Neurogenetics (2006) 7: 81–91
DOI 10.1007/s10048-006-0031-7
ORIGINA L ARTI CLE
Patrick E. Baker . Jessica A. Kearney . Bendi Gong .
Anita P. Merriam . Donald E. Kuhn . John D. Porter .
Jill A. Rafael-Fortney
Analysis of gene expression differences between utrophin/
dystrophin-deficient vs mdx skeletal muscles reveals a specific
upregulation of slow muscle genes in limb muscles
Received: 23 August 2005 / Accepted: 6 December 2005 / Published online: 9 March 2006
# Springer-Verlag 2006
Abstract Dystrophin deficiency leads to the progressive
muscle wasting disease Duchenne muscular dystrophy
(DMD). Dystrophin-deficient mdx mice are characterized
by skeletal muscle weakness and degeneration but they
appear outwardly normal in contrast to DMD patients.
Mice lacking both dystrophin and the dystrophin homolog
utrophin [double knockout (dko)] have muscle degeneration similar to mdx mice, but they display clinical features
similar to DMD patients. Dko limb muscles also lack
postsynaptic membrane folding and display fiber-type
abnormalities including an abundance of phenotypically
oxidative muscle fibers. Extraocular muscles, which are
spared in mdx mice, show a significant pathology in dko
mice. In this study, microarray analysis was used to
characterize gene expression differences between mdx and
dko tibialis anterior and extraocular skeletal muscles in an
effort to understand the phenotypic differences between
these two dystrophic mouse models. Analysis of gene
expression differences showed that upregulation of slow
muscle genes specifically characterizes dko limb muscle
and suggests that upregulation of these genes may directly
account for the more severe phenotype of dko mice. To
P. E. Baker . J. A. Kearney . D. E. Kuhn .
J. A. Rafael-Fortney (*)
Department of Molecular and Cellular Biochemistry,
College of Medicine, The Ohio State University,
Columbus, OH, USA
e-mail: [email protected]
Tel.: +1-614-2927043
Fax: +1-614-2924118
B. Gong . A. P. Merriam . J. D. Porter
Department of Neurology, Case Western Reserve University,
Cleveland, OH, USA
B. Gong . A. P. Merriam . J. D. Porter
University Hospitals of Cleveland,
Cleveland, OH, USA
J. D. Porter
National Institutes of Neurological Disorders and Stroke,
6001 Executive Blvd, NINDS/NSC,
Bethesda, MD 2142, USA
investigate whether any upregulation of slow genes is
retained in vitro, independent of postsynaptic membrane
abnormalities, we derived mdx and dko primary myogenic
cultures and analyzed the expression of Myh7 and Myl2.
Real-time reverse transcriptase-polymerase chain reaction
analysis demonstrates that transcription of these slow genes
is also upregulated in dko vs mdx myotubes. This data
suggests that at least part of the fiber-type abnormality is
due directly to the combined absence of utrophin and
dystrophin and is not an indirect effect of the postsynaptic
membrane abnormalities.
Keywords Muscular dystrophy . Dystrophin . Myh7 .
Myl2 . Myosin heavy chain . Myosin light chain
Introduction
The dystrophin-associated protein complex (DAPC) is a
large, multimeric structure found at the sarcolemma of
muscle fibers that functions to fortify the integrity of the
cell membrane during contraction by linking the cytoskeleton with the extracellular matrix [1–3]. Dystrophin, a
cytoplasmic component of this complex, is bound to
F-actin of the subsarcolemmal cytoskeleton and to a
transmembrane glycoprotein of the DAPC, β-dystroglycan
[4–9]. Mutational inactivation of the dystrophin gene
causes Duchenne muscular dystrophy (DMD), an X-linked
neuromuscular degenerative disease [10, 11]. Patients with
DMD experience gradual muscle weakness beginning in
early childhood and are nonambulant by 12 years of age.
DMD patients typically die due to either cardiac failure
caused by wasting of cardiac muscles or respiratory
infections, a consequence of breathing difficulties caused
by weakening of the diaphragm muscle [12]. Histological
hallmarks of dystrophin-deficient skeletal muscle include
internal nuclei in muscle fibers undergoing cycles of
degeneration and regeneration and infiltration of inflammatory cells to these sites of degeneration [13, 14]. At the
molecular level, the loss of dystrophin is accompanied by
the reduction of DAPC at the sarcolemma [15–17].
82
The mdx (muscular dystrophy on the X-chromosome)
mouse strain has a nonsense mutation in exon 23 of the
dystrophin gene [18]. Although mdx mice lack functional
dystrophin, just as their human DMD counterparts, they
manifest much milder signs of the disease. Although mdx
skeletal muscle is characterized by necrosis, fibrous tissue
deposition, and reduction in force production, the loss of
overall muscle mass in mdx mice is minimal and mdx mice
live an almost normal life span [19]. The presence of the
dystrophin paralog utrophin in mdx skeletal and cardiac
muscles is believed to partially compensate for the lack of
dystrophin in the assembly of the DAPC [20].
In normal skeletal muscle fibers, utrophin is concentrated at the neuromuscular and myotendinous junctions;
however, utrophin is localized under the entire sarcolemma
in newly regenerated fibers present in mdx skeletal muscle
[21, 22]. Overexpression of utrophin in transgenic mdx
mice was shown to inhibit muscular dystrophy [20]. Mice
deficient for both dystrophin and utrophin [double knockout (dko)] exhibit clinical signs similar to that seen in DMD
patients, including joint contractures, kyphosis, short stature, breathing difficulty, and premature death (10–20
weeks for mice) [23].
Although the amount of muscle damage and regeneration is similar between mdx and dko littermates, skeletal
muscles of dko mice exhibit several additional abnormalities. Dko skeletal muscle is characterized by an almost
complete lack of postsynaptic folding at the neuromuscular
junctions (NMJs) and a fiber-type abnormality [23–26].
Quadriceps and tibialis anterior (TA) muscles from dko
mice almost exclusively contain oxidative skeletal muscle
fibers in contrast to the mixture of glycolytic and oxidative
fibers in these muscles from normal and mdx mice [27]. In
addition, dko muscles show a specific absence of fatigable
fibers and a shift toward slower myosin heavy chain protein
isoforms compared to mdx and control muscles [28].
Transgenic expression of truncated dystrophin protein
lacking the β-dystroglycan binding domain (Δcys) is able
to ameliorate both the postsynaptic membrane and fibertype abnormalities of dko skeletal muscle [27, 29].
However, this Δcys dystrophin protein is unable to prevent
muscle degeneration because it does not restore the
mechanical link between dystrophin and the DAPC.
These data indicate that providing a mechanical linkage
between the cytoskeleton and the extracellular matrix is not
the only function of dystrophin and utrophin.
It is important to note that dystrophinopathy does not
have a universal effect on all skeletal muscles. In DMD
patients and mdx mice, the extraocular muscles do not
display any of the functional and structural anomalies of
limb and diaphragm muscles. Extraocular muscles are
skeletal muscles of unique ontogeny with a diverse makeup
of fiber-types and display physiological characteristics
(weak, fast, and fatigue-resistant) that are contradictory in
other skeletal muscles. Fiber-types for the global and
orbital layers of extraocular muscles are broken down into
six categories including singly and multiply innervated
fibers of the orbital layer; red, intermediate, and pale singly
innervated fibers of the global layer; and global multiply
innervated fibers (reviewed in [30]). Adult extraocular
muscles continue expression of embryonic isoforms of
myosin heavy chain, coexpress myosin heavy chain
isoforms within fibers, and express a unique isoform in
addition to a slow-tonic isoform and the four isoforms of
myosin heavy chains present in other adult skeletal muscle
(reviewed in [30]).
In mdx extraocular muscle, utrophin protein is found at
elevated levels compared to both normal extraocular
muscles and to limb muscle of mdx mice [31]. This
increase in utrophin in mouse extraocular muscle appears
to spare this muscle group from destabilization of the
sarcolemma and the degeneration of fibers observed in mdx
limb muscle. This hypothesis is further supported by the
pathology in dko extraocular muscles [31]. However,
because some muscle fibers in extraocular muscle (the
orbital layer of the rectus muscle) of dko mice retain
protection against pathology, utrophin upregulation alone
cannot be responsible for sparing these muscle fibers.
To further characterize the differences between dko and
mdx skeletal muscles, we used microarray analysis to
compare global gene expression in TA and extraocular
muscles from dko vs mdx mice. This microarray analysis
showed mildly enhanced expression of genes that characterize the dystrophic process, which were similarly
shown to be greatly upregulated in mdx vs wild-type limb
muscles [32]. It is notable that slow muscle fiber-type
genes were upregulated in TA muscles from dko compared
to mdx mice. This class of genes was not upregulated in
dko vs mdx extraocular muscles nor was it reported to be
upregulated in mdx vs normal hind limb muscle, suggesting
that the combined loss of utrophin and dystrophin in limb
muscle affects the fiber-type at the level of transcription.
To determine whether any differences in slow muscle
gene expression in the combined absence of utrophin and
dystrophin compared to dystrophin alone persists in vitro,
we derived primary myogenic cultures from hind limbs of
mdx and dko mice. We performed real-time quantitative
reverse transcriptase-polymerase chain reaction (RT-PCR)
to quantitate Myl2 and Myh7 transcripts that were shown by
microarray analysis to be upregulated in dko muscle. The
Myl2 gene is normally expressed in cardiomyocytes and
slow, skeletal muscle. The Myh7 gene encodes the slow,
heavy chain isoform in adult skeletal muscle and in
prenatal ventricular muscle [33, 34]. We show that some
upregulation of the Myl2 and Myh7 genes persists in dko
myogenic cultures indicating that at least some of the
regulation of these slow muscle genes is a direct
consequence of the absence of dystrophin and utrophin
and is independent of the postsynaptic abnormalities in dko
skeletal muscle.
83
Materials and methods
Mouse husbandry
Male and female mdx;utrn+/− (mdx) mice were mated to
generate mdx;Utrn−/− (dko) mice and mdx;utrn+/+ (mdx)
mice. All mice were genotyped for the utrophin knockout
allele status by PCR as previously described [35, 36]. Wildtype C57BL/10 (C57) mice were maintained as a separate
inbred line. Mice were treated in accordance with the
Institutional Laboratory Animal Care and Use Committee.
Microarray analysis
DNA microarray methods were done as described
previously [37–39]. Briefly, DNA microarray analysis
was performed as three independent triplicates containing
RNA from muscles of four 8-week-old mice of each
genotype. Total RNA used for microarray runs were
extracted from TA and extraocular muscles (four rectus and
two oblique muscles were extracted from 12 mice for each
mdx and dko genotype) using the TRIzol reagent (GibcoBRL, Rockville, MD, USA). The resultant RNA pellets
were resuspended at 1 μg/μl in diethylpyrocarbonatetreated water and 8 μg of RNA was used in a reverse
transcription reaction (SuperScript II, Life Technologies,
Rockville, MD, USA) to generate single strand cDNA.
Double strand cDNA was generated and used in an in vitro
transcription (IVT) reaction to generate biotinylated cRNA.
Fifteen micrograms of fragmented cRNA was used in a
300-μl hybridization cocktail containing herring sperm
DNA and bovine serum albumin (BSA) as carrier
molecules, spiked IVT controls, and buffering agents. A
200-μl aliquot of this cocktail was used for hybridization to
Affymetric (Santa Clara, CA, USA) MG-U74Av2 microarrays for 16 h at 45°C. The microarrays were analyzed
using the manufacturer’s posthybridization wash, doublestain, and scanning protocols on an Affymetrix GeneChip
Fluidics Station 400 and with a Hewlett-Packard Gene
Array scanner.
The raw data from the microarray scans were analyzed
with both Microarray Affymetrix Suite (MAS) 5.0 and
Robust Multichip Average (RMA) algorithm [40] in
ArrayAssist 2.0 (Iobion Informatics, La Jolla, CA, USA).
Transcripts absent from all samples were excluded from
analysis. The MAS filter required that transcripts meet two
criteria. First, the transcripts must have consistent increase/
decrease call across all replicate comparisons at a given
time point based upon Wilcoxon’s signed rank test
(algorithm assesses probe pair saturation, calculates a
P value, and determines increase, decrease, or no change
calls). Secondly, the transcripts should have an average
fold difference value ≥2.0. Similarly, the RMA filter also
required that transcripts show an average fold difference
value ≥2.0. Transcripts were ultimately defined as
differentially expressed only if they passed filtering by
both algorithms.
The microarray raw data series and CEL files were
posted on the National Center for Biotechnology Information, Gene Expression Omnibus database (http://www.
ncbi.nlm.nih.gov/geo/) under series record accession
number GSE1463.
Cell culture
Total hind limb was dissected from 5-day-old mice and
dissolved in a filtered-sterilized solution of 1 mg/ml
collagenase (Sigma-Aldrich, St Louis, MO, USA) and
1 mg/ml BSA (Sigma-Aldrich) in phosphate-buffered saline
(Cellgro, Herndon, VA, USA) for 15 min at 37°C. The
supernatants were collected and filtered through a 40 μM
filter (BD Falcon, San Jose, CA, USA). The filtrates were
centrifuged at 1,000 rpm for 10 min. The pellets were
resuspended in 10 ml of F10 HAM nutrient solution
containing 15% horse serum, 200 mM L-glutamine,
2.5 ng/ml of amphotericin B, 1× penicillin/streptomycin,
and 6 ng/ml of basic fibroblast growth factor (bFGF) (all
from Sigma-Aldrich) and were plated onto gelatin-coated
(0.2%) plates. Cells were passaged by trypsinization and
plated onto uncoated plastic for 1 h to remove fibroblasts.
The media containing myogenic cells was then transferred to
gelatin-coated plates. Cells were fed every 12 h with growth
media. After 1 week, cells were switched to media
containing 2 ng/ml bFGF. When cells were approximately
95% confluent, media containing 5% horse serum and no
bFGF was added to the cells for 24 h to initiate differentiation. Twenty-four hours later, the primary cultures were
refed with media containing 10% horse serum. Nine-day
postdifferentiation fused myotubes were harvested and
suspended in RNALater (Ambion, Austin, TX, USA)
stabilization buffer and stored at −80°C.
Immunocytochemistry
Cells were plated onto chambered glass slides coated with
poly-D-lysine and laminin (BD Biosciences, San Jose, CA,
USA). Cultured cells were allowed to grow to confluency
and switched as described previously to serial differentiation media to promote myotube formation. Immunostaining was performed on day 9 postdifferentiated cultures.
Cells were fixed by the addition of an equal volume of
3.7% formaldehyde solution in potassium phosphatebuffered saline (KPBS) to the cell culture followed by
incubation on ice for 5 min. Media/formaldehyde solution
was removed and 3.7% formaldehyde was added for 3 min
on ice. Slides were rinsed in cold KPBS and then extracted
with 0.5% Triton X-100 (BioRad, Hercules, CA, USA) in
KPBS for 5 min on ice and rinsed in KPBS. Samples were
blocked in 1% normal goat serum (NGS) (Vector
Laboratories, Burlingame, CA, USA) in KPBS for
30 min. The slides were incubated overnight at 4°C with
monoclonal antibodies against fast (MHC-f; 1:25), slow
(MHC-s; 1:40), or developmental (MHC-d; 1:25) myosin
84
heavy chain isoforms (Novocastra Laboratories, Newcastle
Upon Tyne, UK) in KPBS containing 1% NGS and 0.1%
Tween-20 (Sigma-Aldrich). Slides were rinsed five times
in KPBS and incubated with 1:200 anti-mouse-CY3labeled secondary (Jackson Laboratories, Bar Harbor, ME,
USA) in 1% NGS and 0.1% Tween-20 solution for 1 h at
room temperature. Slides were rinsed five times and
mounted in Vectashield (Vector Laboratories) containing
1 μl/ml DAPI (Sigma-Aldrich).
Real-time RT-PCR
To generate standards for the real-time RT-PCR, cDNA was
first synthesized from C57 heart RNA using the SuperScript First Strand Synthesis System for RT-PCR (Invitrogen, Carlsbad, CA, USA). The cDNA served as template
for subsequent PCR using the 18S-T7/18S-R, the Myl2-T7/
Myl2-R, and the Myh7-T7/Myh7-R primer pairs (Table 1)
in separate reactions. The sequence for the T7 promoter
was included at the 5′-end of the forward primers of each
pair. This design resulted in the synthesis of PCR products
containing the T7 promoter, which allowed the PCR
products to be used as DNA templates for IVT with the
MEGA Shortscript Kit (Ambion). The Myh7, Myl2, and
18S RNA from the IVT reactions were quantified using a
spectrophometer and were aliquoted at concentrations of
100, 10, 1, 0.1, and 0.01 ng/4 μl.
For real-time RT-PCR, total RNA was isolated from
material that was frozen at −80°C or stored in RNALater
using the RNeasy Mini kit (BioRad). The sources of the
RNA were C57 heart, dko heart, mdx myotube cultures,
dko myotube cultures, C57 quadriceps, dko quadriceps,
and mdx quadriceps. The RNA was quantified using a
spectrophotometer. For 18S detection, 100 ng/4 μl of
sample RNA was used; 500 ng/4 μl of RNA was used for
Myl2 detection; and 400 ng/4 μl of RNA was used for
Myh7 detection. Four microliters of RNA at the appropriate
concentration, 6.6 μl of sterile water, 1.3 μl of 50 mM Mn
(OAc)2, and 0.75 μl of 0.25 μM of forward and reverse
primers (18SII-F/18SII-R, Myl2-F/Myl2-R, or Myh7IV-F/
Myh7IV-R) (Table 1) were added to 7.5 μl of LightCycler
Table 1 Primers used in RT-PCR
Primers
Primer sequence
18S-T7
5′-TAATACGACTCACTATAGGGTGACTCAA
CACGGGAAACCTCAC-3′
5′-TGACTCAACACGGGAAACCTCAC-3′
5′-ATCCAATCGGTAGTAGCGACGG-3′
5′-TAATACGACTCACTATAGGGCGACAAGAAT
GACCTAAGGGACAC-3′
5′-CGACAAGAATGACCTAAGGGACAC-3′
5′-GCCAAGACTTCCTGTTTATTTGCG-3′
5′-TAATACGACTCACTATAGGGCACTATGCTGG
CACTGTGGACTAC-3′
5′-CACTATGCTGGCACTGTGGACTAC-3′
5′-TGGGTTCAGGATGCGATACCTC-3′
18S-F
18S-R
Myl2-T7
Myl2-F
Myl2-R
Myh7-T7
Myh7-F
Myh7-R
RNA Master SYBR Green I (Roche, Germany). The RTPCR reactions were carried out using the LightCycler realtime PCR machine (Roche) in capillary tubes (Roche)
specifically designed for the apparatus. The reverse
transcription phase was performed at 61°C for 20 min
followed by the amplification phase. In the amplification
phase, the samples underwent 30 cycles of 95°C for 5 s,
50°C for 50 s, and 72°C for 13 s. After the amplification
phase, the samples were denatured at 95°C for 2 min.
Melting curve analysis was conducted after the following
treatment: 95°C for 5 s, 65°C for 15 s, and 95°C for 0 s.
To calculate the actual number of transcripts in each
RNA sample, standard curves were generated from analysis of the known amounts of Myl2, Myh7, and 18S RNA
from IVT. The LightCycler software calculated the amount
of PCR product yielded from the standards after the RTPCR and this data was used to create standard curves,
which allowed the calculation of the copy number of Myl2,
Myh7, or 18S RNA in the RNA samples from the muscle
and cell culture samples.
Real-time RT-PCR for each gene was performed in
separate runs each consisting of the RNA standards and
RNA samples from C57 heart, dko heart, mdx myotube
culture, dko myotube culture, C57 quadriceps, dko quadriceps, and mdx quadriceps. The raw data for each sample
was the mean of triplicate RT-PCR runs for each gene. The
mean of these runs was corrected by dividing by 4 (for the
Myh7 assays) or 5 (for the Myl2 assays) because either four
or five times as much RNA was used for the Myh7 and
Myl2 assays than what was used in the 18S assay. This
corrected mean copy number of the three runs for a given
sample was then divided by the mean of the copy number
of three runs of 18S for that sample. Statistical analysis of
data was performed using one-way ANOVA with the
Scheffe multiple comparison post hoc test (SSPS version
12.0, Chicago, IL, USA).
Results
Genome-wide expression changes in dko vs mdx
skeletal muscles
We used DNA microarray analyses to identify global
changes in gene expression in dko skeletal muscles
compared to those of mdx mice. We have previously
identified different sets of gene expression changes in
extraocular vs limb muscles in response to dystrophindeficiency [37, 38]. We have also shown that dko mice
show a pathology in extraocular muscles, which are spared
in mdx [31]. Therefore, we compared expression profiles
between dko and mdx mice for the limb muscle, TA, and
extraocular muscles. Independent triplicate analyses were
conducted for each muscle group and mouse strain and the
data were analyzed using both the Affymetrix and RMA
algorithms.
Seventy-five and 30 transcripts in TA and extraocular
muscles, respectively, were identified to be differentially
expressed ± two-fold between dko and mdx using the
85
Affymetrix algorithm. In both muscles, the overwhelming
majority of differentially regulated transcripts were increased in expression level in dko (83% for TA and 87% for
extraocular muscle). For the hind limb, transcriptional
patterns previously reported for mdx vs the wild-type
control strain for both mdx and dko were further enhanced
in dko vs mdx muscles [32]. We observed further increases
above those reported for mdx in the expression of
transcripts encoding proteins contributing to the extracellular matrix (e.g., Ctgf, Mmp3, Col8a1, and Col3a1),
cytoskeleton (e.g., Tmsb 10 and Tubb2), muscle regeneration (e.g., Myh3, Myog, and Acta2), inflammation (e.g.,
Cd53, Ccl8, C1qb, and Gp49a), and proteolysis (e.g., Lzps, Lyzs, and Capn6) functional categories that represent the
major events in dystrophic skeletal muscle. In contrast, the
extraocular muscles showed nearly no changes in extracellular matrix transcripts and upregulation of only some
genes related to inflammation (e.g., Ccl2, Lgals3, and
Gp49b) and proteolysis (Lzp-s and Lyzs).
Determination of differentially expressed transcripts
using the RMA algorithm and a ± twofold difference
cutoff identified fewer differentially expressed genes in
dko vs mdx comparisons, 28 for hind limb and 15 for
extraocular muscle. We chose to designate only those genes
in the intersection of the Affymetrix and RMA algorithms
as differentially expressed in dko muscles, resulting in 22
transcripts for TA (Table 2) and 12 transcripts for
extraocular muscle (Table 3).
In addition to the categories of differentially expressed
genes that generally characterize dystrophic muscle, dko
TA showed an upregulation of genes that characterize slow
muscle fiber-type. This class of genes was not shown to be
differentially expressed between mdx and wild-type limb
muscles [32]. The differentially expressed slow fiber
transcripts include those that encode sarcomeric proteins
(Myl2, Myh7, Tnnc1, Tnnt1, Tnnt2, Tpm3, and Tnni1) and
the slow isoform of a sarcoplasmic reticulum pump
(Atp2a2). This induction of slow fiber transcripts represented a significant functional response, amounting to 42%
of the upregulated genes observed for the TA. None of
these genes nor any other genes linked to slow fiber
function were differentially expressed in dko vs mdx
extraocular muscle (moreover, none of the eight slow fiber
genes identified for dko TA even met the threshold for
extraocular muscle for either the Affymetrix or RMA
algorithms).
Analysis of the Myl2 and Myh7 genes in dko limb
muscle and myogenic culture samples
Because postsynaptic membrane and fiber-type abnormalities are both present in dko mice and are both ameliorated
by the Δcys transgene, it was not determined whether these
two phenotypes are linked or whether the absence of
utrophin and dystrophin directly leads to alterations in
Table 2 Gene expression differences in dko vs mdx TA muscle
Affymetrix probe ID
Slow muscle transcripts
93050_at
98616_f_at
101063_at
161361_s_at
99570_s_at
100593_at
93266_at
98561_at
Other transcripts
160753_at
103570_at
93294_at
101993_at
92593_at
93037_i_at
100325_at
160487_at
93038_f_at
100611_at
102990_at
98942_r_at
101489_at
93584_at
Accession no.
Gene (symbol)
Fold difference
M91602
AJ223362
M29793
AV213431
AF029982
L47600
U04541
AJ242874
Myosin, light polypeptide 2, regulatory, cardiac, slow (Myl2)
Myosin, heavy polypeptide 7, cardiac muscle, beta (Myh7)
Troponin C, cardiac/slow skeletal (Tnnc1)
Troponin T1, skeletal, slow (Tnnt1)
ATPase, Ca++ transporting, cardiac muscle, slow twitch 2 (Atp2a2)
Troponin T2, cardiac (Tnnt2)
Tropomyosin 3, gamma (Tpm3)
Troponin I, skeletal, slow 1 (Tnni1)
24.7
22.3
5.9
3.7
3.2
2.8
2.7
2.2
M74753
AI315647
M70642
X56304
D13664
M69260
M65027
M19436
M69260
M21050
AA655199
AW125284
D12780
V00821
Myosin, heavy polypeptide 3, skeletal muscle, embryonic (Myh3)
C1q and tumor necrosis factor related protein 3 (C1qtnf3)
Connective tissue growth factor (Ctgf)
Tenascin C (Tnc)
Periostin, osteoblast-specific factor (Postn)
Annexin A1 (Anxa1)
Glycoprotein 49 A (Gp49a)
Myosin, light polypeptide 4 (Myl4)
Annexin A1 (Anxa1)
Lysozyme (Lyzs)
Procollagen, type III, alpha 1 (Col3a1)
RIKEN cDNA 2310032D16
S-adenosylmethionine decarboxylase 1 (Amd1)
Immunoglobulin heavy chain 6 (heavy chain of IgM) (Igh-6)
3.8
3.6
3.2
3.1
2.5
2.5
2.4
2.3
2.2
2.1
2.1
−2.1
−2.4
−3.4
86
Table 3 Gene expression differences in dko vs mdx extraocular muscle
Affymetrix probe ID
Accession no.
Gene (symbol)
Fold difference
102736_at
97519_at
95706_at
98543_at
100325_at
101753_s_at
160469_at
99071_at
100611_at
98575_at
94057_g_at
101566_f_at
M19681
X13986
X16834
AJ223208
M65027
X51547
M62470
L20315
M21050
X13135
M21285
M17818
Chemokine (C-C motif) ligand 2 (Ccl2)
Secreted phosphoprotein 1 (Spp1)
Lectin, galactose binding, soluble 3 (Lgals3)
Cathepsin S (Ctss)
Glycoprotein 49 A (Gp49a)
P lysozyme structural (Lzp-s)
Thrombospondin 1 (Thbs1)
Macrophage expressed gene 1 (Mpeg1)
Lysozyme (Lyzs)
Fatty acid synthase (Fasn)
Stearoyl-Coenzyme A desaturase 1 (Scd1)
Major urinary protein 1 (Mup1)
6.1
5.6
5
4.5
3.4
2.8
2.6
2.6
2.4
−2
−2.3
−2.8
fiber-type pathways [27]. It is possible that the postsynaptic
membrane abnormality present in dko muscle may alter the
signal that the muscle fiber receives from the motor neuron
through the NMJ. Skeletal muscle was shown to switch
muscle fiber type in vivo in response to changes in
innervation, physical activity, hormones, and aging [41].
Therefore, we sought to determine whether slow muscle
gene expression differences between dko and mdx muscles
persist in the absence of these in vivo factors.
To investigate whether the upregulation of Myl2 and
Myh7 (two slow fiber-type genes that demonstrated 24.7and 22.3-fold increases in expression compared to mdx) is
independent of dko in vivo postsynaptic NMJ abnormalities, we derived primary myogenic cultures from dko and
control mdx littermates. Hind limb muscle from day 5
neonatal littermates was used to isolate myogenic cells. In
parallel, the mice were genotyped and cultures derived
from two dko and two mdx littermates were expanded.
Each culture was passed nine times to eliminate any carryover fibers from the original muscle and preplated on
uncoated plastic culture dishes to remove the majority of
fibroblasts. At 95% confluency, passage nine dko and mdx
cultures were switched from growth media to differentiation media and allowed to fuse for a period of 9 days. The
morphology, doubling rate, and rate of fusion were
indistinguishable between dko and mdx cultures.
To determine differences in gene expression of the Myh7
and Myl2 genes, we performed real-time RT-PCR in
triplicate independent runs on total RNA derived from
day 9 postdifferentiation dko and mdx cultures. Real-time
RT-PCR enables us to determine target transcript copies in
a given sample. To control for possible variations in the
amount of RNA between samples, the raw data was
normalized with the copy number of 18S ribosomal RNA
calculated for each sample in a parallel real-time RT-PCR
run. The 18S was chosen as a control because other
housekeeping genes related to oxidative and glycolytic
pathways are known to vary in the dko mice with skeletal
muscle fiber-type abnormalities compared to controls. To
confirm the data from the microarray analysis performed
on dko and mdx skeletal muscle and to compare this data
with data obtained from real-time RT-PCR analysis of the
cell cultures, we included in this study RNA from
quadriceps muscles isolated from dko, mdx, and C57
mice. Also, as positive controls, we included total heart
RNA from C57 and dko mice where Myl2 and Myh7
should be expressed. The degree of statistical significance
of differences in gene expression between samples was
determined by one-way ANOVA with the Scheffe multiple
comparison post hoc test.
C57 heart expressed higher levels of Myh7 than dko
heart did [146 copies (per 106 copies of 18S) vs 74 copies; a
twofold difference] although this is not statistically significant (Fig. 1). As predicted, dko quadriceps (280±199
copies) expressed higher levels of Myh7 than mdx (2±0.4
copies; a 114-fold difference) and C57 (2±0.6 copies; 163fold difference) quadriceps (Fig. 1). Myh7 was also found
to be upregulated in the dko myotube culture samples
compared to the mdx culture (337±112 copies vs 74±3
copies; a 4.6-fold difference) (Fig. 1). The increase of
Myh7 transcripts in dko in the cell culture group and the
quadriceps group was determined to be statistically significant. The low expression level of Myh7 in C57 quadriceps
muscle was consistent with previous studies of Myh7 in
quadriceps using conventional RT-PCR experiments [42].
Real-time RT-PCR analysis showed that Myl2 (Fig. 2)
has a more robust expression compared to Myh7 (Fig. 1)
expression in C57 and dko heart. This data is consistent
with reports of downregulation of Myh7 expression in the
heart shortly after birth [33, 34]. A higher level of Myl2
gene expression was observed in dko quadriceps (249±149
copies) than in C57 quadriceps (10±4 copies; 25-fold
difference) and in mdx quadriceps (4±2; 63-fold difference)
(Fig. 2). Elevated levels of Myl2 were also observed in dko
vs mdx myotube cultures (46±24 copies vs 9±3 copies;
fivefold difference) and dko vs C57 heart RNA samples
(19,894±3,700 copies vs 9,862±1,203 copies, respectively;
twofold difference) (Fig. 2). Although the number of Myl2
copies detected in dko vs mdx myotubes and quadriceps
had nonoverlapping standard deviations, statistical evalua-
87
copies of Myh7/ million copies of 18S rRNA
600.0
500.0
400.0
300.0
200.0
†
100.0
‡
‡
C57
Quadricep
MDX
Quadricep
0.0
Dko Heart
C57 Heart
MDX
Dko Myotube
Dko
Myotube
Quadricep
elevated levels of Myh7 RNA compared to mdx myotube cultures
and mdx quadriceps, respectively. However, Myh7 expression in dko
heart was lower than in C57 heart sample. A P value of less than
0.05 denotes statistical significance under analysis with one-way
ANOVA. Dagger (†) denotes that result is significantly different
(P<0.05) from dko myotube culture. Double dagger (‡) denotes that
result is significantly different (P<0.05) from dko quadriceps sample
tion determined that the elevated levels detected in dko
myotubes and quadriceps were not significantly different
from mdx myotubes and quadriceps, respectively.
Real-time RT-PCR was able to detect the low levels of
Myl2 transcripts in the C57 and the mdx quadriceps RNA
samples (9.9±4.4 copies and 3.6± 2.6 copies, respectively)
(Fig. 2). This low level detection underscores the greater
sensitivity of real-time analysis compared to Northern
blotting, which in at least one previous study, failed to
show any evidence of Myl2 in quadriceps muscle [43].
Copies of Myl2 RNA per 1 million copies of 18S rRNA
Fig. 1 Expression of Myh7 in dko, mdx, and C57 muscle samples.
Real-time RT-PCR analysis was performed to quantitate Myh7
transcripts present in total RNA derived from each sample listed on
the x-axis. Each bar represents the mean (±SD) of Myh7 transcripts
per million copies of 18S from three separate runs of RT-PCR using
the same amount of total RNA for Myh7 (100 ng) and 18S (25 ng)
assays. Dko myotube cultures and dko quadriceps expressed
100000.0
†
‡
10000.0
‡
1000.0
‡
100.0
†
†
†
‡
†
‡
†
10.0
1.0
Dko Heart
C57 Heart MDX Myotube Dko Myotube
Fig. 2 Expression of Myl2 in dko, mdx, and C57 muscle samples.
Real-time RT-PCR analysis was performed to quantitate Myl2
transcripts present in total RNA derived from each sample listed on
the x-axis. Each bar represents the mean (±SD) of Myl2 transcripts
per million copies of 18S from three separate runs of RT-PCR using
the same amount of total RNA for myplc (125 ng) and 18S (25 ng)
assays. Dko myotube cultures, dko quadriceps, and dko hearts
expressed elevated levels of Myl2 RNA compared to mdx myotube
Dko
C57
Quadricep Quadricep
MDX
Quadricep
cultures, mdx quadriceps, and mdx hearts, respectively. Statistical
analysis using one-way ANOVA determined significant difference
between both heart samples and the rest of the samples, but failed to
find significant differences between the remainder of the samples.
Dagger (†) denotes that result is significantly different (P<0.05)
from dko heart. Double dagger (‡) denotes that result is significantly
different (P<0.05) from C57 heart
88
Localization of myosin heavy chain isoforms in mdx
and dko myotube cultures
Because dko skeletal muscle in vivo is characterized by
oxidative enzymes and slow muscle protein isoforms, we
tested whether the upregulation of slow muscle gene
expression observed in vitro correlated with a shift to slow
muscle protein isoform in myotubes 9 days after switching
the dko myogenic cultures to differentiation medium.
Immunocytochemistry was conducted on mdx and dko
myogenic cultures with antibodies against slow, fast, and
developmental myosin heavy chain isoforms (MHC-s,
MHC-f, and MHC-d) (Fig. 3). Although each of these
antibodies reacts with similar affinities to dko muscles in
vivo, antibodies against MHC-s only detect very low
amounts of protein in a small percentage of myotubes in
both dko and mdx (Fig. 3b). Most of the myotubes in both
the dko and mdx cultures are brightly stained with antibody
against MHC-f (Fig. 3a), which was shown to positively
stain 2-week-old dko and mdx diaphragm muscle fibers
[26]. Only a few fibers in both the dko and mdx cultures are
stained with MHC-d (Fig. 3c). This result is in contrast
Fig. 3 Immunocytochemistry for MHC isoforms on differentiated
myotubes derived from mdx and dko primary cultures. Dko and mdx
myotubes were incubated with antibodies against a MHC-f,
b MHC-s, and c MHC-d and were visualized using anti-Cy3labeled secondary antibody (red). Nuclei were stained with DAPI
(blue). Merged images of red and blue channels are shown. No
differences were seen between the dko and mdx genotypes for any of
the three antibodies. MHC-slow staining was overall less intense
than the other two isoforms. Although MHC-d staining is intense,
fewer positively stained myotubes than those immunostained with
the fast and slow isoforms were present
with dko diaphragm muscle and mdx and dko limb muscle,
which show robust staining with MHC-d antibody in twoand four-week-old neonates, respectively [26, 44]. Taking
this into consideration, our data suggests that myotubes
9 days after the switch to differentiation media are not
mature enough to show incorporation of slow MHC
proteins into the contractile apparatus due to the increased
levels of transcription of Myh7.
Discussion
We utilized DNA microarray analysis in an attempt to
identify genes with different expression in mdx vs dko TA
and extraocular skeletal muscles that may contribute to
differences between these two dystrophic mouse models in
the severity of muscle pathologies, fiber-type, and overt
clinical features. Twenty-two transcripts were differentially
expressed greater than twofold between mdx and dko TA
muscles. Eight of these transcripts (Myl2, Myh7, Tnnc1,
Tnnt1, Tnnt2, Tpm3, Tnni1, and Atp2a2) were from genes
that are characteristic of slow fiber-type muscle. These
genes were upregulated in dko TA muscle when compared
to mdx and were not previously shown to be differentially
expressed in previous comparisons of limb muscles from
mdx vs the wild-type control for mdx and dko using
microarray technology [32]. These data therefore correlate
with the previous observations of increases in slower
myosin heavy chain protein isoforms and oxidative
metabolism in dko compared to mdx limb muscles [27, 28].
Despite these interesting findings from microarray
analysis, this technique was unsuccessful in identifying
differential expression of genes involved in signaling
pathways that could bridge the gap between dystrophin and
utrophin loss with fiber-type alteration and postsynaptic
abnormalities present in TA or the pathology present in
extraocular muscles from dko mice. Therefore, the
different effects of dystrophin and/or utrophin deficiency
on extraocular muscle compared to TA muscles may be
more attributable to their differing developmental origin,
functional properties, hormonal response, and myosin
phenotype than transcriptional modulation.
From the finding that slow-type genes were upregulated
in dko limb muscle, a central question emerged: whether
upregulation of slow muscle fiber type genes in dko is a
direct result of the loss of dystrophin and utrophin and
independent of in vivo extrinsic factors such as neuronal
input, physical activity, and hormones. To address this
central question and to confirm the microarray data, we
examined the expression profile of Myh7 and Myl2 in
muscle cell culture and skeletal muscle tissue using realtime RT-PCR. Real-time RT-PCR was utilized for this
study because of its superior sensitivity and quantitative
capability compared to Northern blot analysis and
conventional RT-PCR. The real-time data for both genes
showed the same trends of upregulation in dko vs mdx limb
muscle as the microarray data. Furthermore, the upregulation of Myh7 and Myl2 was also observed in dko myotubes
compared to mdx myotubes. Higher levels of Myh7 and
89
Myl2 in dko vs mdx myotubes suggest that differentiated
myogenic culture can mimic the gene expression profile
observed in muscle tissue. Another implication of these
data is that the upregulation of these genes is due, at least in
part, to the absence of utrophin and dystrophin proteins and
is not due to external cues caused by postsynaptic
abnormalities in vivo.
It is also unlikely that the primary cultures are still under
the influence of in vivo factors. Several studies reported
significant variation between the fiber-type phenotype of
myotube cultures and that of the muscle tissue from which
they were derived [45–47]. A recent study compared
myosin heavy chain isoform expression in several different
muscle types with primary cell cultures that were derived
from these same muscles [42]. This study found that
regardless of the fiber-type phenotype of the donor site, the
cell cultures all expressed similar levels of Myh7. This
homogeneity in the cell cultures and their dissimilarity with
the donor muscle were attributed to the great care taken to
remove the initial muscle fibers from the proliferative
satellite cells. Possible contamination with this muscle
tissue could provide signals for the direction of the fibertype phenotype of the satellite cells. Our cell culture was
performed with similar stringency, therefore suggesting the
validity of the data showing Myh7 and Myl2 upregulation
in dko myotube cultures.
The real-time data for in vivo skeletal muscle samples
indicate a substantially greater level of upregulation in dko
than the microarray study in the expression of both Myl2
(63× vs 24.7×) and Myh7 (114× vs 22.3×) genes. These
differences could be explained by a greater sensitivity of
real-time RT-PCR in quantitative analysis compared to
microarray, the extremely stringent statistical analysis
applied to generate the microarray data, or the difference
in abundance of the control (18S) and experimental (Myl2
and Myh7) RNAs analyzed by RT-PCR. Although the
number of copies of Myh7 transcript is higher in dko
myotubes than in dko quadriceps, the upregulation in dko
compared to mdx in the differentiated myotube culture is
only 4.7-fold compared with a 63-fold increase for dko vs
mdx quadriceps samples. This difference is due to the
presence of a higher number of copies in the mdx myotube
culture (74 copies) than in the mdx quadriceps (2 copies).
Another explanation for lower level of upregulation of
Myh7 gene expression in culture compared to in vivo may
be due to the immaturity of the myotubes. Our analysis of
MHC protein isoforms by immunocytochemistry confirms
that gene expression changes were not yet entirely
incorporated into protein. It is possible that the real-time
RT-PCR data acquired from RNA isolated from day 9
myotubes may be measuring levels of gene expression that
are continuing to increase in dko vs mdx cultures. However,
after this time point, the myotubes began to vigorously
contract and lift off the coated culture surface and therefore
could not be analyzed at much later time points.
The third and possibly most plausible hypothesis for the
differences in the amount of upregulation of gene expression seen in vivo and in vitro is that the gene expression of
Myh7 and Myl2 is partially dependent upon downstream
factors caused by the structural alterations in the postsynaptic membrane in the combined absence of utrophin and
dystrophin. Physiological experiments conducted nearly
40 years ago showed that electrical stimulation of muscle
fibers at different frequencies can lead to different fibertype properties of muscle [48]. These seminal experiments
demonstrated that nerve fibers excised from fast muscle
and, subsequently, used to reinnervate slow muscle could
induce the slow muscle to become fast. These data would
suggest that the absence of innervation in cultured
myotubes might account for the attenuated upregulation
of slow-muscle gene expression. However, the persistence
of upregulation of slow-muscle gene expression observed
in this study between dko and mdx myogenic cultures is the
first evidence that the absence of utrophin and dystrophin
in dko muscle has an effect on downstream gene expression that is independent of their role at the postsynaptic
membrane.
It is of interest that the RNA levels of Myh7 and Myl2 are
elevated in dko quadriceps where these genes are normally
found in only trace amounts [42, 43]. This observation
raises the question of what is the consequence of the
improper expression of these genes. The Myh7 gene is
normally expressed during embryonic development in the
heart of large mammals but diminishes at birth, while
expression remains in slow skeletal muscle [34, 49].
However, there are some instances of abnormal Myh7
expression that suggest that dysregulation of Myh7
expression may have a pathological role. Myh7 is known
to be expressed abnormally in the adult heart as a result of
diabetes, mechanical overload, and alterations in thyroid
hormone levels [33, 34]. Also, a missense mutation in
Myh7 gene causes improper accumulation of myosin and
formation of aberrant filamentous structures in skeletal
muscle resulting in skeletal myopathy but without cardiomyopathy [50]. Recently, mutations in Myh7 were shown
to account for a large number of cases of hypertrophic and
dilated cardiomyopathy [51, 52]. These associations
between Myh7 and disease suggest that the upregulation
of Myh7 in dko may contribute to differences in phenotypic
severity in dko vs mdx mice.
Acknowledgements This work was supported by the Muscular
Dystrophy Association (to JRF), NIH R01 EY12779 (to JDP), and in
part, by a Career Award in the Biomedical Sciences from the
Burroughs Wellcome Fund (to JRF). PEB was supported by an NIH
supplement (AR47034-S). We would like to thank Jaimy Lekan,
Katherine L. Gardner, Jonathan Edwards, and Chad Groer for
technical assistance and the Biochemistry and Molecular Biology
Core, Department of Veterinary Biosciences, OSU for the use of the
real-time PCR machine.
References
1. Campbell KP (1995) Three muscular dystrophies: loss of
cytoskeleton-extracellular matrix linkage. Cell 80:675–679
2. Ahn AH, Kunkel LM (1993) The structural and functional
diversity of dystrophin. Nat Genet 3:283–291
90
3. Tinsley JM, Blake DJ, Zuellig RA, Davies KE (1994)
Increasing complexity of the dystrophin-associated protein
complex. Proc Natl Acad Sci U S A 91:8307–8313
4. Hemmings L, Kuhlman PA, Critchley DR (1992) Analysis of
the actin-binding domain of alpha-actinin by mutagenesis and
demonstration that dystrophin contains a functionally homologous domain. J Cell Biol 116:1369–1380
5. Rybakova IN, Amann KJ, Ervasti JM (1996) A new model for
the interaction of dystrophin with F-actin. J Cell Biol 135:
661–672
6. Way M, Pope B, Cross RA, Kendrick-Jones J, Weeds AG
(1992) Expression of the N-terminal domain of dystrophin in E.
coli and demonstration of binding to F-actin. FEBS Lett
301:243–245
7. Suzuki A, Yoshida M, Hayashi K, Mizuno Y, Hagiwara Y,
Ozawa E (1994) Molecular organization at the glycoproteincomplex-binding site of dystrophin. Three dystrophin-associated proteins bind directly to the carboxy-terminal portion of
dystrophin. Eur J Biochem 220:283–292
8. Jung D, Yang B, Meyer J, Chamberlain JS, Campbell KP
(1995) Identification and characterization of the dystrophin
anchoring site on beta-dystroglycan. J Biol Chem 270:
27305–27310
9. Rosa G, Ceccarini M, Cavaldesi M, Zini M, Petrucci TC (1996)
Localization of the dystrophin binding site at the carboxyl
terminus of beta-dystroglycan. Biochem Biophys Res Commun
223:272–277
10. Koenig M, Monaco AP, Kunkel LM (1988) The complete
sequence of dystrophin predicts a rod-shaped cytoskeletal
protein. Cell 53:219–226
11. Monaco AP, Neve RL, Colletti Feener C, Bertelson CJ, Kurnit
DM, Kunkel LM (1986) Isolation of candidate cDNAs for
portions of the Duchenne muscular dystrophy gene. Nature
323:646–650
12. Emery AEH (1993) Duchenne muscular dystrophy, 2nd edn.
Oxford Univ. Press, Oxford
13. Bradley WG, Hudgson P, Larson PF, Papapetropoulos TA,
Jenkison M (1972) Structural changes in the early stages of
Duchenne muscular dystrophy congenital muscular dystrophy.
J Neurol Neurosurg Psychiatry 35:451–455
14. Conen PE, Bell CD (1970) Histopathological changes in
Duchenne muscular dystrophy. J Neurol Sci 10:163–171
15. Matsumura K, Ervasti JM, Ohlendieck K, Kahl SD, Campbell
KP (1992) Association of dystrophin-related protein with
dystrophin-associated proteins in mdx mouse muscle. Nature
360:588–591
16. Matsumura K, Tome FM, Collin H, Leturcq F, Jeanpierre M,
Kaplan JC, Fardeau M, Campbell KP (1994) Expression of
dystrophin-associated proteins in dystrophin-positive muscle
fibers (revertants) in Duchenne muscular dystrophy.
Neuromuscul Disord 4:115–120
17. Ervasti JM, Ohlendieck K, Kahl SD, Gaver MG, Campbell KP
(1990) Deficiency of a glycoprotein component of the
dystrophin complex in dystrophic muscle. Nature 345:315–319
18. Sicinski P, Geng Y, Ryder-Cook AS, Barnard EA, Darlison
MG, Barnard PJ (1989) The molecular basis of muscular
dystrophy in the mdx mouse: a point mutation. Science
244:1578–1580
19. Mattson MP (2001) Pathogenesis of neurodegenerative disorders. In: Contemporary neuroscience. Humana, Totowa, NJ,
pp x and 294
20. Tinsley J, Deconinck N, Fisher R, Kahn D, Phelps S, Gillis JM,
Davies K (1998) Expression of full-length utrophin prevents
muscular dystrophy in mdx mice. Nat Med 4:1441–1444
21. Bewick GS, Nicholson LV, Young C, O’Donnell E, Slater CR
(1992) Different distributions of dystrophin and related proteins
at nerve-muscle junctions. Neuroreport 3:857–860
22. Law DJ, Allen DL, Tidball JG (1994) Talin, vinculin and DRP
(utrophin) concentrations are increased at mdx myotendinous
junctions following onset of necrosis. J Cell Sci 107:
1477–1483
23. Deconinck AE, Potter AC, Tinsley JM, Wood SJ, Vater R,
Young C, Metzinger L, Vincent A, Slater CR, Davies KE
(1997) Postsynaptic abnormalities at the neuromuscular junctions of utrophin-deficient mice. J Cell Biol 136:883–894
24. Lyons PR, Slater CR (1991) Structure and function of the
neuromuscular junction in young adult mdx mice. J Neurocytol
20:969–981
25. Nagel A, Lehmann-Horn F, Engel AG (1990) Neuromuscular
transmission in the mdx mouse. Muscle Nerve 13:742–749
26. Deconinck AE, Rafael JA, Skinner JA, Brown SC, Potter AC,
Metzinger L, Watt DJ, Dickson JG, Tinsley JM, Davies KE
(1997) Utrophin-dystrophin-deficient mice as a model for
Duchenne muscular dystrophy. Cell 90:717–727
27. Rafael JA, Townsend ER, Squire SE, Potter AC, Chamberlain
JS, Davies KE (2000) Dystrophin and utrophin influence fibertype composition and post-synaptic membrane structure. Hum
Mol Genet 9:1357–1367
28. Deconinck N, Rafael JA, Beckers-Bleukx G, Kahn D,
Deconinck AE, Davies KE, Gillis JM (1998) Consequences
of the combined deficiency in dystrophin and utrophin on the
mechanical properties and myosin composition of some limb
and respiratory muscles of the mouse. Neuromuscul Disord
8:362–370
29. Rafael JA, Cox GA, Corrado K, Jung D, Campbell KP,
Chamberlain JS (1996) Forced expression of dystrophin
deletion constructs reveals structure–function correlations.
J Cell Biol 134:93–102
30. Spencer RF, Porter JD (2005) Biological organization of the
extraocular muscles. Prog Brain Res 151:43–80
31. Porter JD, Rafael JA, Ragusa RJ, Brueckner JK, Trickett JI,
Davies KE (1998) The sparing of extraocular muscle in
dystrophinopathy is lost in mice lacking utrophin and
dystrophin. J Cell Sci 111:1801–1811
32. Porter JD, Khanna S, Kaminski HJ, Rao JS, Merriam AP,
Richmonds CR, Leahy P, Li J, Guo W, Andrade FH (2002) A
chronic inflammatory response dominates the skeletal muscle
molecular signature in dystrophin-deficient mdx mice. Hum
Mol Genet 11:263–272
33. Friedman DJ, Nigro JM, Jakovcic S, Rabinowitz M, Umeda PK
(1984) J Biol Chem 259:6674–6680
34. Lyons GE, Schiaffino S, Sassoon D, Barton P, Buckingham M
(1990) Developmental regulation of myosin gene expression in
mouse cardiac muscle. J Cell Biol 111:2427–2436
35. Deconinck N, Tinsley J, De Backer F, Fisher R, Kahn D, Phelps
S, Davies K, Gillis JM (1997) Expression of truncated utrophin
leads to major functional improvements in dystrophin-deficient
muscles of mice. Nat Med 3:1216–1221
36. Matsushita T, Oyamada M, Fujimoto K, Yasuda Y, Masuda S,
Wada Y, Oka T, Takamatsu T (1999) Remodeling of cell–cell
and cell–extracellular matrix interactions at the border zone of
rat myocardial infarcts. Circ Res 85:1046–1055
37. Porter JD, Merriam AP, Leahy P, Gong B, Feuerman J, Cheng
G, Khanna S (2004) Temporal gene expression profiling of
dystrophin-deficient (mdx) mouse diaphragm identifies conserved and muscle group-specific mechanisms in the pathogenesis of muscular dystrophy. Hum Mol Genet 13:257–269
38. Porter JD, Merriam AP, Leahy P, Gong B, Khanna S (2003)
Dissection of temporal gene expression signatures of affected
and spared muscle groups in dystrophin-deficient (mdx) mice.
Hum Mol Genet 12:1813–1821
39. Sanford JL, Mays TA, Rafael-Fortney JA (2004) CASK and
Dlg form a PDZ protein complex at the mammalian neuromuscular junction. Muscle Nerve 30:164–171
40. Irizarry RA, Bolstad BM, Collin F, Cope LM, Hobbs B, Speed
TP (2003) Summaries of Affymetrix GeneChip probe level
data. Nucleic Acids Res 31:e15
41. Pette D, Staron R (2001) Transitions of muscle fiber phenotypic
profiles. Histochem Cell Biol 115:359–372
42. LaFramboise WA, Guthrie RD, Scalise D, Elborne V, Bombach
KL, Armanious CS, Magovern JA (2003) Effect of muscle
origin and phenotype on satellite cell muscle-specific gene
expression. J Mol Cell Cardiol 35:1307–1318
91
43. Lee KJ, Helfman DM, Schroer TA (1992) Myosin light chain-2
luciferase transgenic mice reveal distinct regulatory programs
for cardiac and skeletal muscle-specific expression of a single
contractile protein gene. Nature 359:244–246
44. Grady RM, Teng H, Nichol MC, Cunningham JC, Wilkinson
RS, Sanes JR (1997) Skeletal and cardiac myopathies in mice
lacking utrophin and dystrophin: a model for Duchenne
muscular dystrophy. Cell 90:729–738
45. Janney MJ, Allen RE, Cornelison DD (1994) Single-cell
analysis of regulatory gene expression in quiescent and
activated mouse skeletal muscle satellite cells. J Cell Physiol
159:379–385
46. Holt J, Easton J, Smith K, Goldspink DF, Smith CK, 2nd
(1994) Temporal expression of myogenic regulatory genes
during activation, proliferation, and differentiation of rat
skeletal muscle satellite cells. J Cell Physiol 161:49–54
47. Garry DJ, Hawke TJ (2001) Myogenic satellite cells: physiology to molecular biology. J Appl Physiol 91:534–551
48. Close R (1965) Force: velocity properties of mouse muscles.
Nature 206:718–719
49. Ng W, Grupp I, Subramaniam A, Robbins J (1991) Cardiac
myosin heavy chain mRNA expression and myocardial function in the mouse heart. Circ Res 68:1742–1750
50. Tajsharghi H, Thornell LE, Lindberg C, Lindvall B, Henriksson
KG, Oldfors A (2003) Myosin storage myopathy associated
with a heterozygous missense mutation in MYH7. Ann Neurol
54:494–500
51. Perrot A, Schmidt-Traub H, Hoffmann B, Prager M, BitAvragim N, Rudenko RI, Usupbaeva DA, Kabaeva Z, Imanov
B, Mirrakhimov MM et al (2005) Prevalence of cardiac betamyosin heavy chain gene mutations in patients with hypertrophic cardiomyopathy. J Mol Med 83:468–477
52. Villard E, Duboscq-Bidot L, Charron P, Benaiche A, Conraads
V, Sylvius N, Komajda M (2005) Mutation screening in dilated
cardiomyopathy: prominent role of the beta myosin heavy
chain gene. Eur Heart J 26:794–803