Nerve activity-independent regulation of skeletal muscle atrophy: role of MyoD and myogenin in satellite cells and myonuclei

Jon-Philippe K. Hyatt,1 Roland R. Roy,2 Kenneth M. Baldwin,3 and V. Reggie Edgerton1,2

1Department of Physiological Science and 2Brain Research Institute, University of California, Los Angeles 90095; and 3Department of Physiology and Biophysics, University of California, Irvine, California 92697

Submitted 3 April 2003 ; accepted in final form 21 June 2003


    ABSTRACT
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Electrical activity is thought to be the primary neural stimulus regulating muscle mass, expression of myogenic regulatory factor 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) and a fast ankle flexor (tibialis anterior). The mass of both muscles plateaued at ~60% of control 2 wk after 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 mRNAs and proteins was observed in denervated and SI muscles: at the later time points, these myogenic regulatory factors remained elevated in denervated, but not in SI, muscles. This widespread neural activity-independent influence on MyoD and myogenin expression was observed in 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 in 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.

myogenic regulatory factor; denervation; spinal cord isolation; bromodeoxyuridine


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 ({alpha}-AChR) (2, 18, 29, 37, 54). Furthermore, after denervation, satellite cell proliferation and differentiation are enhanced (3, 16, 33, 36, 48). These observations are consistent with the present 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, because they can modulate satellite cell division and their incorporation as new nuclei within mature muscle fibers. After activation, satellite cells may undergo one or several rounds of proliferation (47), during which MyoD is expressed. Before and just after differentiation, myogenin is upregulated in satellite cell nuclei. Differentiation occurs when the satellite cell exits the cell cycle and fuses 1) to the parent fiber, with which it is associated, 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 a delayed response. For example, increased satellite cell division in the rat extensor digitorum longus muscle can occur as early as 48 h (36) or not until 30 days (48) after denervation. In addition, recent findings have shown that satellite cell numbers diminish with prolonged denervation (>2 yr), presumably related to 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 midthoracic and a high sacral level (Fig. 1A) and then subjected to 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 activity-independent, influences. Assuming that activity is the only means of mediating a neural influence on muscle, 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 time-dependent neural activity-independent influence on each of these parameters.



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Fig. 1. Schematic illustration of spinal cord isolation (SI) procedures showing midthoracic and upper sacral transections of rat spinal cord (A) and intradural bilateral dorsal rhizotomy (yellow) between transection sites (B; T8-T11 shown). To access dorsal roots (yellow), spinous processes were removed, a narrow trough (1-2 mm) was made along the vertebral column between transection sites, and the dura was opened longitudinally. Dorsal roots were cut near their point of entry into the spinal cord and near their exit through the vertebral column. These procedures eliminate all ascending, descending, and peripheral neural input to neurons in the lumbar region of the spinal cord and effectively silence the muscles innervated by the motor pools located in this region of the spinal cord.

 


    METHODS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
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 Sprague-Dawley rats (200-235 g body wt) 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 after surgery. For all surgeries, the animals were deeply anesthetized with a combination of ketamine hydrochloride (70 mg/kg body wt) and acepromazine maleate (5 mg/kg body wt) administered intraperitoneally. Denervation involved removal of a 5- to 10-mm segment of the sciatic nerve in the thigh region bilaterally. The severed proximal end was ligated and then sutured to 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 midthoracic and an upper sacral level, and the dorsal roots were cut bilaterally between the two transection sites (Fig. 1A). For each rat, mini-osmotic pumps (type 2ML2, Alzet, Cupertino, CA) filled with a 3% solution of bromodeoxyuridine (BrdU) were implanted subcutaneously in the midback region. The pumps were immersed in 0.9% sterile saline for 2 h before implantation. The pumps were inserted on the day of surgery (for measurements on days 3 and 14) or on day 14 (for measurement on day 28). After 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 for the 1st wk after surgery and 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., a toe-spreading reflex and a withdrawal reflex to limb extension and toe pinching were absent. 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 because of mechanical stimulation of the spinal cord. On the basis of previous reports (23, 37), however, we conclude that this brief activation was not sufficient to blunt the elevated MRF expression in SI muscles, because 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 the animals were killed, the proximal stump of the sciatic nerve of each denervated rat was stimulated via bipolar silver electrodes (1-10 V, 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 the study. The tibialis anterior (TA) and medial gastrocnemius (MG) muscles were dissected from both hindlimbs of each animal and cleaned of excess connective tissue. Their wet weight was determined, and the mean of the two muscles from each rat was recorded. The masses for the TA and MG muscles 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.

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, Cincinnati, OH) and converted to cDNA as previously described (17). For myogenin amplification, the following primers were used: 5'-ACTACCCACCGTCCATTCAC-3' (5'-sense primer) and 5'-TCGGGGCACTCACTGTCTCT-3' (3'-antisense primer); they yielded a 233-bp product. For MyoD amplification, the following primers were used: 5'-CTACAGCGGCGACTCAGACG-3' (5'-sense primer) and 5'-TTGGGGCCGGATGTAGGA-3' (3'-antisense primer); they yielded a 563-bp product. Alternate 18S internal standards were used (Ambion, Austin, TX); they yielded a 324-bp product. The 18S competimers and primers were mixed in an 8:1 ratio. cDNA solution (1 µl) was added to 19 µl of 1x PCR buffer, 0.2 mM dNTP, 2 mM MgCl2, 18S primer mix, and 0.75 U of Taq DNA polymerase (InVitrogen, Carlsbad, CA). Amplification was performed using a thermocycler (Stratagene, La Jolla, CA): a denaturing step at 96°C for 4 min followed by 1 min at 96°C, 45 s at 55°C, and 45 s 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 was subjected to electrophoresis in a 2% agarose gel containing ethidium bromide. The 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.

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 preweighed frozen samples in 10 volumes of boiling lysis buffer (1% SDS, 10 mM Tris, pH 7.4, and 1 mM sodium orthovanadate). After complete homogenization, the samples were boiled for 15 s and centrifuged for 10 min at 10,000 g. An aliquot of the supernatant was used for determining protein concentration using DC protein assay reagent (Bio-Rad, Hercules, CA), 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/sample for MyoD and myogenin, respectively. Four control standards were run simultaneously with each gel: a biotin-conjugated molecular weight marker (Cell Signaling, Beverly, MA), protein isolated from undifferentiated (highly expressing MyoD) C2C12 cells, protein isolated from differentiated (24-h serum-starved, with high expression of myogenin) C2C12 cells, and protein isolated from neonatal (P7) rat plantaris muscle (with high expression of both MRFs). Protein was denatured by boiling in SDS-PAGE sample buffer (0.2% SDS, 20% glycerol, 25% 4x buffer, 5% {beta}-mercaptoethanol, and 0.025% bromphenol blue) for 3 min and subjected to electrophoresis (30 mA) in an SDS-10% (MyoD) or SDS-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% nonfat dry milk (Bio-Rad) dissolved in Tris-buffered saline with 0.05% Tween 20 for 1 h. The membranes were then incubated in mouse anti-MyoD (1:400; Dako, Carpinteria, CA) or anti-myogenin (1:500; Santa Cruz Biotechnology, Santa Cruz, CA) diluted in blocking solution overnight at 4°C. The membranes were washed six times for 10 min each in Tris-buffered saline with 0.05% Tween 20 and incubated for 1 h in a secondary antibody cocktail [anti-biotin (diluted 1:5,000) or goat anti-mouse IgG (diluted 1:4,000) for MyoD or goat anti-mouse IgG (diluted 1:5,000) for myogenin; Santa Cruz Biotechnology] at room temperature. The membranes were developed using an ECL+ detection kit (Amersham, Piscataway, NJ) according to 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 protein detected 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. Two to four cryosections (10 µm thick) cut from the muscle belly were rehydrated in phosphate-buffered saline (PBS), fixed in 4% paraformaldehyde, washed twice for 10 min each 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 shown 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 are inconsistent with satellite cell position (11). However, there are distinct methodological discrepancies (38) and differences in M-cadherin antibody generation (11) between these studies and the present study that may have resulted in 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 4°C. After serial washes in PBS, a donkey anti-rabbit IgG-FITC secondary antibody (1:75 final dilution; Jackson Labs, 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, Hybridoma Bank, Iowa City, IA) for 1 h at room temperature. The samples were washed in PBS, and a goat anti-mouse IgGRRX secondary antibody (1:75 dilution; Jackson Labs) was applied for 1 h at room temperature. The muscle sections were mounted in Vectashield mounting medium (Vector, Burlingame, CA) containing 4',6-diamidino-2-phenylindole, 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, on the basis of position and frequency, in Schwann cells or fibroblasts (Fig. 2).



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Fig. 2. Cross sections from control medial gastrocnemius (MG, A), denervated tibialis anterior (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 boundaries of the extracellular matrix. Nuclei were tagged using 4',6-diamidino-2-phenylindole (DAPI, blue). Scale bar, 50 µm. Insets: enlarged areas containing the M-cadherin-positive nucleus. B: M-cadherin-positive nucleus (arrow) inside the laminin-positive region. In contrast, an extracellular matrix-derived nucleus (arrowhead) is encapsulated by laminin and is negative for M-cadherin.

 

For MRF localization experiments, a primary antibody cocktail of mouse anti-MyoD or anti-myogenin (Dako) and rabbit anti-M-cadherin, a satellite cell-specific marker, was used. For satellite cell proliferation analysis, an antibody cocktail of mouse anti-BrdU (Becton Dickinson, 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 tagged the inner boundaries of each fiber for determination of differentiated (BrdU-positive) 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 preincubated in 2 N 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 4°C. After three washes for 10 min each in PBS, a donkey anti-mouse IgG-RRX secondary antibody (1:75 dilution; Jackson Labs) and then a donkey anti-rabbit IgGFITC secondary antibody (1:75 dilution; Jackson Labs) were administered for 1 h at room temperature. Samples were washed three times for 10 min each in PBS and mounted as described above. Finally, 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. A primary antibody cocktail of anti-MyoD or anti-myogenin and rabbit anti-laminin (Sigma, St. Louis, MO) was diluted 1:50 and applied to one section for 1 h at room temperature and then overnight at 4°C. A primary antibody mixture of anti-laminin and mouse anti-slow MHC (Novacastra Laboratories, 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.

Muscle cross sections were analyzed using an Axiophot microscope (Carl Zeiss, Thornwood, NY). A KX series imaging system (CRI, Boston, MA) was used to make individual scans for red, green, and blue wavelengths (3-s exposure for each channel) emitted by the secondary antibodies and 4',6-diamidino-2-phenylindole, respectively. False color composite images were constructed on a computer (Optiplex GX1p, Dell Computer, Round Rock, TX) using Image Pro Plus (version 4.0) software (Symantec, Santa Monica, CA). Nuclei labeled for MyoD, myogenin, BrdU, and/or M-cadherin were counted in randomly selected regions. Two to four regions (~0.3 mm2/region) for each muscle section that contained >=25 contiguous fibers and were 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 to 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. Two to four sections, i.e., >=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. Values are means ± SE. Within-group-time and between-group comparisons were performed using a two-way analysis of variance (ANOVA) and Tukey's post hoc analysis. Statistical significance was set at P < 0.05.


    RESULTS
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Body and muscle masses. Mean body and muscle (absolute and relative) masses 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. 3). By 2 wk after 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 day 28: the relative muscle masses in the SI and denervated rats was ~60 and 25% of control, respectively.


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Table 1. Body mass and absolute and relative muscle masses for control, SI, and denervated rats

 


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Fig. 3. Percent atrophy of TA (A) and MG (B) muscles after 3, 14, and 28 days of denervation (DEN) or 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 means ± SE. *Significantly different from DEN; asignificantly different from day 28; bsignificantly different from day 14.

 

MyoD and myogenin mRNA and protein levels. Regulation of the MyoD gene was muscle specific. In the TA muscle, MyoD mRNA was higher at days 14 and 28 in the denervation group and at day 28 in the SI group than in controls, with the largest increase at day 28 (~1.8- and ~2.6-fold in SI and denervation, respectively; Fig. 4A). In the MG muscle, the highest MyoD mRNA levels were observed 3 days after SI (~13-fold) and denervation (~11-fold; Fig. 4B). These levels remained elevated in the denervation, but not the SI, group at days 14 and 28. In addition, the MyoD mRNA levels were lower in SI than in denervated rats at these later time points. Compared with control, the myogenin mRNA levels increased ~22- and 28-fold in the SI and denervated TA muscles, respectively, 3 days after surgery (Fig. 4C). At the same time point, myogenin mRNA was elevated in the SI (~6-fold) and denervated (~9-fold) MG muscles (Fig. 4D). The myogenin mRNA levels in the denervated group remained elevated in both muscles at days 14 and 28. 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 denervated than in SI rats at all time points in both muscles.



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Fig. 4. MyoD (A and B) and myogenin (C and D) mRNA expression in TA (A and C) and MG (B and D) muscles after 3, 14, and 28 days of denervation or SI as determined by RT-PCR analyses. Top: expression patterns for representative rats in each group and at each time point. 18S mRNA subunit served as an internal control for each sample. D, denervation; C, control. Values are means ± SE. *Significantly different from DEN; asignificantly different from day 28; bsignificantly different from day 14; #significantly different from control.

 

Western analysis showed a progressive increase in MyoD protein in the TA and MG muscles of the denervated rats, with higher levels at day 28 than at days 3 and 14 (Fig. 5, A 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 the SI and denervation groups (compare Fig. 4, C and D, with Fig. 5, C and D). These levels in both muscles were higher in the denervated than in the 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 day 3 were higher than those at days 14 and 28. MRF proteins were ubiquitous in muscle nuclei, i.e., satellite cells and muscle fiber nuclei (myonuclei), and were not specific to fast or slow fiber phenotypes in denervated (Fig. 6) and SI muscles (not shown).



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Fig. 5. MyoD (A and B) and myogenin (C and D) protein levels in TA (A and C) and MG (B and D) muscles after 3, 14, and 28 days of denervation or SI as determined by Western blot analyses. Top: expression patterns for representative rats in each group and at each time point. Values are means ± SE. MW, molecular weight marker; Neo, 7-day neonatal plantaris muscle homogenate; C2C12+ and C2C12-, differentiated and undifferentiated cultured muscle cell homogenate, respectively. *Significantly different from DEN; asignificantly different from day 28; bsignificantly different from day 14.

 


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Fig. 6. Serial transverse sections (10 µm thick) from the deep region of a denervated MG muscle at day 3. A: all fibers are outlined by laminin (green), and fibers containing type I (slow) myosin heavy chain (MHC) are identified (red). Nuclei were tagged using DAPI (blue). B: slow MHC fibers identified in A (*). Location of MyoD protein is shown as red/pink staining in muscle-specific nuclei (myonuclei and satellite cells) adjacent to slow and fast fiber types.

 

Localization of MyoD and myogenin proteins. Colabeling with M-cadherin and either MyoD (Fig. 7, A and B) or myogenin antibodies (Fig. 7, C 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. 5). For instance, relatively low MyoD or myogenin protein levels in SI muscles at day 28 (Fig. 5) 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. 7, A and B) was infrequent and, when expressed, was associated largely with satellite cells.



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Fig. 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. A-D: frequency (%) of nuclei containing MyoD (A and B) or myogenin (C and D) proteins from TA (A and C) and MG (B and D) muscles calculated as follows: 1 - (MRF-positive satellite cells/MRF-positive nuclei) x 100. MRF-positive nuclei include satellite cells and myonuclei expressing MyoD or myogenin. Mgn, myogenin. Open bars, control; solid bars, denervation; shaded bars, SI. *Significantly different from denervation; asignificantly different from day 28; bsignificantly different from day 14; #significantly different from control.

 

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. It was not possible to determine whether this level of activation was comparable to control conditions, because these proteins are transiently expressed (57), and the time points used provide only "snapshots" of activated satellite cells. Thus we examined the mitotic activity of satellite cells and connective tissue cells by continuous perfusion of BrdU for 3 or 14 days.

BrdU, a thymidine analog, is selectively incorporated into the DNA of dividing cells or in the repair 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 (day 3 or 14) or delayed (day 28) response. Despite considerable exposure to BrdU (<=2 wk), 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 day 14 than at day 28 (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 in the SI rats. Compared with control, satellite cell division in the TA of SI rats had increased by day 14 and then returned to control levels by day 28, 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 was elevated in the denervated MG muscles at days 14 and 28 (Fig. 8B).



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Fig. 8. Top: MG muscle (day 14) sections labeled for M-cadherin (green) and bromodeoxyuridine (BrdU, red/pink). Nuclei were tagged using DAPI (blue). BrdU was delivered for 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. A and B: frequency (%) of BrdU-positive satellite cells in TA and MG muscles calculated as follows: (BrdU-positive satellite cells/total satellite cells) x 100. Open bars, control; solid bars, denervation; shaded bars, SI. *Significantly different from denervation; asignificantly different from day 28; bsignificantly different from day 14; #significantly different from control.

 


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Fig. 9. Number of BrdU-positive connective tissue cells per 100 fibers after 3, 14, and 28 days of denervation or SI in TA (A) and MG (B) muscles. All fibers and M-cadherin-negative cells (see Fig. 8) within each region of analysis (~0.3 mm2) were counted and calculated as (BrdU-tagged cells/fiber number) x 100 to account for atrophy in SI and denervated muscles. Open bars, control; solid bars, denervation; shaded bars, SI. *Significantly different from denervation; asignificantly different from day 28; bsignificantly different from day 14; #significantly different from control.

 

Satellite cell differentiation in both muscles was higher in denervated than in SI rats at days 14 and 28 (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 in SI muscles (Fig. 10, top). On occasion, evidence for the formation of new fibers was detected in denervated and, to a lesser extent, SI muscles (data not shown).



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Fig. 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. A and B: number of BrdU-positive myonuclei per 100 fibers after 3, 14, and 28 days of denervation or SI in TA and MG muscles. 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 fiber atrophy associated with SI and denervation. Open bars, control; solid bars, denervation; shaded bars, SI. *Significantly different from denervation; asignificantly different from day 28; bsignificantly different from day 14; #significantly different from control.

 


    DISCUSSION
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Muscle-derived biochemicals can affect neuronal plasticity, namely, elongation (22), and postsynaptic cell targeting (9) and even survival (51). The major findings in the present study suggest that neural factors also affect postsynaptic 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 over extended periods.

At the whole tissue level, the progressive muscle atrophy was less in SI than in denervated rats at 2 and 4 wk. This relative preservation of muscle mass is consistent with earlier observations in SI rats (20) and rats treated with tetrodotoxin (TTX) (13). The SI and TTX models block neurally mediated activation of the hindlimb muscles while maintaining the muscle-nerve connectivity. TTX is an Na+ 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 with denervation observed in 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 after denervation (55). Taken together, these studies suggest that the observed differences between SI and denervation 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 {alpha}-AChR 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 {alpha}-AChR 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. 4) suggests an initial response to the cessation of electrical stimuli in SI and denervated muscle, because both models reduce activation of the hindlimb muscles. The downregulation of myogenin mRNA at 2 (<7-fold) and 4 wk (<3-fold) after SI, however, does not support the theory that electrical activity is the principal neural stimulus regulating this muscle-specific gene. This compares with an ~10- to 15-fold increase of myogenin transcripts 1 mo after denervation (Fig. 4, C 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 upregulated in denervated, but not in SI, muscles. The increase in myogenin protein after 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 denervation. In the present study, MyoD protein was highly upregulated in denervated, but not SI, muscles, with the greatest differences observed at day 28. The differential expression of MyoD mRNA (Fig. 4) and protein (Fig. 5) in SI muscles suggests that 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 after denervation (5, 19, 28, 34). We are investigating the expression of several degradation pathway-related genes (Atrogin-1, MuRF-1, and polyubiquitin) that may help explain the MRF expression differences observed in SI and denervated muscles. In combination, these results indicate that the expression of these muscle-specific genes is modulated by activity-dependent and -independent mechanisms. Only at days 14 and 28 do the effects of activity-independent influences become clearly differentiated from activity-dependent mechanisms (Figs. 4 and 5).

Interestingly, Maier et al. (32) detected no MyoD mRNA or protein in denervated rat TA and soleus muscle at day 2, 7, or 28. Sakuma and coworkers (44) reported a gradual decrease in MyoD protein in denervated plantaris and gastrocnemius muscles 1-28 days after surgery, whereas myogenin protein was relatively unchanged in these muscles. These findings contrast with other studies showing an increase in these MRFs after denervation (2, 8, 18, 29, 52, 55). It is unknown why such MRF expression differences exist with denervation. It is possible that the species of rat used in each study exhibits different MRF responses: Maier et al. and Sakuma et al. used Wistar rats, 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).

A number of candidate molecules 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 neuromuscular junction. Similarly, Witzemann and Sakmann (55) found MyoD and myogenin mRNA expression at junctional and extrajunctional 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 2 wk 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 1st wk after denervation (14, 26). Therefore, if CNTF-mediated signaling suppresses MyoD or myogenin expression, then these MRFs would remain more elevated in denervated than in SI muscles, as observed in the present study. Although CNTF inhibits acetylcholinesterase production in 7-day denervated rat soleus muscles (7), there is no evidence that CNTF can similarly block MRF expression. Furthermore, we cannot exclude the possibility that the presence of other neurotrophins, such as brain-derived neurotrophic factor (1) or nerve growth factor (4), also could have had activity-independent effects, inasmuch as cellular constituents within muscle tissue also produce these neurotrophins.

Using immunohistochemical analyses, several groups reported a widespread MRF response in denervated muscle nuclei (29, 52, 53). In these studies, the proportion of satellite cells and myonuclei expressing MyoD or myogenin was not ascertained, inasmuch 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 predominantly localized. In the present study, we hypothesized that MyoD and myogenin would be associated primarily with satellite cells on the basis of earlier reports of increased satellite cell activity in denervated muscles (33, 36, 48), that myonuclei are postmitotic, and that the present view is that MRF proteins play a predominant 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, if it is considered that the presence of MyoD and myogenin in myonuclei is 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.

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 colabeled) satellite cells (30, 32). We observed that M-cadherin-positive satellite cells also express MyoD, an MRF associated with proliferating myogenic cells. Cooper et al. (12) also showed that M-cadherin coexpressed 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 colabeled with M-cadherin in cultured denervated flexor digitorum brevis muscle fibers of the rat (30), which supports our findings that, in fact, MRFs are expressed predominantly in myonuclei, rather than in satellite cells, after denervation. This widespread upregulation in myonuclei suggests that MyoD and myogenin could be 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), {alpha}-AChR (39), and fast troponin I (31) genes, suggesting that the early upregulation 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, inasmuch 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 after analysis of cellular activity. Generally, connective tissue cell, as well as satellite cell, activity was lower in SI than in denervated muscles (Figs. 8, 9, 10). Augmented cellular proliferation (36, 46, 48) and satellite cell differentiation after denervation have been observed previously (3, 16). A regenerative-like response in denervated muscles has been attributed to initiation of 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. Although it is possible that migrating satellite cells also differentiated and helped form new fibers, rather than fused with the parent fiber, our 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 that the turnover rate of myonuclei, presumably via apoptosis, confounded our analyses, inasmuch 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 after 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) might preserve the satellite cell pool as well as the cytoplasmic components of the muscle fibers.

In summary, a more normal level of homeostasis was observed in muscle fibers having continuity with their motoneurons, albeit inactive, compared with muscle fibers having no anatomic 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 day 14, at which time there was a general return of MRF mRNA and, to a greater extent, protein levels to control values compared with denervation. In addition, after the initial 2-wk period, satellite cell activity returned to basal levels in SI muscles (day 28 in Figs. 8 and 10). These results highlight two important points: 1) physical contact and the ensuing biochemical communication between the nerve and muscle are key components in the homeostatic process for skeletal muscle, and 2) the role of MRFs in adult muscle is potentially as important in orchestrating an adaptive response of existing muscle fibers as in the regenerative responses of muscle fibers.


    DISCLOSURES
 
This work was supported by National Institute of Neurological Disorders and Stroke Grant NS-16333 (to V. R. Edgerton) and a grant from the Sigma Xi Society (to J.-P. K. Hyatt).


    ACKNOWLEDGMENTS
 
We thank Hui Zhong, Fadia Haddad, Anqi Qin, Ming Zeng, and Gary McCall for technical assistance. We especially thank Maynor Herrera for animal care and surgery assistance.

Illustrations in Fig. 1 were printed by exclusive agreement with Fairman Studios (Waltham, MA) and have not been previously published.


    FOOTNOTES
 

Address for reprint requests and other correspondence: V. R. Edgerton, Dept. of Physiological Science, 621 Charles E. Young Dr., University of California, Los Angeles, Los Angeles, CA 90095 (E-mail: vre{at}ucla.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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