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doi:10.1152/ajpendo.00646.2007
295:785-797, 2008. First published Jul 8, 2008;Am J Physiol Endocrinol Metab
Holger M. Reichardt, J. David Furlow and Sue C. Bodine
David S. Waddell, Leslie M. Baehr, Jens van den Brandt, Steven A. Johnsen,
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The glucocorticoid receptor and FOXO1 synergistically activate the skeletal
muscle atrophy-associated MuRF1 gene
David S. Waddell,
1
Leslie M. Baehr,
1
Jens van den Brandt,
4
Steven A. Johnsen,
2,3
Holger M. Reichardt,
4
J. David Furlow,
1
* and Sue C. Bodine
1
*
1
Department of Neurobiology, Physiology, and Behavior, University of California, Davis, California;
2
European Molecular
Biology Laboratory, Heidelberg;
3
Department of Molecular Oncology, Go¨ttingen Center for Molecular Biosciences;
and
4
Department of Cellular and Molecular Immunology, University of Go¨ttingen, Go¨ ttingen, Germany
Submitted 4 October 2007; accepted in final form 27 June 2008
Waddell DS, Baehr LM, van den Brandt J, Johnsen SA,
Reichardt HM, Furlow JD, Bodine SC. The glucocorticoid receptor
and FOXO1 synergistically activate the skeletal muscle atrophy-associ-
ated MuRF1 gene. Am J Physiol Endocrinol Metab 295: E785–E797,
2008. First published July 8, 2008; doi:10.1152/ajpendo.00646.2007.—
The muscle specific ubiquitin E3 ligase MuRF1 has been implicated as a
key regulator of muscle atrophy under a variety of conditions, such as
during synthetic glucocorticoid treatment. FOXO class transcription fac-
tors have been proposed as important regulators of MuRF1 expression,
but its regulation by glucocorticoids is not well understood. The MuRF1
promoter contains a near-perfect palindromic glucocorticoid response
element (GRE) 200 base pairs upstream of the transcription start site. The
GRE is highly conserved in the mouse, rat, and human genes along with
a directly adjacent FOXO binding element (FBE). Transient transfection
assays in HepG2 cells and C
2
C
12
myotubes demonstrate that the MuRF1
promoter is responsive to both the dexamethasone (DEX)-activated
glucocorticoid receptor (GR) and FOXO1, whereas coexpression of GR
and FOXO1 leads to a dramatic synergistic increase in reporter gene
activity. Mutation of either the GRE or the FBE significantly impairs
activation of the MuRF1 promoter. Consistent with these findings, DEX-
induced upregulation of MuRF1 is significantly attenuated in mice
expressing a homodimerization-deficient GR despite no effect on the
degree of muscle loss in these mice vs. their wild-type counterparts.
Finally, chromatin immunoprecipitation analysis reveals that both GR
and FOXO1 bind to the endogenous MuRF1 promoter in C
2
C
12
myo-
tubes, and IGF-I inhibition of DEX-induced MuRF1 expression corre-
lates with the loss of FOXO1 binding. These findings present new
insights into the role of the GR and FOXO family of transcription factors
in the transcriptional regulation of the MuRF1 gene, a direct target of the
GR in skeletal muscle.
forkhead transcription factor class O; muscle RING finger 1; gluco-
corticoid receptor
SKELETAL MUSCLE IS A DYNAMIC TISSUE that has the capacity to
continuously regulate its size in response to a variety of
external cues, including mechanical load, neural activity, hor-
mones/growth factors, stress, and nutritional status. In addition,
skeletal muscle serves as the most significant repository for
protein in the body, a source that is tapped to provide a pool of
amino acids for tissue repair and gluconeogenesis under con-
ditions of starvation and other metabolic stresses. Muscle loss
or “atrophy” occurs as the result of a number of disparate
conditions, including aging, immobilization, metabolic dis-
eases, cancer, and neurodegenerative diseases, and as a serious
side effect of therapeutic corticosteroid hormone treatment (15,
27, 32). The recently identified E3 ubiquitin ligase, muscle
RING finger 1 (MuRF1), is proposed to be a key regulator of
the atrophy process given that 1) it is expressed predominantly
in skeletal muscle (3), 2) it is upregulated under a variety of
atrophy conditions (3, 12, 25), and 3) deletion of the gene in
mice results in significant muscle sparing following denerva-
tion (3). Although the full physiological functions of MuRF1
are not yet known, it is often assumed that it functions in some
manner to regulate protein degradation since it is expressed
early in the atrophy process and its peak expression usually
occurs during maximum muscle loss. For example, MuRF1 has
been shown to play a direct role in myosin heavy chain
ubiquitination and degradation during synthetic glucocorticoid
treatment (5). However, MuRF1 may have additional impor-
tant functions in skeletal muscle, such as inhibition of protein
synthesis during starvation conditions (22) as well as regulat-
ing carbohydrate metabolism (16). Despite considerable inter-
est in MuRF1 as a regulator of skeletal muscle mass and
metabolism, there are limited data on the transcriptional regu-
lation of the MuRF1 gene.
Both natural and synthetic glucocorticoid hormones are
potent inducers of skeletal muscle atrophy (14). Glucocorti-
coids exert their physiological actions primarily via a nuclear
pathway to directly affect target gene transcription. Natural
glucocorticoids such as cortisol and corticosterone, as well as
synthetic glucocorticoids such as dexamethasone (DEX) and
prednisolone, exert their biological effects predominantly via
the glucocorticoid receptor (GR) (56). The GR is a member of
the nuclear receptor superfamily and acts as a ligand-dependent
transcription factor. Skeletal muscle expresses significant lev-
els of GR, as do cultured myotubes. In the absence of ligand,
the GR is found largely in the cytoplasm in a large complex
that includes chaperones such as heat shock protein 90. Upon
ligand binding, the GR becomes localized in the cell nucleus
and binds to DNA sequences called glucocorticoid response
elements (GREs). The consensus GRE sequence is AGAA-
CANNNTGTTCT, where two GRs bind to each six-nucleotide
half-site of the palindome as a homodimeric complex (2). The
three-base pair (bp) spacer sequence (NNN) can consist of any
nucleotide combination, although that particular sequence
length is critical. However, the perfect consensus sequence is
rarely found in native glucocorticoid-responsive promoters,
with two or more nucleotide differences often found in either
* J. D. Furlow and S. C. Bodine contributed equally to this study.
Address for reprint requests and other correspondence: J. D. Furlow or S. C.
Bodine, Section of Neurobiology, Physiology & Behavior, Univ. of California
Davis, One Shields Ave., Davis, CA 95616 (e-mail: jdfurlow@ucdavis.edu or
scbodine@ucdavis.edu).
The costs of publication of this article were defrayed in part by the payment
of page charges. The article must therefore be hereby marked “advertisement”
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Am J Physiol Endocrinol Metab 295: E785–E797, 2008.
First published July 8, 2008; doi:10.1152/ajpendo.00646.2007.
0193-1849/08 $8.00 Copyright ©2008 the American Physiological Societyhttp://www.ajpendo.org E785
on October 22, 2008 ajpendo.physiology.orgDownloaded from
or both half-sites (45). In addition, GR may positively or
negatively influence transcription of target genes in a GRE-
and homodimer-independent manner via interaction with pro-
moter-bound STAT5, AP-1, or NF-B transcription factors
(reviewed in Ref. 50). Development of a mouse strain express-
ing a dimerization-deficient GR [GR homodimerization mutu-
ant (GR
dim
)] has allowed the identification of physiological
processes and specific gene expression that are dependent on
the classical GRE binding activity of the GR vs. an indirect
action through other transcription factors (38, 39).
Although the synthetic glucocorticoid DEX induces MuRF1
mRNA accumulation in vivo as well as in cultured myotubes
(3, 48), it is currently unknown whether the gene is directly
regulated by ligand-bound GR in vivo.DEX induction could
be mediated by the increased expression of other transcription
factors and associated coactivators that in turn bind to and
activate the MuRF1 promoter. For example, the class O type
forkhead transcription factors (FOXO), including FOXO1,
FOXO3a, and FOXO4, have recently been implicated as key
regulators of gene expression during skeletal muscle atrophy
(42, 48). FOXO1, FOXO3a, and FOXO4 are all expressed in
skeletal muscle (17), and FOXO1 and FOXO3a mRNA in
particular are upregulated during fasting and DEX treatment
(9, 25). Constitutively active FOXO proteins can activate the
endogenous MuRF1 gene (42, 48); however, there is no infor-
mation on the direct effect of FOXO proteins on the MuRF1
promoter.
Recently, it has been demonstrated that upregulation of
MuRF1 expression following DEX treatment or starvation of
C
2
C
12
cells can be suppressed by IGF-I (41, 42, 48). The
mechanism by which IGF-I is able to suppress MuRF1 tran-
scription is believed to be at least in part via the phosphatidyl-
inositol 3-kinase/Akt pathway. Akt phosphorylates members of
the FOXO class of forkhead transcription factors (40, 60), and
phosphorylated FOXOs are sequestered in the cytoplasm by
14-3-3 proteins, thereby inhibiting transcription of FOXO
target genes. In addition, FOXO proteins lacking Akt phos-
phorylation sites prevent IGF-I inhibition of DEX induction of
MuRF1 (48). Nevertheless, significant questions remain as to
whether nuclear FOXO transcription factors alone are suffi-
cient to activate transcription of the MuRF1 gene. In the
present study, we provide a detailed analysis of the regulatory
elements governing MuRF1 induction following glucocorti-
coid treatment, with particular attention to the role of the GR
and specific FOXO transcription factors.
MATERIALS AND METHODS
Cell culture. HepG2 and C
2
C
12
cells were cultured in DMEM
supplemented with 10% FBS, nonessential amino acids, and antibi-
otics and grown at 37°C in 5% CO
2
.C
2
C
12
cells were cultured in
DMEM, supplemented with 10% FBS, nonessential amino acids, and
antibiotics, and grown at 37°C in 5% CO
2
.C
2
C
12
myoblasts were
differentiated to myotubes by switching confluent cells (usually
24 – 48 h postsplit) to DMEM supplemented with 2% charcoal dex-
tran-treated FBS, nonessential amino acids, and antibiotics and grown
for an additional 48 –96 h at 37°C in 5% CO
2
. All cell culture reagents
except charcoal dextran-treated FBS (Hyclone) were from Invitrogen.
Plasmids. The pSG5-GR construct was a kind gift from Dr. Stoney
Simons, the pcDNA3-FoxO1 construct was provided by Dr. Masahiko
Negishi, pCMV5-cMyc-FoxO3a was provided by Dr. Dominic Accili,
and pcDNA3-FoxO4 was provided by Dr. Karen Arden. The
pcDNA3.1-GRwt construct contains the full-length mouse GR cDNA,
and the pcDNA3.1-GR
dim
construct was derived from pcDNA3.1-
GRwt by overlap PCR introducing the A458T point mutation and a
novel BsrGI restriction site, similar to the strategy used to create the
same point mutation in the GR
dim
mouse targeting vector described
previously (38). The mouse MuRF1 proximal promoter region was
obtained by PCR from bacteria artificial chromosome (BAC) clones
RP23-40E12 (Children’s Hospital Oakland Research Institute). BAC
DNA was isolated from bacterial cultures using the BACMAX DNA
purification kit from Epicentre according to the manufacturer’s pro-
tocol. Primers were designed to amplify approximately ⫺5,000,
⫺2,000, ⫺1,000, and ⫺500 bp from near the transcription start site.
The primer sequences used to generate the MuRF1 promoter frag-
ments are MuRF1-Pro5000-F 5⬘-GGA CAG TGC ATC ATG ACC
CAG-3⬘, MuRF1-Pro2000-F 5⬘-CCA GAA CTA CAC CAG AAA CTC-
3⬘, MuRF1-Pro1000-F 5⬘-GGA GCT GGG AAT ATA GAC TTG-3⬘,
MuRF1-Pro500-F 5⬘-CCT TAG AGC TGT TCA GAA TCC AG-3⬘, and
MuRF-Pro-R 5⬘-CAC TCG GAT CCT CTT TGT CTT C-3⬘. PCR was
performed using TaqPlus Long (Strategene). The resulting PCR prod-
ucts were then subcloned into the pGEMT-EZ vector (Promega) and
sequenced to confirm that the correct amplicon had been obtained.
The promoter fragments were then digested out of the pGEMT-EZ
vector with EcoRI, blunted, and cloned into the SmaI site of pGL3-
Basic vector (Promega), resulting in fusion with the firefly luciferase
reporter gene. The recombinant plasmids were then subjected to
restriction digest analysis and sequenced to confirm correct orienta-
tion. MuRF1 promoter fragments, fused to the secreted alkaline
phosphatase (SEAP) reporter gene, were constructed by digesting the
pGEMT-EZ recombinant plasmids with EcoRI and subcloning the
MuRF1 fragments into the EcoRI site of the pSEAP-Basic vector.
These constructs were then subjected to restriction digestion analysis
and sequenced to confirm correct orientation. Oligonucleotides cor-
responding to the predicted FOXO binding site [FOXO binding
element (FBE)] and the GRE in the MuRF1 promoter, as well as a
string of six diaminofluorescein (Daf-16) binding elements (6X-
DBE), were designed and ordered from Invitrogen. The oligo se-
quences are MuRF1-FBE-F 5⬘-CTA GTT CTT GTT TAC GAC C-3⬘,
MuRF1-FBE-R 5⬘-CTA GGG TCG TAA ACA AGA A-3⬘, MuRF1-
GRE-F 5⬘-CTA GGC TCT GAA CAG TCT GTT CTT GTT-3⬘, MuRF1-
GRE-R 5⬘-CTA GAA CAA GAA CAG ACT GTT CAG AGC-3⬘,
6X-DBE-F 5⬘-CTA GAA GTA AAC AAC TAT GTA AAC AAC TAT
AAG TAA ACA ACT ATG TAA ACA ACT ATA AGT AAA CAA CTA
TGT AAA CAA GAT C-3⬘, and 6X-DBE-R 5⬘-CTA GGA TCT TGT
TTA CAT AGT TGT TTA CTT ATA GTT GTT TAC ATA GTT GTT
TAC TTA TAG TTG TTT ACA TAG TTG TTT ACT T-3⬘. The
complementary oligo sequences were annealed by mixing and heating
to 95°C and slowly cooling to 25°C over a 35-min period using a
thermocycler. The annealed oligos were then ligated into the SpeI site
of the thymidine kinase (TK)-Luc vector (kindly provided by Dr.
Ronald Evans). The resulting recombinant plasmids were then restric-
tion digested to confirm the presence of an insert and sequenced to
determine the number and orientation of the concatemerized oligos.
Site-directed mutagenesis of the FBE and GRE in the pGL3- and
pSEAP-MuRF1-Pro500 constructs was performed essentially as de-
scribed in the site-directed mutagenesis kit protocol from Strategene.
The primers used to mutate the FBE and GRE are: MuRF1Pro-GRE-
Mut-F 5⬘-CCT GGC TCT GGT CAG TCT GAC CTT GTT TAC G-3⬘,
MuRF1Pro-GRE-Mut-R 5⬘-CGT AAA CAA GGT CAG ACT GAC
CAG AGC CAG G-3⬘, MuRF1Pro-FBE-Mut-F 5⬘-CTG TTC TTG
GTG ACG ACC CCC-3⬘, and MuRF1Pro-FBE-Mut-R 5⬘-GGG GGT
CGT CAC CAA GAA CAG-3⬘. The resulting clones were sequenced to
confirm that the correct mutation had been obtained. The mutated
nucleotides are underlined.
Cell culture reporter gene and Northern blot assays. One hundred
twenty-five to 150 ⫻10
3
HepG2 cells/well were plated into 12-well
plates and cultured for 24 h or until an approximate confluency of
30 – 40% was reached. Using FuGene 6 (Roche), 1 g of total
DNA/well was transiently transfected [including 0.250 g/well of the
E786 MuRF1 GENE REGULATION BY GR AND FOXO1
AJP-Endocrinol Metab •VOL 295 •OCTOBER 2008 •www.ajpendo.org
on October 22, 2008 ajpendo.physiology.orgDownloaded from
indicated reporter construct, 0.070 g/well of SV40-Renilla luciferase
(Promega), 0.250 g/well GR expression vector (i.e., pSG5-GR,
pcDNA3.1-GRwt, or pcDNA3.1- GR
dim
), 0.125 g/well FOXO ex-
pression vector, i.e., pcDNA3-FoxO1, pCMV5-cMyc-FoxO3a, or
pcDNA3-FOXO4, and pBluescript as filler DNA] for 12–16 h. Cells
were then treated for 18 –24 h with either vehicle, 1 M DEX (from
a 1-mM stock in ethanol; Sigma), 20 ng/ml IGF-I (Sigma), or both as
indicated. Dual-luciferase reporter assays were performed in HepG2
cells essentially as described previously (7). Briefly, transfected cells
were lysed with 1X passive lysis buffer (Promega), scraped, trans-
ferred to microfuge tubes, and centrifuged at high speed for 5 min to
clear cellular debris, and then 25 l of supernatant from each tube
(corresponding to each well) was transferred to a 96-well plate. Both
working firefly buffer (25 mM glycylglycine, pH 8.0, 5 mM K
2
HPO
4
,
4 mM EGTA, 15 mM MgSO
4
, 4 mM ATP, 1.25 mM DTT, 0.1 mM
CoA, 80 MD-luciferin) and working Renilla buffer (1.1 M NaCl, 2.2
mM Na
2
EDTA, 0.22 M K
2
HPO
4
, pH 5.1, 0.5 mg/ml BSA, 1.5 mM
NaN
3
, and 1.5 M coelenterazine) were prepared fresh prior to each
assay. D-luciferin was purchased from ICN. Five milligrams of lucife-
rin was dissolved in 18 ml of 25 mM glycylglycine (pH 8.0), aliquoted,
and stored at ⫺80°C. Coelenterazine (Promega) was dissolved in EtOH
to a final concentration of 3 mM. All other chemicals were purchased
from Sigma. Following treatments, cells were lysed and supernatants
analyzed for luciferase activity using a Beckman Coulter LD 400 lumi-
nometer programmed to dispense 100 l of working firefly buffer with a
1-s delay and a 10-s integration, followed by injection of 100 lof
working Renilla buffer with a 2-s delay and a 10-s integration. Firefly
luciferase activities were normalized to Renilla luciferase activity to
correct for variations in transfection efficiency.
For C
2
C
12
transfection experiments, 125–150 ⫻10
3
cells/well
were plated into 12-well plates and cultured as described above for
24 h. Using FuGene 6 (Roche), 1 g of total DNA/well was tran-
siently transfected (including 0.250 g/well of the indicated reporter
construct, 0.125 g/well of a TK--galactosidase construct, and
pBluescript as filler DNA) for 12–16 h. Cells were then switched to
DMEM containing 2% charcoal/dextran-treated FBS. Twenty-four
hours following the switch, the cells were treated with vehicle and
either 1 M DEX (from a 1-mM stock in ethanol; Sigma), 20 ng/ml
IGF-I (Sigma), or both as indicated. Medium from each well of the
transfected differentiated C
2
C
12
myotubes was sampled immediately
prior to ligand treatment and then every 24 h following ligand
treatment for 3 days. Fresh ligand was added to the differentiated
C
2
C
12
myotubes every 24 h. Ten to 15 microliters of each collected
medium was then processed and analyzed using the Great EscAPe
SEAP Detection Kit (BD Biosciences), following the instructions of
the manufacturer. At the end of the experiment, the C
2
C
12
myotubes
were lysed and -galactosidase activities determined and used to
correct for variations in SEAP activities resulting from variations in
transfection efficiencies.
For C
2
C
12
Northern blot experiments, cells were differentiated and
treated as above, and total RNA was isolated using using RNeasy
columns (Qiagen) according to the manufacturer’s instructions.
Northern analysis was conducted using random hexamer-labeled
MuRF1 and rpL8 cDNAs essentially as described previously (8).
Animal studies. The generation of GR
dim
mice was described
previously (38). The mice were backcrossed to Balb/c mice for more
than 20 generations. Homozygous mutants and wild-type controls
were obtained by intercrossing heterozygous GR
dim
mice. Animals
were housed individually in ventilated cages under specific pathogen-
free conditions with ad libitum access to food and water and a normal
dark-light cycle. For the 6- and 24-h DEX treatments, 10 mg/kg
water-soluble DEX (Sigma) was given by intraperitoneal injection,
and 50 mg/l DEX was subsequently provided in the drinking water.
For the 3- and 8-day DEX treatments, 50 mg/l water-soluble DEX was
provided in the drinking water and replenished every 2nd day. At each
time point, animals were weighed and euthanized, and spleens as well
as tibialis anterior and gastrocnemius muscles were dissected and
weighed. Muscles were frozen in liquid nitrogen until further process-
ing. Splenocytes were isolated by passing the freshly isolated spleen
through a 40-m nylon mesh, washed in PBS, and subsequently
treated with OptiLyse (Beckman Coulter) to remove the erythrocytes.
Total splenocyte numbers were determined by counting the cells
under the microscope with a Neubauer chamber. Total RNA from
treated wild-type and GR
dim
gastrocnemius muscle was isolated and
analyzed by Northern hybridization using MuRF1, FOXO1 and
FOXO3a, and rpl32 cDNA probes as described above. All animal
experimentation was conducted in accordance with standards of
human animal care and approved by the Lower Saxony state author-
ities (Niedersa¨chisches Landesamt fu¨r Verbraucherschutz und Leb-
ensmittelsicherheit, Braunschweig, Germany).
Chromatin immunoprecipitation. Chromatin immunoprecipitations
(ChIP) were performed as modified by Metivier et al. (30) and Nelson
et al. (35). Chemicals were obtained from Sigma unless stated other-
wise. C
2
C
12
cells were grown to confluence in 100-mm tissue culture
plates in DMEM supplemented with 10% FBS. Cells were then
shifted to 2% FBS and allowed to differentiate into myotubes for 4
days. One plate per time point for the 1-M DEX time course
treatment or four sets of plates in quadruplicate were treated with 1
M DEX, 20 ng/ml R3-IGF-I, or a combination of 1 M DEX and 20
ng/ml R3-IGF-I for 15 or 30 min. Untreated cells were used as a
control. At the indicated times, medium was removed and cells were
incubated with 1% formaldehyde in PBS for 10 min at 37°C. Form-
aldehyde cross-linking was quenched by adding glycine to a final
concentration of 125 mM and incubating for an additional 10 min at
room temperature. Cells were washed twice in PBS, scraped in 10
mM Tris䡠HCl, pH 8.0, 150 mM NaCl, and 1 mM EDTA, and
collected by centrifugation at 3,000 gfor 5 min. Cell pellets were
resuspended in 400 l of 1% SDS, 10 mM EDTA, 50 mM Tris䡠HCl,
pH 8.0, 10 mM -glycerophosphate, 1 mM sodium orthovanadate,
and protease inhibitor cocktail (Roche). Cells were sonicated twice for
30 s each on setting “3” and “pulsed” using a Branson microtip
sonicator. The resulting lysates were then centrifuged at full speed in
an Eppendorf microfuge for 10 min. Supernatants were diluted to 10
ml with ChIP dilution solution (0.01% SDS, 1.1% Triton X-100, 1.2
mM EDTA, 16.7 mM Tris䡠HCl, pH 8.0, 167 mM NaCl, protease
inhibitor cocktail, 10 mM -glycerophosphate, and 1 mM sodium
vanadate). The indicated antibody [anti-HA tag IgG HA probe (Y-11),
anti-FOXO1 IgG (H-128), and anti-glucocorticoid receptor (M-20)
were obtained from Santa Cruz Biotechnology] was added to the
diluted chromatin at a concentration of 1 g/ml, and samples were
incubated overnight at 4°C with tumbling. Fifty microliters of protein
A (50% slurry) preabsorbed with sheared salmon sperm DNA was
added and incubated with tumbling at 4°C for 2 h. Immunoprecipita-
tions were transferred to Bio-Rad minicolumns (prerinsed with 500 l
of ChIP dilution solution), with an additional 500-l dilution solution
to rinse the tubes and ensure complete transfer of beads to the
minicolumns. Columns were then washed twice each with 1 ml TSEI
(0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris䡠HCl, pH
8.0, 150 mM NaCl), TSEII (0.1% SDS, 1% Triton X-100, 2 mM
EDTA, 20 mM Tris䡠HCl, pH 8.0, 500 mM NaCl), and TSEIII (1 mM
EDTA, 10 mM Tris䡠HCl, pH 8.0, 1% NP-40, 1% sodium deoxy-
cholate, 0.25 M LiCl) followed by three washes with Wash Buffer IV
[10 mM Tris (pH 8.0), 1 mM EDTA]. Washed beads were transferred
to new tubes with 400 l of Wash Buffer IV, adding an additional 500
l Wash Buffer IV to the columns to ensure that all beads were
transferred. The beads were then pelleted by centrifugation at 3,000 g
for 2 min. After the supernatant was removed by careful aspiration,
100 l of 10% Chelex-100 (Bio-Rad) was added to the beads,
followed by vortexing and incubation at 95°C for 10 min to reverse
the cross-linking. Two microliters of 20 U/ml proteinase K solution
(Invitrogen) was added, followed by vortexing and incubation at 55°C
for 30 min and heat inactivation at 95°C for 10 min. After centrifu-
gation at full speed in an Eppendorf microfuge for 2 min, the
supernatant was transferred to a new tube and combined with a second
E787MuRF1 GENE REGULATION BY GR AND FOXO1
AJP-Endocrinol Metab •VOL 295 •OCTOBER 2008 •www.ajpendo.org
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extraction of the Chelex beads with 100 l of water. Input (nonim-
munoprecipitated chromatin) was prepared by adding 100 l of 10%
Chelex directly to 100 l of the diluted chromatin extract and
processed as above. Quantitative PCR of input and immunoprecipi-
tated chromatin samples was performed using 2 l of DNA as
template and PerkinElmer 2X SYBR Green master mix on an ABI
7700. Cycling conditions were one cycle at 95°C for 10 min, followed
by 40 cycles at 95°C for 10 s and 60°C for 60 s. Primers flanking the
putative mouse MuRF1 GRE/FBE region were forward 5⬘-
TATCTGGCTCTCCCCTGAAC-3⬘and reverse 5⬘-CCTCAAAGATTTG-
GCCCTCT-3⬘. Values for each time point and hormone treatment were
normalized to input values. For agarose gel analysis of PCR products,
reactions were stopped at 30 cycles, and samples from each treatment
group were pooled and run on a 2% agarose gel containing ethidium
bromide.
RESULTS
The MuRF1 promoter contains a conserved near-perfect
palindromic glucocorticoid response element. Since MuRF1 is
known to be induced by synthetic glucocorticoids, as well as in
several catabolic conditions associated with elevated endoge-
nous glucocorticoids, we sought to identify the key glucocor-
ticoid-responsive elements in the gene. We began by isolating
BAC clones encompassing the entire mouse MuRF1 transcrip-
tion unit and amplifying 5,000 bp upstream of the transcription
start site of the gene. Human, rat, and mouse promoter se-
quences were aligned to detect conserved sequences that may
be functionally relevant for MuRF1 expression. The MuRF1
(Fig. 1A) promoter shows a high degree of homology across
Fig. 1. Schematic of the muscle RING finger 1
(MuRF1) promoter and sequence alignment of the
proximal regulatory regions. A: promoter sequences
from mouse, rat, and human MuRF1 [5,000 base
pairs upstream of the transcription start site (⫹1)
through the first exon] were downloaded from the
Ensembl database (www.ensembl.org) and aligned
using the ClustalW algorithm. Approximate positions
of potential transcription factor binding sites are in-
dicated in the schematics of the MuRF1 promoter at
top and/or boxed in the alignment at bottom. The
forkhead transcription factor class O (FOXO) fork-
head binding site (G/A)TAAA(T/C)AA (black
ovals), glucocorticoid response element (GRE) (A/
T)GAACANNNTGTTC(A/T) (hatched rectangle),
CCAAT/enhancer-binding protein (C/EBP) TT(G/
T)NGNAA ({), NF-B consensus binding sequence
GGG(G/A)N(C/T)(C/T)(C/T)CC (gray hexagons),
and muscle-specific E-box CANGTG (MyoD, etc.);
N represents any nucleotide. The sequence align-
ments for ⬃400 base pairs upstream and 26 base
pairs downstream of the rat, mouse, and human
MuRF1 promoters are shown below the correspond-
ing promoter schematics. Identical sequences for the
indicated regions are highlighted in black. Arrow
indicates transcription start site. B: comparison of a
consensus GRE and perfect palindrome GRE and
putative GREs from the mouse (mMuRF1), rat
(rMuRF1), and human MuRF1 (hMuRF1) promoters.
Arrows indicate half-sites.
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species for about 400 bp upstream of the transcription start site,
and there are additional sporadic pockets of homology far
upstream of the proximal promoter, 3⬘of the transcription unit,
and within some introns (not shown). We searched for poten-
tial transcription factor binding sites by scanning the promoters
for published and well-established consensus sequences. Pre-
vious reports identified and characterized consensus Class O
forkhead (FOXO) binding sites in the promoter of the muscle
atrophy-related muscle atrophy F-box gene (42), and we found
multiple potential AT-rich FBEs in the MuRF1 promoter as
well, including one overlapping the consensus TATA box (Fig.
1A). Importantly, the MuRF1 promoter has a near-perfect
consensus GRE ⬃200 bp upstream of the transcription start
site, supporting our hypothesis that this gene could be a direct
target of the ligand-activated GR. The potential GREs from the
human, mouse, and rat MuRF1 genes differ only by a single
nucleotide in the 3-bp spacer region, whose exact sequence is
not critical for GR binding (Fig. 1B). Interestingly, a consensus
FBE was found directly adjacent to the putative MuRF1 GRE,
raising the possibility that FOXO and the GR function together
to regulate the MuRF1 promoter. In addition to FOXO and GR
binding sites, we detected a cluster of potential NF-B re-
sponse elements in the MuRF1 promoter, consistent with the
upregulation of this promoter by proinflammatory cytokines
and upon activation of NF-B (4). Several C/EBP sites were
also detected in the MuRF1 promoter, which is of interest
because C/EBPand -␦are upregulated by DEX in skeletal
muscle cells (57, 58). Finally, numerous conserved consensus
muscle-specific E-box sites were found in the promoter, as
expected given its highly selective cardiac and skeletal muscle
expression pattern.
The MuRF1 promoter is a direct target of activated GRs.
Since the mere presence of consensus transcription factor
binding sites does not guarantee their functional importance,
MuRF1 promoter fragments, or the predicted GRE and FBE in
a minimal heterologous promoter, were placed in front of the
firefly luciferase reporter and transiently transfected into
HepG2 cells along with expression vectors for the murine GR
and/or FOXO family members. For these initial studies,
HepG2 cells were chosen for their ease of transfection and their
previous use in transfection studies of gluconeogenic gene
promoters that are also regulated by both GR and FOXO
family members (23, 24, 52). Consistent with the presence of
the consensus GRE within 500 bp of the transcription start site,
DEX was able to induce all MuRF1 promoter constructs
containing between 500 and 5,000 bp (Fig. 2A). Indeed, the
isolated MuRF1 GRE is sufficient to support potent DEX-
induced transcription either alone (14-fold) or as a multimer
(19- to 45-fold) (Fig. 2B).
FOXO transcription factors differentially activate the
MuRF1 promoter. Next, we tested whether the native MuRF1
promoter would respond to cotransfected FOXO1, FOXO3a, or
FOXO4 expression vectors, since these proteins are candidates
for intermediate DEX-inducible regulatory factors. As shown
in Fig. 3A, FOXO factors only marginally induce the proximal
MuRF1 promoter (2- to 3-fold). As with the GRE, we tested
whether the isolated FOXO binding site (FBE) would respond
to cotransfected FOXO1, FOXO3a, or FOXO4 expression
vectors; in this context, only FOXO3a and FOXO4 induced
transcription of the reporter containing a single FBE (Fig. 3B).
However, FOXO1 is able to activate multimerized DBE or the
MuRF1 FBE, suggesting that this FOXO family member is
most active as a multimeric complex rather than a monomer
(Fig. 3C).
The MuRF1 promoter is synergistically activated by FOXO1
and GR. Remarkably, however, FOXO1 showed strong syner-
gistic activation with the GR on the 500-bp MuRF1 promoter
(Fig. 4A). This potent synergy was consistently observed (20-
to 40-fold) over multiple experiments and on all promoter
fragments tested ⱕ5,000 bp from the transcription start site.
FOXO3a and FOXO4 did not synergize with the GR; indeed,
expression of FOXO3a somewhat inhibits GR activation of the
MuRF1 promoter. The GR-FOXO1 synergy is also reflected in
dose-response experiments, where this combination supports
strong activation of the MuRF1 promoter with as little as 1 nM
DEX in the culture medium (Fig. 4B). Both the GRE and the
adjacent FBE play an important role in the observed GR-
FOXO synergy. DEX induction of the 500-bp MuRF1 pro-
moter is nearly abolished when the GRE is mutated and is
completely abolished when the GRE is intact but the adjacent
FOXO site is mutated (Fig. 4, Cand D).
DEX induction of MuRF1 expression is inhibited in GR
dim
mice that express a dimerization-deficient GR. The functional
significance of the near-perfect GRE in the MuRF1 promoter
was tested in vivo through the use of the GR
dim
mouse strain
that expresses a GR with a point mutation in the DNA binding
domain that prevents binding of the GR to classical palin-
Fig. 2. Dexamethasone (DEX)-activated glucocorticoid receptor (GR) induces
the MuRF1 proximal promoter. A: HepG2 cells were transfected with lucif-
erase reporter constructs (Luc) containing varying lengths of the MuRF1
(pGL3-MuRF1) promoter, an SV40-Renilla luciferase reporter construct, and
a mouse GR expression vector (pSG5-GR). Cells were treated with or without
1M DEX for 24 h. Luciferase activities in cell extracts were normalized to
Renilla luciferase activity to control for transfection efficiency. Numbers on
the y-axis indicate distance from the transcription start site fused to the reporter
pGL3-Basic, a promoterless LUC vector. B: the isolated MuRF1 GRE supports
DEX-mediated transcriptional activation of a heterologous promoter. The DEX
regulation of the thymidine kinase (TK) promoter was tested with 0 (control),
1 (1XGRE), 3 (3XGRE), and 4 (4XGRE) inserted GREs. The arrows below the
indicated GRE constructs labeled on the y-axis depict the orientation of each
individual GRE oligonucleotide. In Aand B, fold induction was obtained by
dividing values from DEX-treated samples by the mean of values from
matched untreated samples. Each condition was done in triplicate, and error
bars reflect SD.
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dromic GREs (38). This mutant GR still retains the ability to be
tethered to target genes via protein-protein interactions such as
the NF-B or AP-1 transcription factor complexes. Both wild-
type and GR
dim
mice were treated with water-soluble DEX
either by injection (6- and 24-h time points) or in the drinking
water (24-h, 3-day, and 8-day time points). The experiment
was terminated at 8 days of treatment due to the severe weight
loss observed in the wild-type mice, which was not observed in
the GR
dim
mice (P⬍0.001). As shown in Fig. 5A, despite the
inhibited overall body weight decline in GR
dim
vs. wild-type
mice in response to DEX, the degree of atrophy induced in the
tibilialis anterior and gastrocnemius muscles was not statisti-
cally different in the two strains. By contrast, DEX-induced
splenocyte apoptosis was strongly inhibited in GR
dim
mice
(P⬍0.001).
We next examined MuRF1 expression as well as FOXO1
and FOXO3a expression in GR
dim
vs. GR
⫹/⫹
mice (Fig. 5B).
MuRF1 expression is clearly upregulated by 6 h after DEX
injection and continues to increase ⱕ3 days with continuous
DEX treatment in wild-type mice. MuRF1 induction is
strongly inhibited in GR
dim
mice, which is particularly appar-
ent at the earliest time points. After 8 days of DEX treatment,
however, MuRF1 expression declines in wild-type mice until it
is essentially equal to its induced expression in GR
dim
animals.
In addition, FOXO1 and FOXO3a are also rapidly induced by
DEX in wild-type mice, in agreement with a previous study
(9), and expression is reduced in GR
dim
mice at all time points
tested. We then tested the ability of the homodimerization
mutant GR expressed in GR
dim
mice to upregulate the proximal
MuRF1 promoter, again using transient transfection assays
(Fig. 5C). The homodimerization mutant GR lost the ability to
induce the MuRF1 promoter on its own, as expected, and
showed a strongly reduced, but not completely abolished,
ability to augment FOXO1 induction of the promoter. These
transfection results may at least partially explain the residual
DEX-induced expression of MuRF1 in GR
dim
mice.
The GR and FOXO1 bind to the endogenous MuRF1 pro-
moter in C
2
C
12
myotubes. Given the strong dependence on GR
homodimerization in vivo for full MuRF1 gene induction, we
next sought to determine the role of the GRE and FOXO1 sites
in cultured skeletal myotubes. As we observed in HepG2 cells,
the 500-bp MuRF1 promoter is also DEX inducible in differ-
entiated C
2
C
12
mouse skeletal myotubes (Fig. 6A). DEX in-
duction of the promoter is nearly abolished when the GRE is
mutated, and the adjacent FOXO site is intact and completely
abolished when the GRE is intact but the adjacent FOXO site
is mutated (Fig. 6B). Furthermore, addition of 20 ng/ml IGF-I
strongly inhibits DEX induction of the endogenous MuRF1
gene, as has been reported previously (Fig. 6C) (41, 48), and
IGF-I likewise inhibits DEX induction of the MuRF1 promoter
(Fig. 6D). Therefore, the 500-bp proximal promoter of MuRF1
contains the critical elements necessary to respond positively to
glucocorticoids (via synergy with a forkhead site) and nega-
Fig. 3. FOXO transcription factor induction of the MuRF1 promoter. A: HepG2 cells were transfected with 500-base pair MuRF1-promoter-LUC constructs,
SV40-Renilla luciferase reporter construct, and expression vectors for murine FoxO1, FoxO3A, or FoxO4 (pcDNA3-FoxO1, pcDNA3-FoxO3A, or pcDNA3-
FoxO4) or pcDNA3 alone (dash). Luciferase activities in cell extracts were normalized to Renilla luciferase activity to control for variations in transfection
efficiency. Each point was done in triplicate; error bars reflect SD. B: the isolated MuRF1 FOXO binding element (FBE) supports FOXO3a- and FOXO4- but
not FOXO1-mediated transcriptional activation of a heterologous promoter. FOXO regulation of the TK promoter was tested by inserting a single FBE and
cotransfecting HepG2 cells with the control pcDNA3 vector alone (dash) or each FOXO expression vector as in A.C: concatamerized FOXO binding elements
from the MuRF1 promoter (FBE-4X) and Daf-16 binding elements (DBE-6X) support transcriptional activation of the TK promoter by FOXO1. Open bars,
normalized luciferase values from the cells transfected with the TK-Luc-4XFBE and TK-Luc-6XDBE in the absence of exogenous FOXO1 expression; black
bars, normalized luciferase values from the TK-Luc-4XFBE- and TK-Luc-6XDBE-transfected cells in the presence of exogenous FOXO1 expression. Data were
processed as in Aand B.
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tively to IGF-I in a manner identical to the endogenous gene in
differentiated myotubes.
To determine whether FOXO1 and GR can directly occupy
the MuRF1 promoter in vivo, we established a modified ChIP
protocol suitable for use with differentiated C
2
C
12
myotubes.
C
2
C
12
myoblasts were plated on 100-mm dishes and cultured
until confluent and then differentiated for 4 days in reduced
serum medium. Myotubes were then treated with vehicle, 1
M DEX, 20 ng/ml IGF-I, or 1 M DEX plus 20 ng/ml IGF-I
for the indicated times. Sonicated chromatin from fixed cells
was analyzed with control IgG, an anti-FOXO1 antibody, or
two anti-GR antibodies. Both GR and FOXO1 are present on
the endogenous MuRF1 promoter in untreated C
2
C
12
myotubes
(Fig. 7A), and binding of each factor is enhanced by DEX
treatment as soon as 15 min after ligand addition (Fig. 7, Band
C), especially the GR. GR binding to the MuRF1 promoter is
unaffected by IGF-I (Fig. 7B), although the DEX induction of
the endogenous gene or the transfected MuRF1 promoter is
strongly inhibited. However, IGF-I addition rapidly induces the
loss of FOXO1 from the MuRF1 promoter in the presence or
absence of DEX (Fig. 7C). We also attempted to detect
FOXO3a binding to the MuRF1 promoter by ChIP as well;
only one of two commercially available antibodies we tested
(Upstate vs. Santa Cruz) gave a weak positive signal, and the
binding pattern was highly similar to FOXO1 (not shown).
DISCUSSION
Although it is known that MuRF1 expression increases
under numerous atrophy-inducing conditions (11), the mecha-
nism of transcriptional regulation of this gene is poorly under-
stood. The present study provides new insights into how the
GR and the FOXO family of transcription factors regulate the
transcription of the MuRF1 gene. Specifically, we demonstrate
here that 1) the MuRF1 proximal promoter is directly activated
by glucocorticoids via a conserved GRE that depends strongly
on GR homodimerization for full activation, at least in the
earliest time points after DEX treatment, 2) FOXO1 and GR
synergistically and specifically induce the MuRF1 proximal
promoter, and 3) IGF-I inhibition of DEX-induced MuRF1
expression correlates with the loss of FOXO1 binding to the
endogenous MuRF1 promoter. Importantly, these data also
reveal that not all FOXO family members equally activate the
FOXO binding motif in the MuRF1 promoter.
The presence of an essentially perfect palindromic GRE in
the MuRF1 proximal promoter likely explains the strong in-
duction of the gene by exogenous synthetic glucocorticoids (3,
48) and how the gene is induced under a variety of catabolic
conditions associated with increased endogenous corticoste-
roid levels (25, 53). Each half-site of the MuRF1 GRE is
perfectly conserved between the human, rat, and mouse genes,
with the only difference found in the spacer region that,
nevertheless, still retains the three-base pair length necessary
for GR homodimer binding (43). It is surprisingly rare to find
such a perfect GRE in an endogenous glucocorticoid target
gene promoter, although the specific sequences of the diver-
gent half-sites found in most direct GR target genes are
strongly conserved across species (45, 46). Most GREs, such
as the pair found in the well-studied phosphoenolpyruvate
Fig. 4. DEX-activated GR synergizes with FOXO1 to potently induce the MuRF1 promoter. A: HepG2 cells were transfected with the pGL3-MuRF1 promoter
(⫺500) reporter construct, SV40-Renilla luciferase reporter construct, pSG5-GR, and/or pcDNA3-FOXO1, pcDNA3-FoxO3A, or pcDNA3-FOXO4. Cells were
treated with or without 1 M DEX for 24 h. Firefly luciferase activities in cell extracts were normalized to Renilla luciferase activity to control for variations
in transfection efficiency. Fold induction was obtained by dividing values from DEX-treated samples by the mean of values from matched untreated samples.
Each point was done in triplicate, and errors reflect SD. B: HepG2 cells were transfected with pGL3-MuRF1 promoter (⫺500), SV40-Renilla, and pBluescript
as filler DNA (control) and either GR (GR) or GR and FoxO1 (GR ⫹FoxO1) expression vectors. Twenty-four hours posttransfection, cells were treated with
the indicated DEX concentration and incubated overnight. Each point was done in triplicate. Cand D: mutation of either the GRE or the FBE is sufficient to
abolish DEX-induced MuRF1 promoter activity. HepG2 cells were transfected with either the wild-type MuRF1 promoter construct (⫺500) or MuRF1 promoter
constructs that have either the GRE mutated (GRE-Mut) or the FBE mutated (FBE-Mut) as shown in Cin combination with expression vectors for FoxO1 and/or
GR as indicated. The cells where then treated and assayed for luciferase activity as in A.
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carboxykinase (PEPCK) gene in the liver (18) or the first-
described GREs in the mouse mammary tumor virus long-
terminal repeat (37), have multiple base pair changes compared
with the perfect palindrome consensus GRE that is present in
the MuRF1 promoter. Previous studies on the estrogen-regu-
lated family of vitellogenin genes in the frog liver demonstrate
that a single perfect estrogen response element supports essen-
tially the same fold induction by estradiol as a pair of imperfect
palindromic elements (29). The physiological relevance of
having such a well-conserved consensus GRE in the MuRF1
promoter is an interesting and important question.
Analysis of MuRF1 induction by DEX in GR
dim
mice
revealed a critical role for GR homodimerization, which is
most evident at the earliest time points examined in these
experiments. Interestingly, despite the clear overall reduction
in MuRF1 expression, as well as FOXO1 and FOXO3a, the
degree of muscle atrophy in GR
dim
mice after 8 days of DEX
treatment was exactly the same in the mutant as in wild-type
mice. Consequently, the residual level of MuRF1 expression in
GR
dim
mice may well be enough to support full DEX-induced
atrophy. The rapid rise and gradual decline of MuRF1 expres-
sion we observed in DEX-treated Balb/c mice is in agreement
with experiments in rats dosed with the synthetic glucocorti-
coid prednisolone (1). Concerning the mechanism of residual
MuRF1 induction in GR
dim
mice, it is conspicuous that co-
transfection of FOXO1 with the homodimerization mutant GR
0
5
10
15
20
25
30
35
40
Control GR FOXO1 GR + GRdim GRdim +
FOXO1 FOXO1
Fold induction (+DEX/-DEX)
35
40
45
50
55
Tibialis anterior weight (mg)
0GRdim
1
2
4
8
16
32
64
128
256
GRdim
70
90
110
Gastrocnemius weight (mg)
0GRdim
-20
-15
-10
-5
0
5
Reduction in body weight (%)
GR+/+ GRdim
GR+/+
GR+/+ GR+/+
-DEX
+DEX
-DEX
+DEX
-DEX
+DEX
A
B
C
GR+/+ GRdim
Untreated 3d DEX 8d DEX
MuRF1
FOXO1
FOXO3a
rpL32
GR+/+ GRdim GR+/+ GRdim
GR+/+ GRdim
Untreated 6 hr DEX 24 hr DEX
GR+/+ GRdim GR+/+ GRdim
MuRF1
FOXO1
FOXO3a
rpL32
Splenocytenumber (x106)
Fig. 5. DEX-induced MuRF1 expression is inhibited in mutant GR homodimerization mutant (GR
dim
) mice vs. wild-type GR
⫹/⫹
mice. A:GR
⫹/⫹
and GR
dim
mice were treated with DEX in their drinking water for 8 days, weighed (top left), and killed for determination of splenocyte number (top right) as well as
gastrocnemius (bottom left) and tiblialis anterior (bottom right) weights. Open bars, vehicle treated; black bars, DEX treated. GR
⫹/⫹
,n⫽8; GR
dim
,n⫽7.
B: Northern analysis of MuRF1, FOXO1, and FOXO3a expression in gasctocnemius muscle from DEX-treated GR
⫹/⫹
and GR
dim
mice. B,top: control, 6-h-,
and 24-h-treated mice. B,bottom: control, 3-day-, and 8-day-treated mice. Only results from 4 of the 8-day-treated GR
⫹/⫹
and GR
dim
mice are presented. rpL32
expression is shown as a loading control below each set of Northerns. Bars represent means ⫾SE. C: effect of wild-type GR and the homodimerization mutant
GR on DEX induction of the MuRF1 promoter in a transient transfection assay. HepG2 cells were transfected with the pGL3-MuRF1 promoter (⫺500) reporter
construct, SV40-Renilla luciferase reporter construct (control), pcDNA3.1-GRwt (GR) or pcDNA3.1-GR
dim
(GR
dim
), and/or pcDNA3-FoxO1 (FOXO1). Cells
were treated with or without 1 M DEX for 24 h. Firefly luciferase activities in cell extracts were normalized to Renilla luciferase activity to control for variations
in transfection efficiency. Fold induction was obtained by dividing values from DEX-treated samples by the mean of values from matched untreated samples.
Each point was done in triplicate, and errors reflect SD.
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supports at least modest induction of the MuRF1 promoter in
transient transfection assays, whereas the induction without
FOXO1 is completely lost (Fig. 5). This is in line with the
previous observation that the GR
dim
receptor on its own fails to
bind to a palindromic GRE. Nevertheless, it is conceivable that
FOXO1, when present together, still tethers the homodimer-
ization mutant GR to the composite GRE-FBE motif, since
protein-protein interactions with other transcription factors are
not compromised by the GR
dim
mutation. Regardless, it will be
important to identify DEX-responsive genes that are less af-
fected than MuRF1 and FOXOs in GR
dim
mice that may
ultimately be more important for the loss of muscle mass in
response to DEX. For example, activation of the N-end rule
ubiquitin pathway was recently implicated in MyoD degrada-
tion in response to DEX in C
2
C
12
cells (49). In addition,
transgenic overexpression of MuRF1 was not sufficient to
induce atrophy but led to altered levels of enzymes involved in
carbohydrate metabolism (16). Thus, consideration of these
and other studies, and our present results with GR
dim
mice,
may force a reevaluation of the role of MuRF1 (and possibly
even FOXO1 and FOXO3a) in the reduction of muscle mass in
catabolic states, although the protein certainly may still play a
significant role in structural protein degradation in denervation-
induced atrophy. Further analysis of the GR
dim
model may
allow for a genetic dissection of the induction of atrophy vs.
metabolic effects of glucocorticoids in this important GR target
tissue.
Several recent papers have implicated FOXO family tran-
scription factors as being important for the upregulation of
gene expression during skeletal muscle atrophy, including after
DEX treatment or nutrient starvation in C
2
C
12
cells. For
example, Stitt et al. (48) reported that activated FOXO1 is
necessary but not sufficient to activate the MuRF1 gene in
cultured myotubes; however, the effects of FOXO3a or
Fig. 6. DEX induces the MuRF1 promoter in differentiated C
2
C
12
myotubes.A:C
2
C
12
myoblasts were transfected with a reporter construct containing 500 base
pairs of the MuRF1 promoter fused to the secreted alkaline phosphatase (SEAP) gene. The myoblasts were then differentiated by switching to low-serum medium
followed by treatment with 1 M DEX over a period of 3 days. The medium was sampled every 24 h to measure for SEAP activity. Conditions were done in
triplicate and SEAP numbers normalized with -galactosidase to correct for variations in transfection efficiency. Each point was done in triplicate, and errors
reflect SD. B: mutation of either the GRE or the FBE is sufficient to abolish DEX-induced MuRF1 promoter activity in C
2
C
12
cells. C
2
C
12
myoblasts were
transfected with either the wild-type MuRF1 promoter construct (⫺500) or MuRF1 promoter constructs that have either GRE-Mut or FBE-Mut SEAP constructs,
differentiated, and treated with 1 M DEX over a period of 3 days. The medium was sampled every 24 h to measure for SEAP activity as in A. Each time point
was done in triplicate and normalized with -galactosidase to correct for variations in transfection efficiency. C: Northern blot analysis of C
2
C
12
cells
differentiated for 48 h and then treated with DEX (10 M), IGF-I (20 ng/ml), or DEX (10 M) ⫹IGF-I (20 ng/ml) for 24, 48, and 72 h. Ligand was refreshed
every 24 h. D: IGF-I potently inhibits DEX-induced activation of the MuRF1 promoter. C
2
C
12
myotubes were treated for 24 h with either 1 M DEX, 20 ng/ml
IGF-I, or 1 M DEX and 20 ng/ml IGF-I followed by sampling of the medium for SEAP activity, as described in A. Fold induction was obtained by dividing
values from DEX-treated samples by the mean of values from matched untreated samples. Each point was done in triplicate, and errors reflect SD.
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FOXO4 were not reported. Interestingly, selective transgenic
overexpression of FOXO1 in skeletal muscle produced mice
with significantly smaller muscles than wild-type litter mates
without apparent upregulation of MuRF1 (19). The lack of
MuRF1 induction in these instances is not surprising given our
present findings. Assuming that FOXO1 was the only tran-
scription factor overexpressed in these transgenic mice, and
that circulating corticosteroid levels are normal, then MuRF1
would not necessarily be upregulated. The reduction in muscle
mass observed in these mice may be due to the activation of
other atrophy-inducing pathways. For example, overexpression
of FOXO1 leads to an upregulation of 4E-BP1 and inhibition
of mTOR signaling, resulting in a decrease in protein
synthesis (47).
There is important precedence for the collaborative role of
FOXO1 on GR target genes from studies of inversely regulated
insulin and glucocorticoid-responsive genes in the liver. The
IGF-binding protein-1 (10) and glucose-6-phosphatase pro-
moters contain GREs with directly adjacent and even overlap-
ping FOXO binding sites, and the PEPCK (13) and pyruvate
dehydrogenase kinase-4 (24) genes have FOXO sites in rela-
tively close proximity to their GREs. In each case, integrity of
the FOXO site is important for maximal glucocorticoid respon-
siveness.
In addition, like MuRF1, the ability of FOXO1 and FOXO3a
to have differential effects on a single promoter has recently
been demonstrated by Onuma et al. (36) for the glucose-6-
phosphatase promoter. It is intriguing that key genes involved
in gluconeogenesis have clustered functional GREs and FOXO
binding sites in critical glucocorticoid and insulin-sensitive
enhancers reminiscent of our findings with the muscle-specific
MuRF1 gene promoter. It is tempting to speculate that if
indeed one role of MuRF1, as an E3 ubiquitin ligase, is to
target specific skeletal muscle proteins for degradation, then a
gene involved in providing a source of amino acid substrates
for the liver to use to make glucose de novo might logically be
under similar regulatory control as important gluconeogenic
enzymes. The common regulatory strategy of MuRF1 expres-
sion with liver gluconeogenic enzymes also supports the notion
of a potential prominent role for MuRF1 in skeletal muscle
glucose metabolism.
The role of FOXO1 as a key component of a composite
glucocorticoid-responsive unit in MuRF1 gene regulation is
further emphasized by our ChIP data. IGF-I not only virtually
eliminates DEX upregulation of the 500-bp MuRF1 promoter
and the endogenous MuRF1 gene (see Fig. 7) but also rapidly
clears the MuRF1 promoter of FOXO1 binding without affect-
ing GR binding. On the PEPCK enhancer in rat liver H4IIE
cells, insulin treatment causes a rapid loss of FOXO binding in
control and DEX-treated cells, with significant reduction al-
ready apparent by as little as 3 min after insulin addition (13).
In this case, GR binding was reduced to ⬃40% compared with
the DEX only-induced level after 30 min of treatment, which
we did not observe upon IGF-I addition to C
2
C
12
cells. Nev-
ertheless, the most impressive change in promoter occupancy
examined in both our experiments and the PEPCK gene ex-
periments was decreased FOXO1 binding. In addition to inhi-
bition of FOXO1 (and to a lesser extent GR) binding to the
PEPCK promoter in response to insulin treatment, dramatic
changes in histone posttranslational modifications that paral-
leled decreased transcription were observed (13). Whether
Fig. 7. GR and FOXO1 associate directly with the MuRF1 promoter in C
2
C
12
myotubes. A:C
2
C
12
myotubes were treated with 1 M DEX for 0 or 60 min
(1 10-cm plate/time point), and cross-linked chromatin was immunoprecipi-
tated with normal rabbit IgG, an anti-FOXO1 antibody, or one of two anti-GR
antibodies (P20 or M20). After reversal of cross-links, immunoprecipitated
MuRF1 promoter fragments were detected by PCR using primers flanking the
predicted GRE and FOXO sites in the mouse MuRF1 promoter, followed by
agarose gel electrophoresis. PCRs of a no-template control (NTC) and input
DNAs are shown at left.Band C:C
2
C
12
myotubes were treated with vehicle
(open bars), 1 M DEX, 20 ng/ml R3-IGF-I, or 1 M DEX ⫹20 ng/ml
R3-IGF-I for 15 min (black bars) or 30 min (gray bars) before harvest. Four
plates for each treatment were processed in parallel as in A. Values (fold
enrichment) are expressed as the mean MuRF1 promoter copies immunopre-
cipitated with either the anti-GR antibody (B) or anti-FOXO1 antibody (C)
divided by that immunoprecipitated by a nonspecific antibody (anti-HA), as
determined by quantitative PCR using the same primer pairs as in A. Error bars
reflect SE.
E794 MuRF1 GENE REGULATION BY GR AND FOXO1
AJP-Endocrinol Metab •VOL 295 •OCTOBER 2008 •www.ajpendo.org
on October 22, 2008 ajpendo.physiology.orgDownloaded from
these changes occur on the MuRF1 promoter in response to
DEX and/or IGF-I treatment remains an interesting question,
as is whether removal of FOXO1 is the key event triggering the
reversal of transcriptional activation and associated chromatin
modifications induced by DEX.
The exact nature of the synergy between the GR and FOXO1
is clearly an important area for further investigation because it
is also not well understood for liver gluconeogenic genes.
FOXO1 has been demonstrated to physically interact in solu-
tion with several steroid hormone receptors, such as the estro-
gen, progesterone, and androgen receptors (21, 26, 44, 59), but
not, to date, with the glucocorticoid receptor. The functional
significance of this interaction (either repressing or enhancing
receptor activity) varies greatly depending on the cell type and
target gene promoter context. For example, FOXO1 synergis-
tically activates the IGF-binding protein-1 promoter with
ligand-activated progesterone receptors in endometrial adeno-
carcinoma cells but not in endometrial fibroblasts (20).
FOXO1, FOXO3a, and FOXO4 all contain an LXXLL motif
that can interact with a ligand-induced pocket on the hormone
binding domain of steroid receptors (59). Since FOXO1 bind-
ing to the MuRF1 promoter is (at least modestly) enhanced by
DEX treatment of C
2
C
12
cells along with the strong recruit-
ment of GR (Fig. 7), a situation also observed on the PEPCK
and glucose-6-phosphatase genes (13, 51), these proteins may
be mutually bound and stabilized on the DEX-activated
MuRF1 promoter. Synergy may also be achieved via corecruit-
ment of transcriptional coactivators, such as histone acetyl-
transferases like CBP/p300, that have been shown to be inde-
pendently recruited by the liganded GR or FOXO1 in other
contexts (28, 34). Irrespective of the precise nature of the
GR-FOXO1 synergy, our results reveal that each factor is
required to create an optimal glucocorticoid-inducible MuRF1
promoter.
There are several important implications of our findings on
MuRF1 gene regulation. First, the activated GR may be an
important player in the control of muscle atrophy, but the
presence of nuclear FOXO1 is critical to achieve full induction
of the MuRF1 promoter and, quite possibly, multiple glucocor-
ticoid-regulated genes in skeletal muscle. Furthermore, the
glucocorticoid concentration needed to induce MuRF1 is sig-
nificantly reduced in the presence of FOXO1. A logical pre-
diction on the basis of these results is that conditions with both
elevated corticosteroids and active FOXO1 would lead to the
greatest degree of muscle wasting and altered metabolism. In
fact, diabetes mellitus presents such a condition (53). In several
models of diabetes, Akt is less active, potentially leading to
decreased inhibition of FOXO activity and/or increased nuclear
localization (55). Elevated nuclear FOXO1 would be available
to interact with the GR on the MuRF1 promoter to further
induce the gene above the level induced by corticosteroids
alone. Prolonged glucocorticoid exposure also leads to insulin
resistance (54), which might in turn lead to higher MuRF1
induction, accelerated muscle atrophy, and metabolic changes
as part of a vicious cycle. In contrast, the need for both nuclear
FOXO1 and activated GR to fully activate MuRF1 may be
protective and prevent unnecessary muscle breakdown. For
example, following intense exhaustive exercise, circulating
cortisol levels increase (6, 31), yet elevated cortisol is generally
not associated with an increase in the breakdown of contractile
proteins, nor is loss of muscle mass a consequence of endur-
ance training (33). Thus, the need for additional factors to fully
activate MuRF1 gene expression could protect the muscle from
breakdown under certain physiological conditions.
In summary, our results demonstrate a potent synergy be-
tween the GR and FOXO1 in transcriptional control of an
important gene linked to skeletal muscle atrophy and metabolic
control. Further understanding the molecular details of the
opposing effects of corticosteroids and insulin/IGF-I on gene
expression in skeletal muscle, the breadth of the GR-FOXO1
cooperation on muscle gene expression in general, and the very
nature of this robust synergy should have important ramifica-
tions for preventing inappropriate induction of genes involved
in muscle atrophy and altered metabolism in catabolic disease
states.
ACKNOWLEDGMENTS
We acknowledge Dr. Frank Gannon and members of the Gannon Labora-
tory at the European Molecular Biology Laboratory for generous support and
advice on the ChIP experiments.
GRANTS
This work was supported by a Kirchstein National Research Service Award
to D. S. Waddell and National Institute of Diabetes and Digestive and Kidney
Diseases Grant RO1-DK-075801 to J. D. Furlow and S. C. Bodine.
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