Articles in PresS. Am J Physiol Cell Physiol (July 2, 2003). 10.1152/ajpcell.00128.2003
Nerve activity-independent regulation of skeletal muscle atrophy: role of MyoD and
Myogenin in satellite cells and myonuclei
Jon-Philippe K. Hyatt1, Roland R. Roy2, Kenneth M. Baldwin3, V. Reggie Edgerton1,2
1
Department of Physiological Science, 2Brain Research Institute, 621 Charles E. Young
Dr., University of California, Los Angeles, CA 90095, 3Department of Physiology and
Biophysics, D352 Medical Sciences 1, University of California, Irvine, 92697
Running head: Activity-independent modulation of muscle plasticity
Address correspondence to:
V. Reggie Edgerton, Ph.D.
Department of Physiological Science
621 Charles E. Young Dr.
University of California, Los Angeles
Los Angeles, CA 90095
Phone: (310) 825-1910
Fax: (310) 267-2071
e-mail: [email protected]
Submitted to American Journal of Physiology: Cell on March 28, 2003.
Key words:
Myogenic Regulatory Factor; denervation; spinal cord isolation,
bromodeoxyuridine
Copyright (c) 2003 by the American Physiological Society.
ABSTRACT
Electrical activity is thought to be the primary neural stimulus regulating muscle
mass, expression of Myogenic Regulatory Factor (MRF) genes, and cellular activity
within skeletal muscle. However, the relative contribution of neural influences that are
activity-dependent and -independent in modulating these characteristics is unclear.
Comparisons of denervation (no neural influence) and spinal cord isolation (SI, neural
influence with minimal activity) after 3, 14, and 28 days of treatment were used to
demonstrate whether there are neural influences on muscle that are activity independent.
Furthermore, the effects of these manipulations were compared for a fast ankle extensor
(medial gastrocnemius, MG) and a fast ankle flexor (tibialis anterior, TA). The mass of
both muscles plateaued at ~60% of control 2 weeks post-SI, whereas both muscles
progressively atrophied to <25% of initial mass at this same time point after denervation.
A rapid increase in Myogenin and, to a lesser extent MyoD, RNAs and proteins was
observed in denervated and SI muscles: at the later time points, these MRFs remained
elevated in denervated, but not SI muscles. This widespread neural activity-independent
influence on MyoD and Myogenin expression was observed in both myonuclei and
satellite cells and was not specific for fast or slow fiber phenotypes. Mitotic activity of
satellite and connective tissue cells also was consistently lower in SI than denervated
muscles. These results demonstrate a neural effect independent of electrical activity that
1) helps preserve muscle mass, 2) regulates muscle-specific genes, and 3) potentially
spares the satellite cell pool in inactive muscles.
2
INTRODUCTION
Skeletal muscle size, phenotype, and composition are regulated, in part, by the
nervous system. Eliminating neurally-induced electrical activity to skeletal muscles via
peripheral nerve axotomy (denervation) triggers rapid atrophy and augments the
expression of muscle-specific genes, notably Myogenic Regulatory Factors (MRFs) (2;
18; 29; 52; 53; 55), type II (fast) myosin heavy chain (MHC) isoforms (24; 35), and the
alpha subunit of the acetylcholine receptor (αAChR) (2; 18; 29; 37; 54). Furthermore,
after denervation both satellite cell proliferation and differentiation are enhanced (3; 16;
33; 36; 48). These observations are consistent with the current idea that electrical activity
is the primary neural stimulus modulating skeletal muscle plasticity.
MyoD and Myogenin proteins are basic helix-loop-helix transcription factors
localized within muscle-specific nuclei. These MRFs are traditionally thought to be
markers of skeletal muscle growth and hypertrophy since they can modulate satellite cell
division and their incorporation as new nuclei within mature muscle fibers. Following
activation, satellite cells may undergo one or several rounds of proliferation (47) during
which MyoD is expressed. Prior to and just following differentiation Myogenin is upregulated in satellite cell nuclei. Differentiation occurs when the satellite cell exits the
cell cycle and fuses either 1) to the parent fiber that it is associated with, thereby adding
to the nuclear population of the fiber, or 2) with other satellite cells to form a new fiber.
Denervation of a skeletal muscle enhances satellite cell activity, although it is
unclear whether this is an immediate or delayed response. For example, increased
satellite cell division in the rat extensor digitorum longus muscle can occur as early as 48
hours (36), or not until 30 days (48) after denervation. In addition, recent findings have
3
shown that satellite cell numbers diminish with prolonged denervation (>2 years),
presumably resulting from an increased rate of differentiation (3; 16).
Denervation eliminates neurally-mediated, activation-induced modulation of the
skeletal muscle properties. In contrast, spinal cord isolation (SI) eliminates all ascending,
descending, and peripheral neural input to the skeletal muscles, but leaves the
motoneuron-muscle connectivity intact (Fig. 1). In this preparation, the spinal cord is
transected at a mid-thoracic and a high sacral level (Fig. 1a) followed by bilateral dorsal
rhizotomy between the transection sites (Fig. 1b). These procedures effectively isolate
the lumbar region of the spinal cord from activity-dependent, but not activityindependent, influences. Assuming that activity is the only means of mediating a neural
influence on muscle, then we would hypothesize that the effects of SI and denervation on
muscle mass, the levels of MRF expression, and satellite cell activity should be similar.
In general, the results do not support this hypothesis and indicate that there is a timedependent neural activity-independent influence on each of these parameters.
4
METHODS
Animals and surgical procedures
The experimental and animal care procedures used in this study were approved
under the guidelines outlined by the University of California, Los Angeles Animal
Research Committee and the American Physiological Society. Adult female SpragueDawley rats (200-235 g body weight) were assigned randomly to a control (n = 5/time
point), denervation (n = 6/time point), or SI (n = 7/time point) group. Rats from each
group were examined at 3, 14, or 28 days post-surgery. For all surgeries, the animals
were deeply anesthetized with a combination of ketamine hydrochloride (70 mg/kg body
weight) and acepromazine maleate (5 mg/kg body weight) administered intraperitoneally.
Denervation involved removing a 5-10 mm segment of the sciatic nerve in the thigh
region bilaterally. The severed proximal end was ligated and then sutured into
surrounding denervated muscles to prevent reinnervation of the distal musculature. The
details of the SI procedures have been reported previously (20; 43). Briefly, the spinal
cord was transected completely at a mid-thoracic and an upper sacral level and the dorsal
roots cut bilaterally between the two transection sites (Fig. 1a). For each rat, miniosmotic pumps (Type 2ML2, Alzet Inc., Cupertino, CA) filled with a 3% solution of
bromodeoxyuridine (BrdU) were implanted subcutaneously in the mid-back region. The
pumps were immersed in 0.9% sterile saline for two hours prior to implantation. The
pumps were inserted either on the day of surgery (3- and 14-day groups) or on day 14
(28-day group). Following surgery, all incisions were closed and the animals were
allowed to recover in a heated (37°C) incubator. SI animals were monitored closely for
the duration of the study: the urinary bladder was expressed manually three times per day
5
for the first week following surgery, then twice per day thereafter. The hindlimbs in the
SI rats were manipulated passively through a full range of motion once per day to prevent
adverse conditions associated with inactivity such as ankylosis.
Motor tests were
performed routinely to verify that the muscles in the hindlimbs of the SI rats were
inactive, i.e., there was an absence of a toe spreading reflex and of a withdrawal reflex to
limb extension and toe pinching, respectively. No SI rats showed any response to these
motor reflex tests throughout the study. Occasional movement in the hindlimbs of SI rats
was observed during daily bladder expression, most likely due to a mechanical
stimulation of the spinal cord. Based on previous reports (23; 37), however, we conclude
that this brief activation was not sufficient to blunt the elevated MRF expression in SI
muscles since MyoD and Myogenin can remain elevated after continuous exogenous
electrical stimulation for several minutes (23) or hours (37).
Our procedures for
maintaining spinally injured animals have been reported in detail previously (42).
On the day of termination, the proximal stump of the sciatic nerve of each
denervated rat was stimulated via bipolar silver electrodes (1-10 volts, range of frequency
from 1-100 Hz for 300 ms) to ensure that reinnervation had not occurred in the distal
musculature. Complete denervation was verified in each animal in this study. The TA
and MG muscles were dissected from both hindlimbs of each animal, cleaned of excess
connective tissue, and wet weighed. The mean of the two muscles from each rat was
recorded. The muscle masses for the TA and MG in each group then were averaged and
compared. Each muscle was pinned on a cork at approximately the in situ length and
submerged in isopentane cooled by liquid nitrogen. The muscles were stored at -70°C
until further analysis.
6
RNA extraction and RT-PCR analysis
Total RNA from muscle homogenates of randomly selected ipsi- or contralateral
limbs was extracted according to the manufacturer’s protocol (Molecular Research
Center, Inc., Cincinnati, OH) and converted to cDNA as previously described (17). For
Myogenin
amplification,
the
ACTACCCACCGTCCATTCAC-3’
following
(5’
primers
sense
were
primer)
used:
5’-
and
5’-
TCGGGGCACTCACTGTCTCT-3’ (3’ antisense primer) and yielded a 233-bp product.
For
MyoD
amplification,
the
CTACAGCGGCGACTCAGACG-3’
following
(5’
primers
sense
were
primer)
used:
5’-
and
5’-
TTGGGGCCGGATGTAGGA-3’ (3’ antisense primer) and yielded a 563-bp product.
Alternate 18S internal standards were used (Ambion, Inc., Austin, TX) and yielded a
324-bp product. The 18S competimers and primers were mixed in an 8:1 ratio. A 1 µl of
cDNA solution was added to 19 µl (1x PCR buffer, 0.2 mM dNTP, 2 mM MgCl 2, 18S
primer mix, and 0.75 units of Taq DNA polymerase (InVitrogen, Inc., Carlsbad, CA).
Amplification was performed using a Stratagene thermocycler (Stratagene, Inc., La Jolla,
CA) which commenced with a denaturing step (96°C for 4 min) followed by 1 min at
96°C, 45 sec at 55°C, and 45 sec at 72°C. Myogenin and MyoD were amplified for 25
and 26 cycles, respectively. A final elongation step was performed at 72°C for 3 min.
Each sample then was electrophoresed in a 2% agarose gel containing ethidium bromide.
The obtained film negatives were analyzed by laser-scanned densitometry and quantified
as previously described (56). Each sample was run in duplicate, normalized to the 18S
subunit, averaged, and statistically compared.
7
Protein extraction and Western analysis
For MyoD and Myogenin immunoblotting, total muscle protein containing
cytoplasmic and nuclear fractions was extracted from randomly chosen ipsi- or
contralateral limbs by rapid homogenization of a pre-weighed frozen samples in 10volume of boiling lysis buffer (1% SDS, 10mM Tris pH 7.4, and 1 mM sodium
orthovanadate). After complete homogenization, the samples were boiled for 15 sec and
centrifuged for 10 min at 10,000 g. An aliquot of the supernatant was used for
determining protein concentration (using Bio-Rad, Inc., Hercules, CA, DC protein assay
reagent), and the remainder of the supernatant was stored at 4°C for subsequent Western
analysis. Optimal loading for immunoblotting was determined to be 90 and 75 µg per
sample for MyoD and Myogenin, respectively.
Four control standards were run
simultaneously with each gel: a biotin-conjugated molecular weight marker (Cell
Signaling, Inc., Beverly, MA); protein isolated from undifferentiated (highly expressing
MyoD) C2C12 cells; protein isolated from differentiated (24-hour serum starved; highly
expressing Myogenin) C2C12 cells; and protein isolated from neonatal (P7) rat plantaris
muscle (highly expressing both MRFs). Protein was denatured by boiling in SDS-PAGE
sample buffer (0.2% SDS, 20% glycerol, 25% 4x buffer, 5% β-mercaptoethanol, and
0.025% bromophenol blue) for 3 min and electrophoresed (30 mA) in either a SDS-10%
(MyoD) or -12.5% (Myogenin) polyacrylamide gel. The proteins were transferred to
nitrocellulose membranes for 3 h at 500 mA. The membranes were immersed in a
blocking solution containing 5% non-fat dry milk (Bio-Rad, Inc.) dissolved in Trisbuffered saline with 0.05% Tween-20 (T-TBS) for 1 h. The membranes then were
8
incubated in either mouse anti-MyoD (1:400) (Dako, Inc., Carpinteria, CA) or antiMyogenin (1:500) (Santa Cruz, Inc., Santa Cruz, CA) diluted in blocking solution
overnight at 4ºC. The membranes were washed 6 x 10 min in T-TBS and incubated for 1
h in a secondary antibody cocktail (anti-biotin, 1:5000; goat anti-mouse IgG, 1:4000 for
MyoD or 1:5000 for Myogenin; Santa Cruz, Inc.) at room temperature. The membranes
were developed using an ECL+ detection kit (Amersham, Inc., Piscataway, NJ) per the
manufacturer’s instructions. Densitometry and quantification were performed as
described above. MyoD and Myogenin proteins in control muscles were generally below
detectable levels and are not shown. Any detected protein in the control muscles was
averaged and then subtracted from each denervated or SI protein value at the
corresponding time point. For each PCR and Western analysis, at least one sample per
group per time point was performed for within-group-time and between-group
comparisons.
Immunohistochemistry
Between 2-4 cryosections (10 µm-thick) cut from the muscle belly were
rehydrated in phosphate-buffered saline (PBS), fixed in 4% paraformaldehyde, washed 2
x 10 min in PBS, and immersed in 5% normal donkey serum for 15 min. Satellite cells
were detected with an antibody for M-cadherin (a gift from A. Wernig, Bonn University),
which was generated from the methods described by Rose et al. (41) and has showed
specific labeling for quiescent and activated satellite cells (27). There are several studies
reporting in situ M-cadherin mRNA labeling in cultured Schwann cells and fibroblasts
(38) and M-cadherin protein localization in muscle regions, e.g., extracellular matrix, that
9
are inconsistent with satellite cell position (11). However, there are distinct
methodological discrepancies (38) and differences in M-cadherin antibody generation
(11) with these studies and the present one that may have attributed varied findings. To
test for M-cadherin antibody specificity, we initially incubated control, denervated, and
SI muscles from each time point with rabbit anti-M-cadherin for 1 h at room temperature
and then overnight at 4ºC. Following serial washes in PBS a donkey anti-rabbit IgGFITC secondary antibody (1:75 final dilution) (Jackson Labs, Inc., West Grove, PA) was
applied for 1 h at room temperature. The samples were washed in PBS, blocked in 5%
normal goat serum for 15 min., and then incubated with a mouse anti-laminin (1:50 final
dilution) (2E8 from Hybridoma bank, Iowa City, IA) for 1 h at room temperature. The
samples were washed in PBS and a goat anti-mouse IgG-RRX secondary antibody (1:75)
(Jackson Labs, Inc.) was applied for 1 h at room temperature. The muscle sections were
mounted in Vectashield mounting media (Vector, Inc., Burlingame, CA) containing
DAPI, a general tag for all nuclei. We observed M-cadherin-positive nuclei adjacent to
parent fibers inside the laminin-positive regions. No M-cadherin-positive nuclei were
observed in the extracellular matrix or, based on position and frequency, in Schwann
cells or fibroblasts (Fig. 6).
For MRF localization experiments, a primary antibody cocktail of mouse antiMyoD or -Myogenin (Dako, Inc.) and rabbit anti-M-cadherin, a satellite cell-specific
marker, was used. For satellite cell proliferation analysis, an antibody cocktail of mouse
anti-BrdU (Beckton Dickinson, Inc., San Diego, CA) and anti-M-cadherin was used. For
satellite cell differentiation experiments, an antibody cocktail of anti-BrdU and rabbit
anti-dystrophin (a gift from K. Campbell, University of Iowa) was used. Dystrophin
10
tagged the inner boundaries of each fiber in order to determine differentiated (BrdUpositive) satellite cells. The antibodies were diluted in a PBS solution containing 0.5%
carrageenan and 0.02% sodium azide at a 1:50 final dilution for all antibodies. For BrdU
staining, samples were pre-incubated in 2N HCl for 1 h at room temperature. The primary
antibody solutions were applied to the muscle sections for 1 h at room temperature and
then overnight at 4ºC. Following 3 x 10 min washes in PBS, a donkey anti-mouse IgGRRX secondary antibody (1:75) (Jackson Labs, Inc.) was administered followed by a
donkey anti-rabbit IgG-FITC secondary antibody (1:75) (Jackson Labs, Inc.) for 1 h at
room temperature. Samples were washed 3 x 10 min in PBS and mounted as described
above. Lastly, we determined whether MyoD and Myogenin proteins were associated
with Type I (slow) or Type II (fast) MHC muscle fibers. For these experiments, we
examined serial sections from the deep region (close to the bone, mixed fiber type
composition) of control, denervated, and SI MG muscles. Sections were fixed and
prepared as described above. On one section, a primary antibody cocktail of either antiMyoD or -Myogenin and rabbit anti-laminin (Sigma, Inc., St. Louis, MO) was diluted
1:50 and applied for 1 h at room temperature and then overnight at 4ºC. A primary
antibody mix of anti-laminin and mouse anti-slow MHC (Novacastra Laboratories, Ltd.,
Burlingame, CA) was applied to the other muscle cross section for 1 h at room
temperature. Laminin was tagged to identify the same fibers in serial sections. A donkey
anti-mouse IgG-RRX secondary antibody was used for MyoD, Myogenin, or slow MHC
detection (1:75 final dilution). A donkey anti-rabbit IgG-FITC secondary antibody was
administered for laminin detection (1:75 final dilution). The sections were mounted as
described above.
11
Muscle cross sections were analyzed using an Axiophot microscope (Carl-Zeiss,
Inc., Thornwood, NY). Using a KX Series Imaging system (CRI, Inc., Boston, MA)
individual scans were made for red, green, and blue wavelengths (3 s exposure for each
channel) emitted by the secondary antibodies and DAPI, respectively. False color
composite images were constructed on a Dell Optiplex GX1p (Dell Computer, Inc.,
Round Rock, TX) computer using Image Pro Plus (version 4.0) software (Sysantec, Inc.,
Santa Monica, CA). Nuclei labeled for MyoD, Myogenin, BrdU, and/or M-cadherin
were counted in randomly selected regions. Between 2-4 regions (region = ~0.3 mm2) for
each muscle section containing at least 25 contiguous fibers and free of damage, large
connective tissue areas, or large vessels were chosen for analysis. For random selection
of these regions, each muscle section was divided into 16 regions and labeled from 1-16.
A random number generator (Microsoft®) was used to determine the regions for analysis.
The myonuclei and satellite cells positive for MyoD, Myogenin, and/or BrdU were
counted in each region and averaged. Between 2-4 sections, i.e., a minimum of 200
fibers, were analyzed per muscle. The counts for each criterion measure were normalized
to the number of fibers per region to account for the muscle fiber atrophy associated with
SI and denervation.
Statistical analyses
All data are reported as mean ± SEM. Within-group-time and between-group
comparisons were performed using a two-way analysis of variance (ANOVA) and
Tukey’s post-hoc analyses. Statistical significance was set at P < 0.05.
12
RESULTS
Body and muscle masses
Mean body and muscle (absolute and relative) mass for each group at each time
point are shown in Table 1. The relative atrophy for each muscle on day 3 was similar in
the denervated and SI groups (Fig. 2). By 2 weeks post-surgery, the TA and MG muscle
masses in SI rats were ~64 and 71% of age-matched controls, respectively. Comparable
values were only ~48 and 46% for the denervation group. The difference in percent
atrophy between the denervated and SI muscles was even more disparate by 28 days: the
relative muscle masses in the SI and denervated rats plateaued at ~60% and 25% of
control, respectively.
MyoD and Myogenin mRNA and protein levels
The regulation of the MyoD gene was muscle-specific. Compared to the TA
muscles in control rats, MyoD mRNA was higher at 14 and 28 days in the denervation
and at 28 days in the SI groups with the largest increase occurring at 28 days (SI, ~1.8fold; denervation, ~2.6-fold; Fig. 3A). In the MG muscle, the highest MyoD mRNA
levels were observed at 3 days following SI (~13-fold) and denervation (~11-fold; Fig.
3B). These levels remained elevated in the denervation, but not SI, group at 14 and 28
days. In addition, the MyoD mRNA levels were lower in the SI than denervated rats at
these later time points. Compared to control, the Myogenin mRNA levels increased ~22and 28-fold in the SI and denervated TA muscles 3 days post-surgery, respectively (Fig.
3C). At the same time point, Myogenin mRNA was elevated in the SI (~6-fold) and
denervated (~9-fold) MG muscles (Fig. 3D).
13
The Myogenin mRNA levels in the
denervated group remained elevated in both muscles at the 14 and 28 day time points. In
contrast, there was a progressive decrease in this response in the SI group, such that the
values were similar to control at the latest time point. In effect, the values were higher in
the denervated than SI rats at all time points in both muscles.
Western analysis showed a progressive increase in MyoD protein in the TA and
MG muscles of the denervated rats with the levels being higher at 28 than 3 and 14 days
(Fig. 4A and B). There were no significant changes in MyoD protein in either muscle of
the SI group throughout the study. Myogenin protein levels generally paralleled the
changes in its mRNA in both the SI and denervation groups (compare Fig. 3C and D with
Fig. 4C and D). These levels in both muscles were higher in the denervated than SI rats at
all time points. Over the 28-day period, the Myogenin protein levels in the TA and MG
muscles were unaffected by denervation, whereas there was a progressive decrease in SI
rats such that the values at 3 days were higher than at 14 and 28 days. MRF proteins were
ubiquitous in muscle nuclei, i.e., both satellite cells and muscle fiber nuclei (myonuclei),
and were not specific to either fast or slow fiber phenotypes in denervated (Fig. 5) and SI
muscles (not shown).
Localization of MyoD and Myogenin proteins
Co-labeling with M-cadherin and either MyoD (Fig. 7A and B) or Myogenin (Fig.
7C and D) revealed that the nuclei containing either MRF occurred predominantly within
myonuclei rather than satellite cells. The frequency of MRF-positive myonuclei and, to a
lesser extent, satellite cells reflected the relative changes in MRF expression as
determined by Western analysis (Fig. 4). For instance, relatively low MyoD or Myogenin
14
protein levels in SI muscles at 28 days (Fig. 4) were associated with less frequent and
intense MRF staining in the muscle cross sections (Fig. 7). In control rats the presence of
MyoD and Myogenin proteins in the TA and MG muscles (Fig. 7A and B) was infrequent
and, when expressed, was associated largely with satellite cells.
Satellite cell proliferation and differentiation
Some satellite cell-specific expression of MyoD and Myogenin was observed in
both muscles of denervated and SI rats (Fig. 7), suggesting that a portion of the satellite
cell pool was activated in both conditions. Determining whether this level of activation
was comparable to control conditions was not possible because these proteins are
transiently expressed (57) and the time points used provide only “snap-shots” of activated
satellite cells. Thus, we examined the mitotic activity of satellite cells and connective
tissue cells by continuous perfusion of BrdU for either 3 or 14 days.
BrdU, a thymidine analog, is selectively incorporated into the DNA of dividing
cells or in the repairing of DNA. Thus, the time course of the BrdU delivery used in the
present study was used to determine whether satellite cell proliferation or differentiation
was an immediate (3 or 14 days) or delayed (28 days) response. Despite considerable
exposure to BrdU ( 2 weeks), previous evidence by Schmalbruch and Lewis
(45)
indicated that BrdU for this duration does not produce toxic effects in vivo. In general,
satellite cell (Fig. 8) and connective tissue cell (Fig. 9) division was greater at 14 than 28
days (p>0.05 for satellite cells). At both of these time points the mitotic activity in both
muscles was consistently higher in the denervated than SI rats. Compared to control,
satellite cell division in the TA of SI rats had increased by 14 days and then returned to
15
control levels by 28 days, whereas these values in the denervated rats were higher than
control at both time points (Fig. 8A). Satellite cell proliferation in the MG muscles of SI
rats was similar to control levels at all time points, but elevated in the denervated MG
muscles at 14 and 28 days (Fig. 8B).
Satellite cell differentiation in both muscles was higher in denervated than SI rats
at both 14 and 28 days (Fig. 10). The majority of differentiated satellite cells were located
at the periphery of the fibers in both conditions; however, when present, centrally located
BrdU-labeled nuclei were observed more frequently in denervated than SI muscles (Fig.
10, top panel). On occasion, evidence for the formation of new fibers was detected in
denervated and, to a lesser extent, SI muscles (data not shown).
16
DISCUSSION
Muscle-derived biochemicals can affect neuronal plasticity, namely elongation
(22), and post-synaptic cell targeting (9) and even survival (51). The major findings in the
present study suggest that neural factors also affect post-synaptic targets at the whole
tissue, molecular, and cellular levels independent of electrical activity. In skeletal muscle,
these influences are sufficient to blunt atrophy, MyoD and Myogenin expression, and
cellular activity, particularly at extended periods.
At the whole tissue level, the progressive muscle atrophy was less in SI than
denervated rats at 2 and 4 weeks. This relative preservation of muscle mass is consistent
with earlier observations in SI (20) and tetrodotoxin (TTX)-treated rats (13). Both the SI
and TTX models block neurally-mediated activation of the hindlimb muscles while
maintaining the muscle-nerve connectivity. TTX is a sodium channel blocker that can
eliminate the propagation of action potentials distal to the site of application to the
peripheral nerve. In this case, the motoneurons presumably remain normally active. In
contrast, SI inactivates the hindlimb muscles at the level of the spinal cord, thus resulting
in the motoneuron, as well as the associated musculature, being electrically silent.
The blunted atrophic response compared to denervation observed in both the SI
and TTX models suggests a similar neurally-mediated activity-independent effect on the
hindlimb muscles. In addition, this activity-independent modulation of muscle plasticity
is supported by the observation that the upregulation of Myogenin mRNA in the rat
triceps surae was ~50% less after 7 days of TTX treatment than denervation (55). Taken
together, these studies suggest that the observed differences between SI and denervation
17
in the present study cannot be attributed to a greater level of residual activation of the SI
muscles.
It has been proposed that MyoD and Myogenin genes are tightly regulated by
electrical activity and serve as intermediaries between electrical activity and the
expression of other muscle-specific genes, such as fast MHC isoforms and the αAChR
subunit. This notion is supported by evidence from cross-reinnervation experiments (25)
and studies showing that exogenous electrical stimulation normalizes the elevated MyoD,
Myogenin, and αAChR mRNA levels in denervated muscle (18; 37). In light of these
earlier reports (18; 37), the early (Day 3) rise in MyoD and Myogenin mRNA (Fig. 3)
suggests an initial response to the cessation of electrical stimuli in SI and denervated
muscle, since both models reduce activation of the hindlimb muscles. The
downregulation of Myogenin mRNA at 2 (<7-fold) and 4 weeks (<3-fold) post-SI,
however, does not support the theory that electrical activity is the principal neural
stimulus regulating this muscle-specific gene. This compares to an ~10-15 fold increase
of Myogenin transcripts one month after denervation (Figs. 3C and D). The blunted
response of Myogenin and, to a lesser degree, MyoD mRNA in SI muscles is consistent
with a neurally-mediated, activity-independent regulation of these muscle-specific genes.
Myogenin protein also was highly up-regulated in denervated, but not in SI,
muscles. The increase in Myogenin protein following denervation is consistent with the
findings of Kostrominova et al. (29), who observed a dramatic increase in Myogenin
protein in the rat TA and gastrocnemius muscles after 3, 10, and 30 days of deneravtion.
In the present study, MyoD protein was highly up-regulated in denervated, but not SI,
muscles, with the greatest differences observed at 28 days. The differential expression of
18
MyoD mRNA (Fig. 3) and protein (Fig. 4) in SI muscles suggests that both
pretranslational and translational processes were being affected by activity-independent
influences. It is possible that the translational proteins also were upregulated in
denervated but not SI muscles, resulting in a greater conversion of MyoD and Myogenin
mRNAs to their respective proteins. Additionally, we cannot discount the possibility that
mRNA or protein degradation occurred differently in SI and denervated muscle. Previous
work has shown that a number of degradation pathways are enhanced following
denervation (5; 19; 28; 34). We are currently investigating the expression of several
degradation pathway-related genes (Atrogin-1, MuRF-1, poly-ubiquitin), which may help
to explain the MRF expression differences observed in SI and denervated muscles.
Combined, these results indicate that the expression of these muscle-specific genes is
modulated by activity-dependent and -independent mechanisms. Only at 14- and 28-day
time points do the effects of activity-independent influences become clearly differentiated
from activity-dependent mechanisms (Figs. 3 and 4).
Interestingly, Maier et al. (32) detected no MyoD mRNA or protein in denervated
TA and soleus rat muscle at 2, 7 or 28 days. Sakuma and coworkers (44) reported a
gradual decrease in MyoD protein in denervated plantaris and gastrocnemius muscles 128 days post-surgery, whereas Myogenin protein was relatively unchanged in these
muscles. These findings contrast with other studies showing an increase in these MRFs
following denervation (2; 8; 18; 29; 52; 55). It is unknown why such MRF expression
differences exist with denervation. It is possible that that the species of rat used in each
study exhibit different MRF responses: Maier et al. and Sakuma et al. used Wistar rats,
19
whereas we and others (18; 52; 53; 55) used Sprague-Dawley rats, although augmented
MRF expression has been reported in denervated muscles of Wistar rats (2; 10).
There are a number of candidate molecules that could mediate the apparent
neurotrophic effects that were observed in the present study, although such biochemicals
may not be nerve-derived. We did not find MRF protein expression to be localized at the
NMJ. Similarly, Witzemann and Sakmann (55) found MyoD and Myogenin mRNA
expression at both junctional and extra-junctional regions of 7-day denervated rat
diaphragm muscles. To this effect, the neurotrophic influence could emanate from local
Schwann cells or fibroblasts, which also can generate neurotrophic signals (1; 4).
Compelling evidence by Helgren et al. (21) showed that delivery of exogenous
recombinant ciliary neurotrophic factor (CNTF) to denervated rat soleus muscles
attenuated the atrophic response and blunted the slow-to-fast MHC (I to IIa) conversion
during the first two weeks after surgery. CNTF is produced abundantly within the
peripheral nervous system by Schwann cells and the CNTF receptor is expressed in
skeletal muscle (15). There is a dramatic drop in CNTF mRNA within the first week after
denervation (14; 26). Therefore, if CNTF-mediated signaling suppresses MyoD or
Myogenin expression, then these MRFs would remain more elevated in denervated than
SI
muscles,
as
observed
in
the
present
study.
Although
CNTF
inhibits
acetylcholinesterase production in 7-day denervated rat soleus muscles (7), there is no
current evidence that CNTF can similarly block MRF expression. Furthermore, we
cannot exclude the possibility that the presence of other neurotrophins, such as brainderived neurotrophic factor (1) or nerve growth factor (4), also could have had activity-
20
independent effects as cellular constituents within muscle tissue also produce these
neurotrophins.
Several groups have reported a widespread MRF response in denervated muscle
nuclei (29; 52; 53) using immunohistochemical analyses. In these studies the proportion
of satellite cells and myonuclei expressing MyoD or Myogenin was not ascertained as no
specific marker for satellite cells was used. Using an antibody for M-cadherin, Weis et al.
(53) showed that MRF4, another member of the MRF family of genes, increased in
satellite cells and myonuclei in denervated rat diaphragm muscle, but there was no
determination in which nuclei the MRF4 proteins predominately localized. In the present
study, we hypothesized that MyoD and Myogenin would be associated primarily with
satellite cells based on earlier reports of increased satellite cell activity in denervated
muscles (33; 36; 48), that myonuclei are post-mitotic, and that the current view is that
MRF proteins play a predominate role in regeneration rather than adaptation in adult
skeletal muscle. Our findings show that MRF proteins accumulate predominantly within
myonuclei rather than satellite cells (Fig. 7).
These findings were unexpected
considering that the presence of MyoD and Myogenin in myonuclei is thought to be only
a short-term response during satellite cell proliferation and differentiation (57). Our
results show, however, that at each time point the MRF proteins were consistently upregulated largely in myonuclei. Identification of satellite cells with M-cadherin has been
used to detect satellite cells using light microscopy (6; 12; 27; 30; 40; 53). It has been
suggested that M-cadherin labels only differentiating (Myogenin-M-cadherin co-labeled)
satellite cells (30; 32). We observed that M-cadherin-positive satellite cells also express
MyoD, a MRF associated with proliferating myogenic cells. Cooper et al. (12) also
21
showed M-cadherin co-expressed with MRFs specific for proliferating satellite cells
(MyoD and Myf5) in regenerating mouse soleus and gastrocnemius muscle, confirming
that M-cadherin can be expressed in non-differentiating satellite cells. Kuschel et al. (30)
showed virtually no Myogenin protein co-labeled with M-cadherin in cultured denervated
flexor digitorum brevis muscle fibers of the rat (30), which support our findings that, in
fact, MRFs are expressed predominantly in myonuclei rather than satellite cells following
denervation. This widespread up-regulation in myonuclei suggests that MyoD and
Myogenin are inducing structural and phenotypic adaptations of the denervated and SI
muscle fibers. Previous studies have shown that MyoD and Myogenin can interact with
promoter elements of the MHC IIb (49; 50), αAChR (39), and fast troponin I (31) genes
suggesting that the early up-regulation of these MRFs may be contributing to the
increased expression of these genes in denervated and SI muscles. However, regulation of
these genes clearly requires a host of transcriptional proteins as the presence of MRFs
alone has been previously shown to be insufficient to activate gene expression (31).
A neural activity-independent influence also was evident following analysis of
cellular activity. Generally connective tissue cell, as well as satellite cell, activity was
lower in SI than denervated muscles (Figs. 8-10). Augmented cellular proliferation (36;
46; 48) and satellite cell differentiation following denervation have been observed
previously (3; 16). A regenerative-like response in denervated muscles has been
attributed to initiate the formation of new fibers, presumably from differentiated satellite
cells (46) that have migrated through the basal lamina (away from the parent fiber) into
the connective tissue spaces. While the possibility exists that migrating satellite cells also
differentiated and helped to form new fibers rather than fusing with the parent fiber, our
22
analyses focused on satellite activity, e.g., BrdU incorporation, in denervated and SI
muscles as a whole. We did not focus on the dynamics of individually migrating satellite
cells.
In addition, we do not believe turnover rate of myonuclei, presumably via
apoptosis, confounded our analyses, as previous evidence showed that not only is the
normal myonuclear turnover rate slow (45), but the presence of apoptotic indicators was
infrequent during the first several weeks following denervation (45; 58). The lower
satellite cell activity observed in SI muscles is consistent with McGeachie’s proposal (33)
that muscle-nerve contact suppresses satellite cell activity.
Given that the presence of
the nerve in the absence of activation is sufficient to suppress satellite cell proliferation
and differentiation, it appears that some neurotrophic factor(s) have the prospect of
preserving the satellite cell pool as well as preserving the cytoplasmic components of the
muscle fibers.
Summary
A more normal level of homeostasis was observed in muscle fibers having
continuity with their motoneurons, albeit inactive, compared to muscle fibers with no
anatomical or functional continuity with their motoneurons. The TA and MG muscles,
which differ functionally, morphologically, and biochemically, responded similarly after
SI surgery: atrophy plateaued at ~60% of control muscle mass at 14 days at which time
there was a general return of MRF mRNA and, to a greater extent, protein levels to
control values when compared to denervation. In addition, after the initial 2-week period
satellite cell activity returned to basal levels in SI muscles (see 28 day time point in Figs.
8 and 10). These results highlight two important points. Firstly, physical contact and the
23
ensuing biochemical communication between the nerve and muscle is a key component
in the homeostatic process for skeletal muscle. Secondly, the role of MRFs in adult
muscle is potentially more important in orchestrating an adaptive response of existing
muscle fibers than in the regenerative responses of muscle fibers.
24
Acknowledgements
This work was supported by grants from NIH (NS16333) to VRE and the Sigma
Xi Society to JPKH. We would like to thank K. Campbell for providing us with antidystrophin and A. Wernig for providing us anti-M-cadherin. We also thank Hui Zhong,
Fadia Haddad, Anqi Qin, Ming Zeng, and Gary McCall for their technical assistance. The
SI illustrations in Figure 1 were reprinted by permission from Fairman Studios, Inc.
(Waltham, MA). Lastly, we especially thank Maynor Herrera for animal care and surgery
assistance.
25
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30
Table 1. Body mass and absolute and relative muscle masses for Control (CON), SI, and Denervated (DEN) groups.
Body Mass
(g)
CON
(n=5)
Day 3
DEN
(n=6)
SI
(n=7)
Start 214±3
End 229±4
218±1
224±4
Absolute
TA 475±13 399±12 ab
Muscle Mass
(mg)
MG 620±18 a 530±16 ab
CON
(n=5)
Day 14
DEN
(n=6)
SI
(n=7)
232±0.6 #*
205±2 #*
223±5
254±6
220±3
235±5
386±6 ab
512±13
496±9 ab
666±18
Relative
TA 2.1±0.03 1.8±0.05 ab 1.9±0.01ab
Muscle Mass
(mg / g)
MG 2.7±0.05 2.4±0.06 ab 2.4±0.04 ab
CON
(n=5)
Day 28
DEN
(n=6)
SI
(n=7)
224±3
184±8 #*
224±4
262±7
222±3
251±4
222±3
207±7 #*
226±9 a
236±26
546±18
143±4
259±17*
279±11 a
339±24
722±33
169±12
375±21*
2.0±0.04 0.96±0.04 a 1.3±0.01*
2.1±0.03 0.57±0.02 1.3±0.08*
2.6±0.08 1.2±0.04 a
2.8±0.07 0.67±0.04 1.8±0.08*
1.9±0.01*
Table 1. Mean (±SEM) changes in body and muscle mass in Control (CON), denervated (DEN), and SI rats. To account for
changes in body mass, both the absolute and relative (mg / g) muscle masses are shown. For all time points, the absolute and relative
muscle masses in the DEN and SI groups are different from CON (P < 0.05). *, #, a, b: significantly different from DEN, CON, 28
days, and 14 days, respectively.
Figure legends
Figure 1. Schematic of the spinal cord isolation (SI) procedures showing the midthoracic and upper sacral transections of the rat spinal cord (A) and the intradural
bilateral dorsal rhizotomy (yellow) between the transection sites (B, T8-11 shown). To
access the dorsal roots (yellow), the spinous processes were removed, a narrow trough (12 mm) made along the vertebral column between the transection sites and the dura
opened longitudinally. The dorsal roots were cut near their point of entry into the cord
and near their exit through the vertebral column. These procedures eliminate all
ascending, descending and peripheral neural input to the neurons in the lumbar region of
the spinal cord and effectively silenced the muscles innervated by the motor pools located
in this region of the spinal cord. These illustrations were reprinted by permission from
Fairman Studios, Inc. (Waltham, MA).
Figure 2. Percent atrophy of the tibialis anterior (TA, A) and medial gastrocnemius (MG,
B) muscles after 3, 14 and 28 days of denervation (DEN) or spinal cord isolation (SI).
Muscle masses were first expressed relative to body weight and then averaged for each
group and are plotted relative to age-matched control values. Values are mean ± S.E.M.
*, a, b: significantly different from DEN, 28 days, and 14 days, respectively.
Figure 3. MyoD (A and B) and Myogenin (C and D) mRNA expression in the TA (A
and C) and MG (B and D) muscles after 3, 14, and 28 days of DEN or SI as determined
by RT-PCR analyses. Expression patterns for representative rats in each group and at
each time point are shown above the graphs The 18S mRNA subunit served as an internal
control for each sample. Values are mean ± S.E.M.
#, significantly different from
control, other symbols and abbreviations are the same as in Fig. 2.
Figure 4. MyoD (A and B) and Myogenin (C and D) protein levels in the TA (A and C)
and MG (B and D) muscles after 3, 14, and 28 days of DEN or SI as determined by
Western blot analyses. Expression patterns for representative rats in each group and at
each time point are shown above the graphs. Values are mean ± S.E.M.
MW,
molecular weight marker; Neo, 7-day neonatal plantaris muscle homogenate; C2C12+
and C2C12-, differentiated and undifferentiated cultured muscle cells, respectively. All
other symbols and abbreviations are the same as in Figs. 2 and 3.
Figure 5. Serial transverse sections (10 µm thick) from the deep region of a 3-day
denervated MG muscle. Top: All fibers are outlined by laminin (green) and the fibers
containing Type I (slow) myosin heavy chain (MHC) are identified (red). Nuclei were
tagged using DAPI (blue). Bottom: The slow MHC fibers identified in the top panel are
identified by asterisks (*). The location of MyoD protein is shown as red/pink staining in
muscle-specific nuclei (myonuclei and satellite cells) adjacent to both slow and fast fiber
types.
Figure 6. Cross-sections from control MG (A), denervated TA (B), and SI TA (C)
muscles (Day 3) showing specificity of the M-cadherin (green) antibody for satellite
cells. An antibody for laminin (red) was used to tag the boundaries of the extracellular
matrix. Nuclei were tagged using DAPI (blue). Scale bar = 50 µm. Insets in each panel
show an enlarged area containing the M-cadherin-positive nucleus. Panel B shows a Mcadherin positive nucleus (arrow) inside the laminin-positive region. In contrast, an
33
extracellular matrix-derived nucleus (arrowhead) is encapsulated by laminin and is
negative for M-cadherin.
Figure 7. Top, TA muscle (Day 14) sections labeled for M-cadherin (green), a satellite
cell-specific marker, and Myogenin (red/pink). Nuclei were tagged using DAPI (blue).
By intensifying the false color background muscle fibers were easily identified for
counting. Scale bar = 25 µm. Bottom, The frequency (%) of nuclei containing either
MyoD (A and B) or Myogenin (C and D) proteins from TA (A and C) and MG (B and D)
muscles was calculated as: [(1 – (MRF-positive satellite cells/MRF-positive nuclei) x
100)]. MRF-positive nuclei in this calculation included both satellite cells and myonuclei
expressing either MyoD or Myogenin. Mgn, Myogenin. Symbols and other abbreviations
are the same as in Figs. 2 and 3.
Figure 8. Top, MG muscle (Day 14) sections labeled for M-cadherin (green) and BrdU
(red/pink). Nuclei were tagged using DAPI (blue). BrdU was delivered for either 3 or 14
days using osmotic pumps. BrdU-tagged and unlabeled satellite cells were counted. All
other BrdU-tagged nuclei (predominantly in connective tissue cells) were counted (see
Fig. 9). Scale bar = 25 µm. Bottom, The frequency (%) of BrdU-positive satellite cells in
the TA (A) and MG (B) muscles was calculated as: [(BrdU-positive satellite cells/total
number of satellite cells) x 100]. Symbols and other abbreviations are the same as in Figs.
2 and 3.
Figure. 9. The number of BrdU-positive connective tissue cells per 100 fibers after 3, 14
and 28 days of denervation or SI is shown for the TA (A) and MG (B). All fibers and Mcadherin-negative cells (see Fig. 8) within each region of analysis (~0.3 mm2) were
34
counted and calculated as [BrdU-tagged cells/fiber number) x 100] to account for atrophy
in SI and denervated muscles. Symbols and other abbreviations are the same as in Figs. 2
and 3.
Figure 10. Top: Muscle cross sections (Day 14) showing differentiated satellite cells in
denervated and SI TA muscles identified as BrdU-positive nuclei (arrows, red/pink
staining) clearly within the muscle fiber boundaries (delineated by dystrophin, green).
Scale bar = 25 µm. Bottom: The number of BrdU-positive myonuclei per 100 fibers after
3, 14 and 28 days of denervation or SI is shown for the TA (A) and MG (B). All fibers
and BrdU-labeled myonuclei within each region of analysis (~0.3 mm2) were counted
and expressed as [BrdU-tagged myonuclei/fiber number) x 100] to account for the fiber
atrophy associated with SI and denervation. Symbols and other abbreviations are the
same as in Figs. 2 and 3.
35
A
B
Fig. 1
36
B
A
100
ab
ab
J
B
J
B
80
100
80
ab
*
*
ab
J
60
J
60
SI
B
a
40
B
DEN
B
20
20
0
3
7
10
14
18
Time (Days)
SI
B
a
40
*J
J
*
24
0
28
3
Fig. 2
37
7
10
14
18
Time (days)
24
28
DEN
A
B
TA
1.2
MG
1.5
ab
#
CON
#
1.2
1
a#
#
0.8
DEN
SI
Myo D
0.9
a#
#
0.6
#
0.6
0.4
*
0.3
0.2
0
0
3
C
14
Time (days)
28
3
14
Time (days)
D
TA
28
MG
ab
1.5
#
1.2
*
0.9
#
ab
#
ab
a#
ab
#*
0.6
#*
0.9
#*
0.6
Myogenin
#
0.3
0.3
#
a#
*
*
*
0
0
3
14
Time (days)
28
3
Fig. 3
38
14
Time (days)
28
A
B
TA
750
MG
750
DEN
SI
500
Myo D
500
a
250
250
a
*
*
ab
0
3
14
Time (days)
C
0
28
3
14
Time (days)
D
TA
400
300
300
ab
28
MG
400
200
*
Myogenin
200
*
100
*
*
*
100
*
0
*
0
3
14
Time (days)
28
3
Fig. 4
39
14
Time (days)
28
Fig. 5
40
Fig. 6
41
Fig. 7
42
A
ab
100
B
TA
# #
#
#
# #
MG
100
a#
#
#
#
b
75
75
#
#*
MyoD
50
50
25
25
0
0
3
C
14
Time (days)
28
D
TA
a # b#
100
# #
3
# #
14
Time (days)
28
MG
# #
#
#
# #
100
75
75
50
50
25
25
Myogenin
0
0
3
14
Time (days)
3
28
Fig. 7
43
14
Time (days)
28
A 50
B 50
TA
40
MG
40
#
#
#
30
30
b#
*
20
#
20
*
10
10
0
ab
*
*
0
3
14
Time (days)
28
3
Fig. 8
44
14
Time (days)
28
A 50
B 50
TA
a#
40
MG
40
a
#*
30
20
10
20
#
#*
ab
ab
a#
30
#
*
10
ab
*
ab ab ab
0
0
3
14
Time (Days)
3
28
Fig. 9
45
14
Time (days)
28
A
B 10
TA
7.5
#
#
MG
7.5
a#
5
5
#
*
*
2.5
*
2.5
*
ab ab
0
0
0
0
0
3
14
Time (Days)
3
28
Fig. 10
46
14
Time (days)
28