin-1
promoters, DNA was amplified by PCR
(Figure 2A and Figure S3C). Clear enrich-
ment of the corresponding promoter sequences in the DNA
immunoprecipitated with antibodies against myogenin com-
pared to IgG was indicative of myogenin binding to the endoge-
nous MuRF1 and atrogin-1 promoters.
We validated in vivo binding of myogenin to the endogenous
MuRF1 and atrogin-1 promoters by performing ChIP assays
using sonicated chromatin extracts from TA muscles harvested
from mice at 3 days and 7 days after denervation (Figure 2B
and Figure S3D). Direct binding of myogenin as a heterodimer
with E12 proteins to the E boxes E2 and E3 in the MuRF1
promoter and to the E box in the atrogin-1 promoter was shown
by gel mobility shift assays (Figure S3E).
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. 37
We further tested the ability of myogenin to activate the
MuRF1 and atrogin-1 promoter regions in vitro by constructing
luciferase reporter plasmids containing the 600 bp genomic
DNA fragment upstream of the MuRF1 gene (MuRF1-Luc) or
712 bp upstream of the atrogin-1 gene (atrogin-1-Luc) upstream
of a luciferase reporter. Mutant versions of these promoter
regions were generated by mutating the myogenin-binding sites
in the promoters. By transfecting C2C12 cells, activation of lucif-
erase was detected in response to myogenin using the wild-type
promoters (Figure 2C). This activation was blunted by mutation
of the E boxes in the promoters (Figure 2C), indicating that the
MuRF1 and atrogin-1 promoter regions contain responsive myo-
genin-binding sites. Similar results were obtained in transfected
COS1 cells (Figure S3F).
MuRF1 and Atrogin-1
(A) ChIP assay performed in C2C12 myotubes
showing myogenin binding to MuRF1 and atro-
gin-1 promoters. Chromatin was immunoprecipi-
tated with antibodies against immunogloblulin G
(IgG), or myogenin. Primers flanking the E boxes
on the MuRF1 and atrogin-1 promoters were
used for amplifying DNA by real-time PCR. Values
indicate the mean of fold enrichment over chro-
matin immunoprecipitated with antibodies against
IgG ± SEM. n = 3.
(B) ChIP assays performed using denervated TA
muscle at 3 and 7 days following denervation
show myogenin binding to the MuRF1 and
atrogin-1 promoters. Values indicate the fold
enrichment over chromatin immunoprecipitated
with antibodies against IgG.
C2C12 myoblasts transfected with luciferase
reporter plasmids ligated to the WT (MuRF1-Luc)
(atrogin-1-Luc), or the mutant constructs of
MuRF1 and atrogin-1 genes, with myogenin (+)
or empty (?) expression plasmid. Data are repre-
sented as mean ± SEM.
(D) b-galactosidase staining of contralateral and
denervated GP muscles isolated from transgenic
mice containing a lacZ transgene under the
control of the WT (MuRF1-WT-lacZ) (atrogin-1-
(atrogin-1-Emut-lacZ) constructs of the MuRF1
or atrogin-1 promoters. Upper panels show
whole muscles. Lower panels show muscle
sections. Scale bar = 20 microns.
See also Figure S3.
2. Myogenin DirectlyRegulates
To test the responsiveness of the E3
ligase gene promoters to atrophic signals
in vivo, transgenic mice were generated
harboring the same upstream regions of
the genes ligated to a lacZ reporter
(Kothary et al., 1989; Williams et al.,
2009). Transgenic mice with the mutated
versions of these
were also generated (MuRF1-Emut-lacZ
and atrogin-1-Emut-lacZ). Seven days
following denervation, b-galactosidase
expression controlled by the wild-type promoters was upregu-
lated in denervated GP muscle fibers compared to the inner-
vated contralateral leg muscles (Figure 2D). The expression of
lacZ in only a subset of myofibers likely reflects the mosaicism
of F0 transgenic mice and, perhaps, variable upregulation of
the E3 ubiquitin ligase genes in different myofibers in response
to denervation (Moriscot et al., 2010). In contrast to the obvious
upregulation of the wild-type transgenes following denervation,
mutation of the E boxes in these promoters abrogated b-galac-
tosidase expression, revealing an essential role for myogenin in
denervation-dependent activation of MuRF1 and atrogin-1
in vivo (Figure 2D). These results show that the MuRF1 and
atrogin-1 genes are targets of myogenin transcriptional activa-
tion in response to denervation.
38 Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc.
Mice Null for Class II HDACs Are Resistant to Muscle
Atrophy upon Denervation
Previous studies showed that the class II HDACs, HDAC4 and
vation (Bodine et al., 2001; Cohen et al., 2007; Tang et al., 2008)
andareresponsible fortherepressionof Dach2,anegativeregu-
the role of myogenin in promoting muscle atrophy, we hypothe-
sized that mice lacking HDAC4 or HDAC5 in skeletal muscle
would be resistant to atrophy following denervation owing to
hereafter referred to as Hdac4skKO) (Potthoff et al., 2007). The
absence of HDAC4 protein upon Hdac4 gene deletion was
confirmed by western blot analysis (Figure S4). Since mice null
for Hdac5 do not display a phenotype (Chang et al., 2004), we
experiments. Fourteen days following denervation, WT dener-
vated TA showed approximately a 50% decrease in weight in
vated TA muscles from Hdac4skKO or Hdac5 KO mice showed
a decrease of about 30% in muscle weight in comparison to
the contralateral muscles (Figure 3A), suggesting that these
mice were partially resistant to muscle atrophy. The weight of
HDAC4 and HDAC5 display functional redundancy in different
tissues and in a variety of developmental and pathological
settings (Backs et al., 2008; Haberland et al., 2009; Potthoff
et al., 2007), so we generated double knockout (dKO) mice by
crossing Hdac4skKO with Hdac5 KO mice to further investigate
the role of HDAC4 and HDAC5 in skeletal muscle atrophy.
The dKO mice were viable and fertile and showed no obvious
phenotype under normal conditions (data not shown). Strikingly,
Figure 3. HDAC4 and HDAC5 Redundantly
Regulate Skeletal Muscle Atrophy
(A) Percentage of TA muscle weight of mice of
the indicated genotype 14 days after denervation,
expressed relative to the contralateral muscle.
Data are represented
**p < 0.005 versus WT. n = 5 for each sample.
(B) Immunostaining for laminin in contralateral and
denervated TA of mice of the indicated genotype,
14 days after denervation. Scale bar = 20 microns.
(C) Morphometric analysis of contralateral and
denervated TA of indicated genotype, 14 days
after denervation. Values indicate the mean of
cross-sectional area of denervated TA fibers as
a percentage of the contralateral fibers ± SEM.
*p < 0.05 and **p < 0.005 versus WT. n = 3
See also Figure S4 and Figure S5.
fourteen days after denervation, the
TA of denervated dKO mice showed
a decrease in weight of only ?10%
(Figure 3A), revealing that the dKO mice were more resistant to
muscle atrophy compared to Hdac4skKO or Hdac5 KO mice.
The weight of the contralateral TA was comparable among the
mice (data not shown). Similar differences were also observed
among GP muscles between WT and dKO mice (Figure S5).
Immunostaining for laminin 14 days after denervation clearly
demonstrated that the denervated TA fibers from Hdac4skKO
and Hdac5 KO mice were larger than the denervated WT fibers
and that the denervated TA from dKO mice had a minimal
decrease in muscle fiber size compared to the contralateral dKO
TA (Figure 3B). Morphometric analysis on TA sections revealed
that, although WT mice showed a reduction of ?70% in the myo-
fiber cross-sectional area between denervated and contralateral
TA, Hdac4skKO denervated TA displayed ?30% reduction in
myofiber cross-sectional area. Hdac5 KO denervated TA also
showed a substantial reduction in myofiber area (?50%) when
compared to the contralateral TA, whereas in dKO mice this
reduction was only ?25% (Figure 3C). From these results, we
conclude that HDAC4 and HDAC5 redundantly regulate skeletal
muscle atrophyand micelackingtheseHDACs inskeletal muscle
are resistant to muscle atrophy upon denervation.
Aberrant Transcriptional Responses to Denervation
in HDAC Mutant Mice
We compared the transcriptional responses to denervation in
WT and dKO mice by real-time PCR analysis of denervation-
responsive transcripts. As reported previously (Cohen et al.,
2007; Tang et al., 2008), Dach2 expression was dramatically
downregulated upon denervation in WT mice. However, Dach2
was only modestly downregulated in the dKO mice (Figure 4).
Consistent with the repressive influence of Dach2 on Myogenin
expression, in WT mice, Myogenin and Myod1 were strongly
upregulated three days after denervation, as were MuRF1 and
atrogin-1 (Figure 4). In contrast, neither Myogenin nor Myod1
transcripts were upregulated following denervation of dKO
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. 39
mice (Figure 4). The upregulation of MuRF1 and atrogin-1 was
also completelyabolished indKOdenervated GP (Figure4), sug-
gesting that the lack of upregulation of MuRF1 and atrogin-1 in
denervated dKO muscles was in part responsible for resistance
Myogenin Overexpression in dKO Muscle Restores
To examine whether forced expression of myogenin was suffi-
cient to overcomethe resistance of the dKOTA muscle to dener-
vation-induced atrophy, we electroporated the TA of dKO mice
with either a myogenin expression plasmid or an empty expres-
sion plasmid. Gene delivery efficiency was monitored by coelec-
troporation with a GFP vector (Dona et al., 2003; Rana et al.,
2004). Three days after electroporation, which is sufficient time
for the electroporated plasmids to be expressed in skeletal
muscle (Dona et al., 2003), we denervated one leg of the dKO
micebycutting thesciatic nerve;theTA muscleswereharvested
10 days after denervation. As seen in Figure 5A, laminin immu-
nostaining of dKO TA muscles clearly revealed a decrease in
Figure 4. dKO Mice Show Altered Gene Expression upon Denervation
Expression of the indicated mRNAs was detected by real-time PCR in WT and dKO denervated GP and normalized to the expression in the contralateral muscle.
Data are represented as mean ± SEM. **p < 0.005 versus dKO. n = 6 for each time point.
Figure 5. Ectopic Expression of Myogenin Induces Muscle Atrophy in dKO Mice Following Denervation
(A) Immunostaining for laminin (red) of cross-section of contralateral and denervated dKO TA electroporated with GFP expression plasmid and control plasmid
(HDAC4/5 dKO Control) or GFP plasmid and myogenin (HDAC4/5 dKO + Myogenin), 10 days after denervation. Histology shows that the dKO denervated
GFP-positive fibers coelectroporated with myogenin are smaller than denervated GFP-positive fibers coelectroporated with control plasmid. Scale
bar = 20 microns.
and control plasmid (Control) or GFP plasmid and myogenin (Myogenin), 10 days after denervation. Values indicate the mean of cross-sectional area of
GFP-positive muscle fibers as a percentage of the contralateral control fibers ± SEM. *p < 0.05 versus control. n = 7 for each condition.
(C) Expression of Myogenin, MuRF1, and atrogin-1 in contralateral (?) and denervated (+) dKO TA muscles electroporated with GFP plasmid and a control
plasmid (Control) or GFP plasmid and myogenin (Myogenin), 10 days after denervation. Values are normalized to the expression in the contralateral control
muscles. Data are represented as mean ± SEM. *p < 0.05 versus control. n = 3 for each sample.
See also Figure S6.
40 Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc.
myofiber size in the denervated TA of dKO mice overexpressing
myogenin compared to the denervated dKO TA electroporated
with the control vector. Morphometric analysis performed on
GFP-positive myofibers showed a significant decrease in the
size of myofibers of the denervated dKO TA electroporated
with myogenin versus control vector (Figure 5B). Real-time
PCR analysis validated the overexpression of Myogenin in elec-
troporated TA muscle of dKO mice and showed an upregulation
the myogenin-dependent regulation of the E3 ubiquitin ligases.
The potential role of myogenin in driving muscle atrophy was
further investigated by overexpressing myogenin in the TA
muscle of WT mice. Morphometric analysis performed on
GFP-positive myofibers showed no significant size difference
between myofibers electroporated with control or myogenin
expression plasmid (Figures S6A and S6B). Real-time PCR anal-
ysis validated the overexpression of myogenin in electroporated
TA muscle of WT mice and showed an upregulation of the
expression of MuRF1 and atrogin-1 (Figure S6C). Taken
genin is necessary but not sufficient to induce muscle atrophy.
known for its function as an essential regulator of myogenesis, in
controlling neurogenic atrophy. Myogenin promotes muscle
atrophy upon denervation by directly activating the expression
of MuRF1 and atrogin-1, which encode E3 ubiquitin ligases
responsible for muscle proteolysis. Upregulation of Myogenin
in response to denervation is controlled by a transcriptional
pathway in which HDAC4 and 5 are initially induced and, in
turn, repress the expression of Dach2 (Tang and Goldman,
2006), a negative regulator of Myogenin (Figure 6).
It is generally accepted that muscle atrophy occurs when
proteolysis exceeds protein synthesis (Eley and Tisdale, 2007;
Glass, 2003; Mammucari et al., 2008; Sandri et al., 2004). Up-
regulation of myogenin in response to denervation has been
proposed as an adaptive mechanism to prevent muscle atrophy
Figure 6. Model for Neurogenic Atrophy
Denervation of skeletal muscle results in the upregulation
regulator of myogenin, resulting in Myogenin expression.
Myogenin activates the expression of MuRF1 and atro-
gin-1, two E3 ubiquitin ligases that participate in the
proteolytic pathway resulting in muscle atrophy. Myoge-
nin also regulates miR-206, which establishes a negative
(Hyatt et al., 2003; Ishido et al., 2004). On the
contrary, we demonstrate here that myogenin
directly regulates MuRF1 and atrogin-1, which
promote the loss of muscle mass in response
to denervation, revealing a mechanistic basis
for neurogenic muscle atrophy and a previously
unrecognized function for myogenin in this
Recently, we showed that microRNA (miR) 206 is also upregu-
lated in denervated skeletal muscle via a series of conserved E
boxes that bind myogenin (Williams et al., 2009). miR-206, in
turn, represses expression of HDAC4 and controls a retrograde
signaling pathway that promotes reinnervation of denervated
tion by activating an elaborate network of transcriptional and
epigenetic pathways, involving positive and negative feedback
loops, which modulate nerve-muscle interactions and muscle
growth and function (Figure 6).
Dual Roles of Myogenin in Muscle Development
Our findings reveal the gene regulatory circuitry for muscle
development is redeployed in adulthood to control aspects of
muscle disease and stress responsiveness. Thus, myogenin
differentiation ordegradation—dependingon thedevelopmental
or pathological setting. These contrasting activities of myogenin
likely reflect differential modulation by signaling pathways and
cofactors that enable myogenin to regulate distinct sets of target
Similar to myogenin, Dach2 is a transcription factor involved in
both muscle development and muscle atrophy. Dach2 is
expressed in the developing somites prior to the onset of
myogenesis and has been shown to regulate myogenic specifi-
cation by interacting with the Eya2 and Six1 transcription factors
(Heanue et al., 1999; Kardon et al., 2002). Indeed, Dach proteins
are required for activation of Six1 targets (Li et al., 2003), sug-
gesting a possible role of Dach proteins in the Six1-mediated
regulation of muscle development (Laclef et al., 2003) or fiber
type specification (Grifone et al., 2004). Following denervation,
Dach2 plays a role in connecting neuronal activity with myogenin
expression (Cohen et al., 2007; Tang and Goldman, 2006; Tang
et al., 2008).
The finding that forced expression of myogenin in HDAC4/5
mutant mice is sufficient to restore muscle atrophy following
denervation indicates that myogenin is a key downstream
mediator of the proatrophic functions of these HDACs. It is
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. 41
noteworthy, however, that the blockade to muscle atrophy and
E3 ligase expression imposed by the combined deletion of
HDACs 4 and 5 is more pronounced than in Myog?/?mice.
This suggests the existence of additional downstream targets
of these HDACs that promote neurogenic atrophy. We also
note that forced overexpression of myogenin in innervated skel-
etal muscle was not sufficient to induce muscle atrophy
(Figure S6) (Hughes et al., 1999). These findings indicate that
myogenin is necessary, but not sufficient, to regulate the genetic
denervation-dependent signals that potentiate the ability of
myogenin to promote atrophy.
MyoD, like myogenin, is upregulated in response to denerva-
tion (Figure 4 and (Charge et al., 2008; Hyatt et al., 2003; Ishido
et al., 2004). In Myog?/?mice, Myod1 expression is dramatically
elevated compared to WT muscles and is super-induced in
response to denervation (Figure 1D). The observation that
Myod1 null mice are not resistant to muscle atrophy following
denervation (Jason O’Rourke and E. Olson, unpublished data)
demonstrates a negligible role for Myod1 in neurogenic atrophy
and points to myogenin as the major myogenic bHLH factor
involved in this process. This is consistent with the finding that,
although MyoD and myogenin bind the same DNA consensus
sequences, they regulate distinct sets of target genes (Blais
et al., 2005; Cao et al., 2006).
A Myogenin-Dependent Transcriptional Pathway
for Muscle Atrophy
We show, both in vivo using denervated muscles and in vitro
using differentiated C2C12 cells, that myogenin binds the
endogenous MuRF1 and atrogin-1 promoters. We observed
a decrease in myogenin expression and binding to these E3
vation (Figure 2B and Figure 4), suggesting an especially impor-
tant role of myogenin in triggering the transcriptional cascade
leading to atrophy. Consistent with our finding that myogenin
regulates MuRF1 and atrogin-1 expression, these E3 ubiquitin
ligases are upregulated upon C2C12 differentiation (Figure S3B)
(Spencer et al., 2000), a process known to be regulated by
myogenin. Although it is well established that MuRF1 and atro-
gin-1 function in driving skeletal muscle atrophy (Bodine et al.,
2001; Clarke et al., 2007; Gomes et al., 2001; Kedar et al.,
2004; Lecker et al., 2004; Li et al., 2004; Li et al., 2007; Willis
et al., 2009), their potential roles in myogenesis have not been
explored. Considering the important role of ubiquitination in
regulating proteolysis, endocytosis, signal transduction (Hicke,
2001), and transcription (Salghetti et al., 2001), it will be inter-
esting to investigate the potential involvement of MuRF1 and
atrogin-1 in muscle development and regeneration.
Numerous disorders, including motor neuron disease, fasting,
cancer cachexia, and sarcopenia, cause muscle atrophy and
the E3 ubiquitin ligase genes are thought to function as final
common mediators of different atrophic stimuli. Myogenin is
upregulated upon denervation and spinal cord isolation (Hyatt
et al., 2003), but is not induced in response to other forms of
atrophy, such as fasting, cancer cachexia, or diabetes (Lecker
et al., 2004; Sacheck et al., 2007). In this regard, we have found
that Myog?/?mice display a normal lossof skeletal muscle mass
in response to fasting, further demonstrating that myogenin is
dedicated to neurogenic atrophy and sensing the state of motor
innervation. The fact that MuRF1 and atrogin-1 are upregulated
in other atrophy conditions in the absence of myogenin upregu-
lation (Lecker et al., 2004; Sacheck et al., 2007) strongly
suggests that other transcription factors known to regulate the
expression of these ubiquitin ligases, such as the FoxO family
or NFkB (Bodine et al., 2001; Sandri et al., 2004; Waddell
et al., 2008), play a role in driving muscle atrophy in a myoge-
Our finding that myogenin, in addition to HDAC4 and HDAC5,
acts as a regulator of neurogenic muscle atrophy through the
activation of E3 ubiquitin ligases provides a new perspective
on potential therapies for muscle wasting disorders. Class II
HDACs are regulated by a variety of calcium-dependent
signaling pathways that control their nuclear export through
etal.,2000).Inapathological condition suchas muscledenerva-
tion, HDAC4 and HDAC5 are upregulated, shuttle into the
myonuclei adjacent to neuromuscular junctions (Cohen et al.,
2007), and are critical regulators of muscle atrophy. Modulation
of the activity of class II HDACs, through pharmacologic inhibi-
tion compatible with the maintenance of steady-state transcrip-
tion of genes regulated by class II HDACs, may represent a new
strategy for ameliorating muscle atrophy following denervation.
Mice used in this study are described in the Extended Experimental Proce-
Inanaesthetized adult mice,thesciatic nerveoftheleft legwascutand a3mm
piece wasexcised. Theright legremained innervatedand wasusedascontrol.
Mice were sacrificed after 3, 7, 10, or 14 days.
DNA Delivery by Electroporation
For gene delivery by electroporation, adult dKO mice were anesthetized; TA
muscles exposed, injected with 30 mg of DNA in a solution of 5% mannitol,
and immediately subjected to electroporation. Electroporation was performed
by delivering 10 electric pulses of 20 V each (five with one polarity followed by
five with inverted polarity). A pair of 3 3 5 mm Genepaddle electrodes (BTX,
San Diego, CA) placed on opposite sides of the muscle was used to deliver
the electric pulses. pCMV-Snap25-GFP (provided by Tullio Pozzan, University
of Padua, Padua, Italy) was used in a 1:1 ratio with pcDNA3.1 (Invitrogen) or
EMSV-myogenin plasmid (Rana et al., 2004).
Cryosections of TA or soleus were fixed in 4% paraformaldehyde in PBS for
10 min at 4?C and washed in PBS. After incubating 30 min with 0.1%
Triton X-100 in PBS, the samples were fixed for 1 hr in 15% goat serum in
PBS supplemented with M.O.M. Mouse IgG blocking reagent (Vector Labora-
tories) (BB) at room temperature. Primary antibodieswere incubatedovernight
at 4?C (1:100 dilution of rabbit polyclonal anti-laminin antibody; 1:16000
anti-type I myosin heavy chain (MHC) (Sigma). Primary antibodies were
detected by Alexa Fluor-488 or -555 goat anti-rabbit antibody (Invitrogen)
diluted 1:800 in BB. DAB staining (Vector Laboratories) was used on soleus
muscle for detecting type I MHC. Soleus muscles were used for metachro-
matic ATPase staining as described elsewhere (Ogilvie and Feeback, 1990).
42 Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc.
Staining of transgenic lines positive for b-galactosidase was performed on GP
muscles, as previously described (Williams et al., 2009).
Myofiber area was assessed on TA cryosections using ImageJ software
(http://rsb.info.nih.gov/ij/) (NIH). Three H&E-stained cross-sections from three
different mice for each genotype were analyzed. Between 100 and 350 GFP-
positive fibers were analyzed for each electroporated TA muscle. The values
are calculated as the percentage of the average of the cross-sectional area
of each TA overthe average cross-sectional area of the contralateral TA fibers.
RNA Isolation and RT-PCR
Total RNA was isolated from GP muscles using Trizol reagent (Invitrogen)
following the manufacturer’s instructions. Three micrograms of RNA was con-
vertedtocDNA usingrandom primers and Superscript IIIreversetranscriptase
(Invitrogen). Gene expression was assessed using real-time PCR with the ABI
PRISM 7000 sequence detection system and TaqMan or with SYBR green
Master Mix reagents (Applied Biosystems). Real-time PCR values were
normalized with glyceraldehyde-3-phosphate dehydrogenase (GAPDH).
A list of Taqman probes and Sybr Green primers are available in the Extended
A list of the plasmids used in this study is available in the Extended Experi-
COS cells were grown in DMEM supplemented with 10% fetal bovine serum
(FBS) and antibiotics (100 U/ml penicillin and 100 mg/ml streptomycin).
C2C12 myoblasts were grown in DMEM supplemented with 20% FBS and
antibiotics and differentiated in DMEM supplemented with 2% horse serum
Chromatin Immunoprecipitation Assay
ChIP assays were performed using C2C12 myotubes at day six of differentia-
tion or using TA muscles three and seven days after denervation with the ChIP
assay kit (Upstate) following the manufacturer’s instructions. Chromatin was
ogenin (M-225; Santa Cruz). The sequences of the ChIP primers are available
in the Extended Experimental Procedures.
C2C12 transfections were performed using Lipofectamine 2000 (Invitrogen) as
previously described (Mercer et al., 2005). COS cells were plated and trans-
fected 12 hr later using FuGENE (Roche Applied Science) following the manu-
facturer’s instructions. The MuRF1 and atrogin-1 reporter plasmid cloning
strategy is described in the Extended Experimental Procedures. Luciferase
assays were performed with the Luciferase Assay kit (Promega) according
to the manufacturer’s instructions.
Mutations were introduced into E boxes E2 and E3 of the MuRF1 promoter
region and in the E box of the atrogin-1 promoter by using the QuikChange II
Site-Directed Mutagenesis Kit (Stratagene). The same E box mutations as
those used in electrophoretic mobility shift assays were introduced within
each E box site in the promoters.
Data are presented as mean ± standard error of the mean (SEM). Statistical
significance was determined using two-tailed t test with a significance level
minor of 0.05.
Supplemental Information includes Extended Experimental Procedures and
six figures and can be found with this article online at doi:10.1016/j.cell.
We thank Marco Sandri for scientific input, Cheryl Nolen and Svetlana
Bezprozvannaya for technical assistance, Jose Cabrera for graphics, and
Jennifer Brown for editorial assistance. Work in the laboratory of E.N.O. was
supported by grants from the National Institutes of Health and the
Robert A. Welch Foundation (grant number I-0025). W.H.K. was supported
by a grant from the Muscular Dystrophy Association and the Robert A. Welch
Foundation. J.B. was supported by the Deutsche Forschungsgemeinschaft
Received: April 20, 2010
Revised: June 1, 2010
Accepted: August 20, 2010
Published: September 30, 2010
Attaix, D., Combaret, L., Bechet, D., and Taillandier, D. (2008). Role of the
ubiquitin-proteasome pathway in muscle atrophy in cachexia. Curr. Opin.
Support. Palliat. Care 2, 262–266.
Attaix, D., Ventadour, S., Codran, A., Bechet, D., Taillandier, D., and
Combaret, L. (2005). The ubiquitin-proteasome system and skeletal muscle
wasting. Essays Biochem. 41, 173–186.
Backs, J., Backs, T., Bezprozvannaya, S., McKinsey, T.A., and Olson, E.N.
(2008). Histone deacetylase 5 acquires calcium/calmodulin-dependent kinase
II responsiveness by oligomerization with histone deacetylase 4. Mol. Cell.
Biol. 28, 3437–3445.
Beehler, B.C., Sleph, P.G., Benmassaoud, L., and Grover, G.J. (2006).
Reduction of skeletal muscle atrophy by a proteasome inhibitor in a rat model
of denervation. Exp. Biol. Med. (Maywood) 231, 335–341.
Blais, A., Tsikitis, M., Acosta-Alvear, D., Sharan, R., Kluger, Y., and Dynlacht,
B.D. (2005). An initial blueprint for myogenic differentiation. Genes Dev. 19,
Bodine, S.C., Latres, E., Baumhueter, S., Lai, V.K., Nunez, L., Clarke, B.A.,
Poueymirou, W.T., Panaro, F.J., Na, E., Dharmarajan, K., et al. (2001). Identifi-
cation of ubiquitin ligases required for skeletal muscle atrophy. Science 294,
Cai, D., Frantz, J.D., Tawa, N.E., Jr., Melendez, P.A., Oh, B.C., Lidov, H.G.,
Hasselgren, P.O., Frontera, W.R., Lee, J., Glass, D.J., et al. (2004). IKKbeta/
NF-kappaB activation causes severe muscle wasting in mice. Cell 119,
Cao, Y., Kumar, R.M., Penn, B.H., Berkes, C.A., Kooperberg, C., Boyer, L.A.,
Young, R.A., and Tapscott, S.J. (2006). Global and gene-specific analyses
show distinct roles for Myod and Myog at a common set of promoters.
EMBO J. 25, 502–511.
Chang, S., McKinsey,T.A., Zhang, C.L., Richardson, J.A., Hill, J.A., and Olson,
E.N. (2004). Histone deacetylases 5 and 9 govern responsiveness of the heart
to a subset of stress signals and play redundant roles in heart development.
Mol. Cell. Biol. 24, 8467–8476.
Charge, S.B., Brack, A.S., Bayol, S.A., and Hughes, S.M. (2008). MyoD- and
nerve-dependent maintenance of MyoD expression in mature muscle fibres
acts through the DRR/PRR element. BMC Dev. Biol. 8, 5–18.
Clarke, B.A., Drujan, D., Willis, M.S., Murphy, L.O., Corpina, R.A., Burova, E.,
Rakhilin, S.V., Stitt, T.N., Patterson, C., Latres, E., et al. (2007). The E3 Ligase
MuRF1 degrades myosin heavy chain protein in dexamethasone-treated
skeletal muscle. Cell Metab. 6, 376–385.
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. 43
Cohen, T.J., Waddell, D.S., Barrientos, T., Lu, Z., Feng, G., Cox, G.A., Bodine,
S.C., and Yao, T.P. (2007). The histone deacetylase HDAC4 connects neural
activity to muscle transcriptional reprogramming. J. Biol. Chem. 282, 33752–
Davis, R.L., Weintraub, H., and Lassar, A.B. (1987). Expression of a single
transfected cDNA converts fibroblasts to myoblasts. Cell 51, 987–1000.
Dona, M., Sandri, M., Rossini, K., Dell’Aica, I., Podhorska-Okolow, M., and
Carraro, U. (2003). Functional in vivo gene transfer into the myofibers of adult
skeletal muscle. Biochem. Biophys. Res. Commun. 312, 1132–1138.
Eley, H.L., and Tisdale, M.J. (2007). Skeletal muscle atrophy, a link between
depression of protein synthesis and increase in degradation. J. Biol. Chem.
Glass, D.J. (2003). Molecular mechanisms modulating muscle mass. Trends
Mol. Med. 9, 344–350.
Gomes, M.D., Lecker, S.H., Jagoe, R.T., Navon, A., and Goldberg, A.L. (2001).
Atrogin-1, a muscle-specific F-box protein highly expressed during muscle
atrophy. Proc. Natl. Acad. Sci. USA 98, 14440–14445.
Grifone, R., Laclef, C., Spitz, F., Lopez, S., Demignon, J., Guidotti, J.E.,
Kawakami, K., Xu, P.X., Kelly, R., Petrof, B.J., et al. (2004). Six1 and Eya1
expression can reprogram adult muscle from the slow-twitch phenotype into
the fast-twitch phenotype. Mol. Cell. Biol. 24, 6253–6267.
Haberland, M., Montgomery, R.L., and Olson, E.N. (2009). The many roles of
histone deacetylases in development and physiology: implications for disease
and therapy. Nat. Rev. Genet. 10, 32–42.
Hasty, P., Bradley, A., Morris, J.H., Edmondson, D.G., Venuti, J.M., Olson,
E.N., and Klein, W.H. (1993). Muscle deficiency and neonatal death in mice
with a targeted mutation in the myogenin gene. Nature 364, 501–506.
Hayashi, S., and McMahon, A.P. (2002). Efficient recombination in diverse
tissues by a tamoxifen-inducible form of Cre: a tool for temporally regulated
gene activation/inactivation in the mouse. Dev. Biol. 244, 305–318.
Heanue, T.A., Reshef, R., Davis, R.J., Mardon, G., Oliver, G., Tomarev, S.,
Lassar, A.B., and Tabin, C.J. (1999). Synergistic regulation of vertebrate
muscle development by Dach2, Eya2, and Six1, homologs of genes required
for Drosophila eye formation. Genes Dev. 13, 3231–3243.
Herbison, G.J., Jaweed, M.M., and Ditunno,J.F. (1979). Muscle atrophy in rats
following denervation, casting, inflammation, and tenotomy. Arch. Phys. Med.
Rehabil. 60, 401–404.
Hughes, S.M., Chi, M.M., Lowry, O.H., and Gundersen, K. (1999). Myogenin
induces a shift of enzyme activity from glycolytic to oxidative metabolism in
muscles of transgenic mice. J. Cell Biol. 145, 633–642.
Hyatt, J.P., Roy, R.R., Baldwin, K.M., and Edgerton, V.R. (2003). Nerve
activity-independent regulation of skeletal muscle atrophy: role of MyoD and
myogenin in satellite cells and myonuclei. Am. J. Physiol. Cell Physiol. 285,
Ishido, M., Kami, K., and Masuhara, M. (2004). In vivo expression patterns of
MyoD, p21, and Rb proteins in myonuclei and satellite cells of denervated
rat skeletal muscle. Am. J. Physiol. Cell Physiol. 287, C484–C493.
Kardon, G., Heanue, T.A., and Tabin, C.J. (2002). Pax3 and Dach2 positive
regulation in the developing somite. Dev. Dyn. 224, 350–355.
Kedar, V., McDonough, H., Arya, R., Li, H.H., Rockman, H.A., and Patterson,
C. (2004). Muscle-specific RING finger 1 is a bona fide ubiquitin ligase that
degrades cardiac troponin I. Proc. Natl. Acad. Sci. USA 101, 18135–18140.
Knapp, J.R., Davie, J.K., Myer, A., Meadows, E., Olson, E.N., and Klein, W.H.
(2006). Loss of myogenin in postnatal life leads to normal skeletal muscle but
reduced body size. Development 133, 601–610.
Kothary, R., Clapoff, S., Darling, S., Perry, M.D., Moran, L.A., and Rossant, J.
(1989). Inducible expression of an hsp68-lacZ hybrid gene in transgenic mice.
Development 105, 707–714.
Laclef, C., Hamard, G., Demignon, J., Souil, E., Houbron, C., and Maire, P.
(2003). Altered myogenesis in Six1-deficient mice. Development 130, 2239–
Lecker, S.H., Jagoe, R.T.,Gilbert, A.,Gomes,M., Baracos, V.,Bailey, J., Price,
S.R., Mitch, W.E., and Goldberg, A.L. (2004). Multiple types of skeletal muscle
atrophy involve a common program of changes in gene expression. FASEB J.
Li, H.H., Kedar, V., Zhang, C., McDonough, H., Arya, R., Wang, D.Z., and
Patterson, C. (2004). Atrogin-1/muscle atrophy F-box inhibits calcineurin-
dependent cardiac hypertrophy by participating in an SCF ubiquitin ligase
complex. J. Clin. Invest. 114, 1058–1071.
Li, H.H., Willis, M.S., Lockyer, P., Miller, N., McDonough, H., Glass, D.J., and
Patterson, C. (2007). Atrogin-1 inhibits Akt-dependent cardiac hypertrophy in
Li, X., Oghi, K.A., Zhang, J., Krones, A., Bush, K.T., Glass, C.K., Nigam, S.K.,
Aggarwal, A.K., Maas, R., Rose, D.W., et al. (2003). Eya protein phosphatase
activity regulates Six1-Dach-Eya transcriptional effects in mammalian
organogenesis. Nature 426, 247–254.
Lu, J., Webb, R., Richardson, J.A., and Olson, E.N. (1999). MyoR: a muscle-
restricted basic helix-loop-helix transcription factor that antagonizes the
actions of MyoD. Proc. Natl. Acad. Sci. USA 19, 552–557.
Mammucari, C., Schiaffino, S., and Sandri, M. (2008). Downstream of Akt:
McKinsey, T.A., Zhang, C.L., Lu, J., and Olson, E.N. (2000). Signal-dependent
nuclear export of a histone deacetylase regulates muscle differentiation.
Nature 408, 106–111.
Medina, R., Wing, S.S., and Goldberg, A.L. (1995). Increase in levels of polyu-
biquitin and proteasome mRNA in skeletal muscle during starvation and
denervation atrophy. Biochem. J. 307, 631–637.
Mercer, S.E., Ewton, D.Z., Deng, X., Lim, S., Mazur, T.R., and Friedman, E.
(2005). Mirk/Dyrk1B mediates survival during the differentiation of C2C12
myoblasts. J. Biol. Chem. 280, 25788–25801.
Merlie, J.P., Mudd, J., Cheng, T.C., and Olson, E.N. (1994). Myogenin and
acetylcholine receptor alpha gene promoters mediate transcriptional regula-
tion in response to motor innervation. J. Biol. Chem. 269, 2461–2467.
Midrio, M., Danieli-Betto, D., Megighian, A., Velussi, C., Catani, C., and
Carraro, U. (1992). Slow-to-fast transformation of denervated soleus muscle
of the rat, in the presence of an antifibrillatory drug. Pflugers Arch. 420,
Moriscot, A.S., Baptista, I.L., Bogomolovas, J., Witt, C., Hirner, S., Granzier,
H., and Labeit, S. (2010). MuRF1 is a muscle fiber-type II associated factor
and together with MuRF2 regulates type-II fiber trophicity and maintenance.
J. Struct. Biol. 170, 344–353.
Nabeshima, Y., Hanaoka, K., Hayasaka, M., Esumi, E., Li, S., and Nonaka, I.
(1993). Myogenin gene disruption results in perinatal lethality because of
severe muscle defect. Nature 364, 532–535.
Nwoye, L., Mommaerts, W.F., Simpson, D.R., Seraydarian, K., and Marusich,
M. (1982). Evidence for a direct action of thyroid hormone in specifying muscle
properties. Am. J. Physiol. 242, R401–R408.
Ogilvie, R.W., and Feeback, D.L. (1990). A metachromatic dye-ATPase
method for the simultaneous identification of skeletal muscle fiber types I,
IIA, IIB and IIC. Stain Technol. 65, 231–241.
produces different single fiber phenotypes in fast- and slow-twitch hindlimb
muscles of the rat. Am. J. Physiol. Cell Physiol. 291, C518–C528.
Potthoff, M.J., Wu, H., Arnold, M.A., Shelton, J.M., Backs, J., McAnally, J.,
Richardson, J.A., Bassel-Duby, R., and Olson, E.N. (2007). Histone deacety-
lase degradation and MEF2 activation promote the formation of slow-twitch
myofibers. J. Clin. Invest. 117, 2459–2467.
44 Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc.
poration of plasmid mixtures into muscle in vivo. Acta Physiol. Scand. 181,
Sacheck, J.M., Hyatt, J.P., Raffaello, A., Jagoe, R.T., Roy, R.R., Edgerton,
V.R., Lecker, S.H., and Goldberg, A.L. (2007). Rapid disuse and denervation
atrophy involve transcriptional changes similar to those of muscle wasting
during systemic diseases. FASEB J. 21, 140–155.
Salghetti, S.E., Caudy, A.A., Chenoweth, J.G., and Tansey, W.P. (2001).
Regulation of transcriptional activation domain function by ubiquitin. Science
Sandri, M., Lin, J., Handschin, C., Yang, W., Arany, Z.P., Lecker, S.H.,
Goldberg, A.L., and Spiegelman, B.M. (2006). PGC-1alpha protects skeletal
muscle from atrophy by suppressing FoxO3 action and atrophy-specific
gene transcription. Proc. Natl. Acad. Sci. USA 103, 16260–16265.
Sandri, M., Sandri, C., Gilbert, A., Skurk, C., Calabria, E., Picard, A., Walsh, K.,
Schiaffino, S., Lecker, S.H., and Goldberg, A.L. (2004). Foxo transcription
muscle atrophy. Cell 117, 399–412.
Sato, Y., Shimizu, M., Mizunoya, W., Wariishi, H., Tatsumi, R., Buchman, V.L.,
and Ikeuchi, Y. (2009). Differential expression of sarcoplasmic and myofibrillar
proteins of rat soleus muscle during denervation atrophy. Biosci. Biotechnol.
Biochem. 73, 1748–1756.
Spencer, J.A., Eliazer, S., Ilaria, R.L., Jr., Richardson, J.A., and Olson, E.N.
(2000). Regulation of microtubule dynamics and myogenic differentiation by
MURF, a striated muscle RING-finger protein. J. Cell Biol. 150, 771–784.
Tang, H., and Goldman, D. (2006). Activity-dependent gene regulation in skel-
etal muscle is mediated by a histone deacetylase (HDAC)-Dach2-myogenin
signal transduction cascade. Proc. Natl. Acad. Sci. USA 103, 16977–16982.
Tang, H., Macpherson, P., Marvin, M., Meadows, E., Klein, W.H., Yang, X.J.,
and Goldman, D. (2008). A histone deacetylase 4/myogenin positive feedback
loop coordinates denervation-dependent gene induction and suppression.
Mol. Biol. Cell 20, 1120–1131.
Tawa, N.E., Jr., Odessey, R., and Goldberg, A.L. (1997). Inhibitors of the
proteasome reduce the accelerated proteolysis in atrophying rat skeletal
muscles. J. Clin. Invest. 100, 197–203.
Vega, R.B., Matsuda, K., Oh, J., Barbosa, A.C., Yang, X., Meadows, E.,
deacetylase 4 controls chondrocyte hypertrophy during skeletogenesis. Cell
Furlow, J.D., and Bodine, S.C. (2008). The glucocorticoid receptor and FOXO1
synergistically activate the skeletal muscle atrophy-associated MuRF1 gene.
Am. J. Physiol. Endocrinol. Metab. 295, E785–E797.
Williams, A.H., Valdez, G., Moresi, V., Qi, X., McAnally, J., Elliott, J.L., Bassel-
Duby, R., Sanes, J.R., and Olson, E.N. (2009). MicroRNA-206 delays ALS
progression and promotes regeneration of neuromuscular synapses in mice.
Science 326, 1549–1554.
Willis, M.S., Rojas, M., Li, L., Selzman, C.H., Tang, R.H., Stansfield, W.E.,
Rodriguez, J.E., Glass, D.J., and Patterson, C. (2009). Muscle ring finger 1
mediates cardiac atrophy in vivo. Am. J. Physiol. Heart Circ. Physiol. 296,
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. 45
EXTENDED EXPERIMENTAL PROCEDURES
Hdac5?/?mice were described previously (Chang et al., 2004). Hdac4 conditional mutant mice were generated by flanking exon 6
with loxP sites, which results in a frame shift mutation in the Hdac4 allele (Potthoff et al., 2007). Transgenic mice harboring genomic
ary et al., 1989; Williams et al., 2009). Among various lines of transgenic mice, three out of five for MuRF1-WT-lacZ and three out of
seven for atrogin-1-WT-lacZ expressed the lacZ transgene in denervated myofibers. Among various lines of transgenic mice
harboring mutant constructs, three out of nine for MuRF1-Emut-lacZ and two out of six for atrogin-1-Emut-lacZ expressed the
lacZ transgene, as seen in denervated myofibers.
Mice harboring the Myogfloxallele mated to CAGGCre-ERTM transgenic mice, which ubiquitously express a conditional Cre-re-
combinase that is activated by intraperitoneal injection of tamoxifen (10 mg/40 g body weight), have been previously described
(Knapp et al., 2006). Tamoxifen injections were performed in two month-old mice. The deleted Myog allele was detected by quan-
titative PCR genotyping from genomic DNA 1 week after tamoxifen injection, using primer sequences described previously (Knapp
et al., 2006). Denervation experiments were performed one month after tamoxifen injection and myogenin deletion was confirmed by
quantitative real time PCR. Littermates injected with tamoxifen, but not expressing the Cre-recombinase, were used as controls. All
mouse studies were approved by the UT Southwestern Institutional Animal Care and Use Committee.
The following plasmids were used: pCMV-Snap25-GFP (provided by Tullio Pozzan, University of Padua, Padua, Italy); EMSV-myo-
genin; pcDNA-E12; pcDNA 3.1 and CMV-lacZ.
Immunoblots were performed from GP muscles as previously described (Vega et al., 2004). Antibodies against HDAC4 (1:500 in
TBST; Sigma) and a-tubulin (1:3,000 in 5% milk TBST; Sigma) were used. ECL Advance Western Blotting Detection Kit (Amersham
Biosciences) was used for signal detection.
Electrophoretic Mobility Shift Assays
Oligonucleotides were synthesized (Integrated DNA Technology) corresponding to the E box sites and mutated binding sites for
generating probes used in the gel mobility shift assays. Unlabeled competitor was 200X of labeled probe. Myogenin and E12
were performed as described (Lu et al., 1999). Oligonucleotides were synthesized as follows (plus strand sequences are shown with
the binding sites in bold and the mutations underlined):
MuRF1 E2: GGAATGCTCAGCTGGTCCCCTC, TCTTGC;
MuRF1 E3: CTGGGGCTCATGTGACAGAGGT, TCGTGC;
atrogin-1 E box: CCCGAGGCCACGTGGCTTTGTT, TCTTGC.
Sybr Green Primers
Dach2 for: 50-ACTGAAAGTGGCTTTGGATAA-30;
Dach2 rev: 50-TTCAGACGCTTTTGCATTGTA-30
Oligonucleotide primers for amplification of E boxes on the MuRF1 promoter are:
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. S1
Oligonucleotide primers for amplification of the E box on the atrogin-1 promoter are:
Oligonucleotide primers for amplification of GAPDH are:
For: 50-ATC CAC GAC GGA CAC ATT GG-30
Rev: 50-TGGTGC TGC CAA GGCTGT GG-30
into the pGL3-Basic reporter (Promega). The MuRF1 genomic fragment was generated by PCR using the following primers:
gene into the pGL3-Basic reporter. The atrogin-1 genomic fragment was generated by PCR using the following primers:
S2 Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc.
Figure S1. Deletion of Myog Allele in Myog?/?Mice, Related to Figure 1
Quantitative PCR genotyping was performed using genomic DNA from Myog?/?and WT mice 1 week after tamoxifen injection. n = 8 for each sample. Data are
represented as mean ± SEM.
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Figure S2. Myog?/?Mice Do Not Show Changes in Fiber Type Composition, Related to Figure 1
(A) Fiber type composition was determined by metachromatic ATPase staining of contralateral and denervated soleus from WT and Myog?/?mice, 14 days after
denervation. Type I fibers stain dark blue. Type II fibers stain light blue. Scale bar = 20 microns.
(B) Type I myosin heavy chain (MHC) immunostaining of contralateral and denervated soleus muscle of WT and Myog?/?mice, 14 days after denervation. Scale
bar = 20 microns.
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Figure S3. Myogenin Binds and Activates MuRF1 and atrogin-1 Transcription In Vitro and In Vivo, Related to Figure 2
(A) Sequence alignment of a fragment of the promoter region of the mouse MuRF1 and atrogin-1 genes from different species shows the conserved upstream
region containing E boxes. Position (0) denotes the transcriptional start site.
(B) Real-time PCR expression of Myogenin, MuRF1 and atrogin-1 in C2C12 cells, 2 and 6 days after switching from growth to differentiation medium. Values are
Cell 143, 35–45, October 1, 2010 ª2010 Elsevier Inc. S5
normalized to transcript expression in growth medium. n = 3 for each time point. Data are represented as mean ± SEM.
(C) Representative ChIP assay performed using C2C12 myotubes shows myogenin binding on the MuRF1 and atrogin-1 genes. Chromatin was immunoprecip-
itated with antibodies against immunogloblulin G (IgG), or myogenin (Myog). A negative control was performed with no antibody (NoAb). Primers flanking the E
boxes on MuRF1 and atrogin-1 promoters were used for amplifying DNA by PCR. GAPDH primers were used as negative control; Input shows similar amount of
chromatin in each sample.
(D) RepresentativeChIPassayperformedusingdenervated TAmuscle at3and 7daysfollowing denervation showsmyogeninbindingontheMuRF1and atrogin-
1 genes. GAPDH primers were used as negative control; Input shows similar amount of chromatin in each sample.
(E) Electrophoretic mobility shift assay shows direct binding of Myogenin/E12 heterodimers to the E box consensus sequences in the MuRF1 (E2 and E3) and
atrogin-1 (E box) promoters. Competitor consists of the E box consensus sequence (w) or a mutated E box (m).
(F) Luciferase assays using extracts of COS cells transfected with luciferase reporter plasmids ligated to WT (MuRF1-Luc) (atrogin-1-Luc), or the mutant (E-mut)
constructs of MuRF1 and atrogin-1 genes and increasing amounts of a myogenin expression plasmid. Empty vector is used as control. Data are represented as
mean ± SEM.
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Figure S4. Absence of HDAC4 Expression in Skeletal Muscle of dKO Mice, Related to Figure 3
Representative immunoblot for HDAC4 in contralateral (Con) and denervated (Den) GP of WT and dKO mice, 7 days after denervation. a-Tubulin antibody is used
as loading control.
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Figure S5. dKO Mice Are Resistant to Muscle Atrophy upon Denervation, Related to Figure 3
Percentageof GP muscle weight of WT and dKO mice 14days after denervation, expressed relative to contralateral muscle. **p < 0.005 versus WT. n= 6 for each
sample. Data are represented as mean ± SEM.
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Figure S6. Ectopic Expression of Myogenin Does Not Induce Muscle Atrophy in WT Mice, Related to Figure 5 Download full-text
(A) Immunostaining for laminin (red) of cross-section of TA WT muscles electroporated with GFP expression plasmid and control plasmid (WT Control) or GFP
plasmid and myogenin (WT + Myogenin). Scale bar = 20 microns.
(B) Morphometric analysis shows that GFP-positive myofibers have no significant (NS) size difference between WT myofibers electroporated with control or my-
ogenin expression plasmid. n = 7 for each sample. Data are represented as mean ± SEM.
(C) Expression of Myogenin, MuRF1 and atrogin-1 in WT TA muscles electroporated with GFP plasmid and a control plasmid (Control) or GFP plasmid and my-
ogenin (Myogenin). Values are normalized to the expression of the control muscles. n = 3 for each sample. Data are represented as mean ± SEM.
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