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J Appl Physiol 97: 1584-1587, 2004. First published March 19, 2004; doi:10.1152/japplphysiol.00534.2003
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Myogenin and oxidative enzyme gene expression levels are elevated in rat soleus muscles after endurance training

Parco M. Siu, David A. Donley, Randall W. Bryner, and Stephen E. Alway

Laboratory of Muscle Biology and Sarcopenia, Division of Exercise Physiology, West Virginia University School of Medicine, Morgantown, West Virginia 26506

Submitted 19 May 2003 ; accepted in final form 12 March 2004


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The intent of this study was to determine whether endurance exercise training regulates increases in metabolic enzymes, which parallel modulations of myogenin and MyoD in skeletal muscle of rats. Adult Sprague-Dawley rats were endurance trained (TR) 5 days weekly for 8 wk on a motorized treadmill. They were killed 48 h after their last bout of exercise. Sedentary control (Con) rats were killed at the same time as TR animals. Myogenin, MyoD, citrate synthase (CS), cytochrome-c oxidase (COX) subunits II and VI, lactate dehydrogenase (LDH), and myosin light chain mRNA contents were determined in soleus muscles by using RT-PCR. Myogenin mRNA content was also estimated by using dot-blot hybridization. Protein expression levels of myogenin and MyoD were measured by Western blots. CS enzymatic activity was also measured. RT-PCR measurements showed that the mRNA contents of myogenin, CS, COX II, COX VI, and LDH were 25, 20, 17, 16, and 18% greater, respectively, in TR animals compared with Con animals (P < 0.05). The ratio of myogenin to MyoD mRNA content estimated by RT-PCR in TR animals was 28% higher than that in Con animals (P < 0.05). Myosin light chain expression was similar in Con and TR muscles. Results from dot-blot hybridization to a riboprobe further confirmed the increase in myogenin mRNA level in TR group. Western blot analysis indicated a 24% greater level of myogenin protein in TR animals compared with Con animals (P < 0.01). The soleus muscles from TR animals had a 25% greater CS enzymatic activity than the Con animals (P < 0.01). Moreover, myogenin mRNA and protein contents were positively correlated to CS activity and mRNA contents of CS, COX II, and COX VI (P < 0.05). These data are consistent with the hypothesis that myogenin is in the pathway for exercise-induced changes in mitochondrial enzymes.

myogenic transcription factors; mitochondrial enzymes; exercise; citrate synthase; MyoD

SKELETAL MUSCLE IS AN EXTREMELY dynamic tissue, which possesses the ability to adapt to a variety of physiological stimuli or external influences (37). Repeated bouts of exercise have been shown to induce a series of physiological and biochemical adaptations in skeletal muscle. These training adaptations normally improve the muscle's working efficacy, resulting in better functional capacity and physical performance of an individual. Endurance exercise training results in the upregulation of several metabolic enzymes (12, 19, 39), and this biochemical adaptation is associated with the training-induced improvement of the muscle's oxidative capacity. Although this endurance training-induced adaptation has been well characterized, the molecular events contributing to this physiological adaptation remain poorly understood.

Transcription of muscle-specific genes, including metabolic enzymes and contractile proteins genes, is regulated by DNA binding proteins, which are called myogenic regulatory factors (MRFs). MRFs belong to a superfamily of basic helix-loop-helix muscle-specific transcription factor proteins, which includes MyoD, myogenin, MRF4, and Myf-5 (36). A basic DNA-binding domain essential for sequence-specific DNA binding and a helix-loop-helix motif are required for heterodimerization in MRFs. This family of myogenic factors forms dimers with ubiquitous E proteins (e.g., E47 or E12), and the resulting E-protein heterodimeric complexes bind to the E-box consensus DNA sequence (5'-CANNTG-3') that is found in the regulatory region of the promoters of many skeletal muscle-specific genes. It has been shown that the E-protein heterodimers form readily and are efficient gene transactivators (11, 34, 35).

Several studies have suggested that MRFs may be involved in determining structural and metabolic phenotype in postmitotic skeletal muscles (1, 20, 22, 26, 32). For example, Hughes and coworkers (22) found that the distributions of MyoD and myogenin mRNAs are regulated in a muscle type-dependent manner. They showed that the MyoD transcript is prevalent in fast glycolytic muscle, whereas the myogenin transcript is mainly found in slow-oxidative muscle (22). Furthermore, the alteration of fast and slow fiber-type distribution by thyroid hormone treatment or by cross-reinnervation causes a corresponding alteration in the MyoD and myogenin transcriptional expression pattern (22). Several studies have shown that myogenin expression is associated with the expression of the slow myosin heavy chain isoform, whereas MyoD is related to the expression of the fast isoform (10, 21, 22). By using transgenic techniques, Hughes et al. (21) reported shifts in myosin heavy chains and fiber-type distribution in muscles of mice lacking a functional MyoD gene. Moreover, Hughes and colleagues (20) reported that overexpression of myogenin in skeletal muscles of transgenic mice shifted the muscle's metabolic enzyme content toward oxidative metabolism, such that levels of oxidative enzymes were increased, whereas glycolytic enzymes were decreased. Adams and coworkers (1) demonstrated that compensatory hypertrophy in rat plantaris muscles increased the expression of myogenin. Fiber-type-specific alterations in MRF transcription have also been reported during skeletal muscle disuse atrophy (26). Taken together, it is reasonable to suggest that myogenin and MyoD may have distinct roles in mediating adaptations to exercise, overload, or disuse in a fiber-type-specific manner. Thus it is possible that MRFs contribute to mechanisms controlling skeletal muscle phenotype. For instance, increased oxidative metabolic enzymes resulting from endurance training may be due, in part, to the altered expression pattern of MRFs following training. MyoD and myogenin act as independent key regulatory factors during early muscle differentiation (36). MRF expression also occurs in activated satellite cells and postmitotic adult skeletal muscles (14, 27, 41). Thus it is likely that MRFs play a role in mature muscle and not just in muscle development (20). Given that these MRFs provide a potential link between electrical activity and gene regulation (16), this raises the possibility that MRFs may reside in the pathway for activity-dependent changes in muscle-specific mitochondrial enzymes. Understanding these molecular links to activity-induced adaptations is important because they may yield suitable markers for tracking improvements in exercise training regimens that are intended to improve human performance. Therefore, the purpose of the present study was to test the hypothesis that endurance exercise training regulates increases in metabolic oxidative enzymes, which parallel elevation of myogenin but not MyoD in skeletal muscles of rats.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals.   Experiments were conducted on 3-mo-old adult Sprague-Dawley rats (Harlan, Indianapolis, IN). The rats were housed in pathogen-free conditions at 20–22°C. They were exposed to a reverse light condition of 12 h of light and 12 h of darkness each day and were fed rat chow and water ad libitum throughout the study period.

Treadmill training.   Sixteen rats with similar body weights were randomly assigned to control (Con, n = 8) or training (TR, n = 8) groups. Animals assigned to the TR group were trained by running on a level motorized rodent treadmill (Columbus Instruments, Columbus, OH) 5 days weekly for a period of 8 wk. During the first 4 wk, the speed of the treadmill and duration of the training sessions were gradually increased from a speed of 10 m/min for 10 min, to a running speed of 28 m/min for 55 min by the end of the 4th week. During weeks 5–8, a 5-min warm-up session at a speed of 20 m/min was followed by the 55-min training session at a speed of 28 m/min. Mild electrical shock stimulation was applied, if necessary, to maintain the running motivation during the training sessions. TR animals were killed 48 h after the last training session. Animals assigned to the Con group were handled daily and were exposed to the noise of the running treadmill by placing their cages next to the treadmill when their TR-matched partner ran on the treadmill. The intent of this procedure was to minimize any possible confounding effect of nontraining external factors on MRF responses in the rat muscles. Con animals were killed at the same time as the TR animals. All animals were killed via CO2 inhalation followed by decapitation, at which time the soleus muscles were quickly removed and frozen immediately in liquid nitrogen and stored at –80°C until further analysis.

All experimental procedures carried approval from the Institutional Animal Use and Care Committee from West Virginia University School of Medicine. The animal care standards were followed by adhering to the recommendations for the care of laboratory animals, as advocated by the American Association for Accreditation of Laboratory Animal Care and following the policies and procedures detailed in the Guide for the Care and Use of Laboratory Animals as published by the US Department of Health and Human Services and proclaimed in the Animals Welfare Act (PL89-544, PL91-979, and PL94-279) and fully conformed with the American Physiological Society's "Guiding Principles for Research Involving Animals and Human Beings" (7).

RT-PCR analyses of mRNA for MRF, myosin light chain kinase, and metabolic enzyme genes.   Total RNA was extracted from rat soleus muscles with TriReagent (Molecular Research Center, Cincinnati, OH), which is based on the guanidine thiocyanate method. Frozen muscles were mechanically homogenized on ice in 1 ml of ice-cold TriReagent. Total RNA was solubilized in RNase-free H2O incubated in DNase I (Ambion, Austin, TX) to remove any DNA present in the sample and quantified in duplicate by measuring the optical density (OD) at 260 nm. Purity of RNA was ensured by obtaining an 260-to-280-nm-OD ratio of ~2.0. Two micrograms of RNA were reverse transcribed with decamer primers and Superscript II RT in a total volume of 20 µl, according to standard methods (Invitrogen, Life Technologies, Bethesda, MD). Control RT reactions were done in which the RT enzyme was omitted. The control RT reactions were PCR amplified to ensure that DNA did not contaminate the RNA. One microliter of cDNA was then amplified by PCR by using 100 ng of each primer, 18S primer pairs (Ambion), 250 µM deoxyribonucleotide triphosphates, 1x PCR buffer, and 2 units of Taq DNA polymerase (Sigma Chemical, St. Louis, MO) in a final volume of 50 µl. The primer pairs were designed from sequences published in GenBank (Table 1). All PCR products were verified by restriction digestion and by sequencing. Preliminary experiments were conducted with each gene to ensure that the number of cycles represented a linear portion for the PCR OD curve for the muscle samples. The cDNA from all muscle samples were amplified simultaneously by using aliquots from the same PCR mixture. After the PCR amplification, 15 µl of each reaction were electrophoresed on 1.5% agarose gels, stained with ethidium bromide. Then images were captured, and the signals were quantified in arbitrary units as OD x band area by using Kodak one-dimensional (1D) image analysis system (Eastman Kodak, Rochester, NY). The size (number of base pairs) of each of the bands corresponded to the size of the processed mRNA. In the present study, ribosomal 18S primers were used as internal controls because 18S rRNA levels are not affected by endurance exercise training (33). All RT-PCR signals were normalized to the 18S signal of the corresponding RT product. This eliminated any potential measurement error from uneven sample loading and provided a semiquantitative measure of the relative changes in gene expression. Our laboratory has previously found that normalizing PCR signals to cyclophilin, which is another housekeeping gene, provides similar relative results as when the PCR signals are normalized to 18S (4, 28).


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  		Table 1. Primers used for PCR amplification of cDNA

 
Riboprobes for dot-blot hybridization.   Riboprobes were generated for rat myogenin that could be used in dot-blot hybridization. Rat myogenin was amplified by RT-PCR by using primers with EcoR I restriction sites (underlined) (upper, 5'-CAGAATTCACTAACACCCAGCCAGCAA-3'; lower, 5'-GCGAATTCTTGCCCCCCAGAGACTTTGG-3'; annealing temperature = 60.3°C; 340 bp). The resulting myogenin product was digested with EcoR I, gel purified, and then cloned into pBluescript II KS (±) Phagemid (Stratagene, La Jolla, CA). The PCR insert was verified by DNA sequencing. A riboprobe to rat GAPDH was generated as a potential housekeeping gene, but the expression of this gene was affected by endurance training, and, therefore, it was an unsuitable housekeeping gene by which to normalize the myogenin signals. Therefore a {beta}-actin probe was used as an internal control, and this was made by transcribing pTRI-Actin DNA template (Ambion). RNA probes were transcribed by using biotin-16-UTP (1388908, Roche) and T7 RNA polymerase (1324, MAXIscript T7/T3 kit, Ambion). The resulting {beta}-actin probe had ~10% of the UTP biotinylated, and 40% of the myogenin riboprobe were biotinylated with UTP.

Dot-blot hybridization for myogenin.   Dot-blot hybridization was carried out to further determine the mRNA level of myogenin in soleus muscles from Con and TR animals. Briefly, 10 µg of RNA were placed on a positively charged nylon membrane (10102, BrightStar-Plus, Ambion) and immobilized by ultraviolet cross-linking (UV cross-linker, Fisher Biotech). The immobilized RNA was prehybridized in ULTRAhyb hybridization buffer (NorthernMax kit, Ambion) at 68°C for 1 h. The myogenin or {beta}-actin riboprobe was added, and hybridization was carried out at 42°C overnight. Negative control experiments were performed by omitting the riboprobe during hybridization. After hybridization, the membrane was washed with Low and High Stringency Wash solutions (NorthernMax kit, Ambion). Biotinylated signals were detected by chemiluminescence (BrightStar BioDetect Nonisotopic Detection kit, Ambion). Signals were quantified in arbitrary units as OD x band area by using Kodak 1D image analysis system (Eastman Kodak). The myogenin signals were normalized to the {beta}-actin signal from the same RNA sample. GAPDH proved to be an inappropriate housekeeping gene in endurance-trained muscles.

Western blot analyses.   Myogenin and MyoD protein expressions were determined in the soleus muscles of Con and TR rats. Protein extracts were obtained from soleus muscle, as previously described (9). The protein contents of the solubilized extracts were quantified in duplicate by using bicinchoninic acid reagents (Pierce, Rockford, IL) and BSA standards. Soluble protein (40 µg) was boiled for 3 min at 100°C in Laemmli buffer and loaded on each lane of a 12% polyacrylamide gel. The proteins were separated by routine SDS-PAGE for 1.5 h at 20°C (2). The gels were blotted to nitrocellulose membranes (Bio-Rad, Hercules, CA) and stained with Ponceau S red (Sigma Chemical) to confirm equal loading and transferring of proteins to the membrane in each lane. As a second approach to verify similar loading between the lanes, gels were loaded in duplicate, and one gel was stained with Coomassie blue.

The membranes were then blocked in 4% nonfat milk in phosphate-buffered saline with 0.05% Tween 20 (PBS-T) and probed with anti-myogenin rabbit polyclonal antibody (M-225; Santa Cruz Biotechnology, Santa Cruz, CA) or anti-MyoD mouse monoclonal antibody (5.8A; PharMingen, San Diego, CA) at a concentration of 1–2 µg/ml in PBS-T in 1% milk. Secondary antibodies were conjugated to horseradish peroxidase (Chemicon), and the signals were developed by chemiluminescence (Pierce). The signals were visualized by exposing the membranes to X-ray films (BioMax MS-1, Eastman Kodak), and digital records of the films were captured with a Kodak 290 camera. The resulting bands were quantified as OD x band area by a 1D image analysis system (Eastman Kodak) and expressed in arbitrary units. The sizes of the proteins were verified by using standard molecular-weight markers (Bio-Rad). Although we had standardized the Western blotting protocol for all of the measurements (e.g., ran the SDS-PAGE for the same voltage and time, probed the membranes with same concentration of fresh antibodies and incubation time, and temperature, etc.), we found that there were blot-to-blot variations for the same sample in our immunoblots when blots were run on different days or weeks. To reduce this variation, we reran the Western blots, but included the same "control" protein sample on one lane of each gel. The signals obtained on the other lanes were then normalized to the signal from the "control" protein sample. In this way, we were able to directly compare the relative differences between samples on blots that were run on different days.

Citrate synthase activity assay.   Approximately 20 mg of soleus muscles were homogenized on ice in 0.1 M Tris buffer containing 0.1% Triton X-100, pH 8.35. Citrate synthase (CS) activity was determined spectrophotometrically, according to the method of Srere (40). The homogenates were frozen under liquid nitrogen and thawed four times to disrupt the mitochondria to release the CS. The assay system was in a total volume of 200 µl: 100 mM Tris buffer (pH 8.35), 5 mM DTNB, 22.5 mM acetyl-CoA, 25 mM oxaloacetate, and 4 µl of homogenate. The rate change in color was monitored at a wavelength of 405 nm at 15-s intervals for a period of 3 min by using a Dynex MRX plate reader, controlled through PC software (Revelation, Dynatech Laboratories). All measurements were performed in duplicate, in the same setting at 20–22°C. The solubilized protein extracts of the homogenates were quantified in duplicate by using bicinchoninic acid reagents (Pierce) and BSA standards. The CS activity was then normalized to the total protein content and was reported in a value in nanomoles per milligram protein per minute.

Immunocytochemistry.   Ten-micrometer-thick muscle sections were cut in a freezing cryostat at –22°C. The tissue sections were fixed in 50% methanol-50% acetone for 10 min, washed in PBS, and permeabilized with 0.2% Triton X-100 in 0.1% sodium citrate for 5 min. The tissue sections were blocked in 10% goat serum in PBS-T at room temperature for 30 min, washed in PBS, and incubated with anti-myogenin rabbit polyclonal antibody (1:10 dilution in 1.5% goat serum-PBS-T, M-225, Santa Cruz Biotechnology) at 4°C for overnight. Negative control experiments were performed by omitting the anti-myogenin antibody on the tissue sections. After a minimum of three washes in PBS-T, the sections were incubated with a biotin-conjugated anti-mouse/rabbit/rat IgG antibody (1:200 dilution in 1.5% goat serum/PBS-T, B1404, Sigma Chemical) at room temperature for 1 h followed by fluorescein avidin DCS incubation (1:10 dilution in sodium bicarbonate, A2011, Vector Laboratories) at room temperature for 45 min. The tissue sections were counterstained with 4',6-diamidino-2-phenylindole dimethylsulfoxide (DAPI; Vectashield mounting medium, Vector Laboratories). Myogenin- and DAPI-positive nuclei were examined with a fluorescence microscope (Biological Research Microscope E800 Eclipse, Nikon, NY), and images were obtained with a SPOT RT camera (Diagnostic Instruments). Metamorph software (Universal Imaging, Downingtown, PA) was used to superimpose images of DAPI and myogenin-positive nuclei.

Statistics.   Statistical analyses were performed by using the SPSS 10.0 software package. Student's t-test was used to examine differences between Con and TR groups. Relationships between given variables were examined by calculating the Pearson product-moment correlation coefficient, r. Statistical significance was accepted at P < 0.05. All data are given as means ± SE.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Body mass.   At the start of the study, animals in both groups had similar body weights (210–240 g). All animals assigned to the TR group successfully completed the treadmill training protocol. After the training, the animals were ~5 mo old. Animals undergoing endurance training weighed less than the sedentary Con animals at the time of death (Con: 449 ± 12.9 g; TR: 386.1 ± 10.2 g; P < 0.05).

MRF, MLC, and metabolic enzyme mRNA levels estimated by RT-PCR.   The total RNA content was similar in soleus muscles of Con and TR groups (Con: 0.98 ± 0.06 µg/µl; TR: 1.07 ± 0.11 µg/µl; P > 0.05). Transcriptional expression of myogenin, MyoD, CS, cytochrome-c oxidase (COX) subunits II and VI, lactate dehydrogenase (LDH), and myosin light chain (MLC) genes were analyzed by semiquantitative RT-PCR in soleus muscles of TR and Con animals. Representative ethidium bromide-stained gels for the mRNA after RT-PCR are shown in Fig. 1. A 25% higher level of myogenin mRNA (P < 0.01) was observed in TR animals compared with Con animals (Fig. 1), whereas the estimates of mRNA signals for CS, COX II, COX VI, and LDH were 20, 17, 16, and 18% greater (P < 0.05), respectively, in TR animals compared with Con animals (Fig. 1).



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  		Fig. 1. A: myogenin, MyoD, citrate synthase (CS), cytochrome-c oxidase (COX) subunits II and VI (COX II and COX VI), and lactate dehydrogenase (LDH) mRNA contents were estimated by RT-PCR. Normalized data are presented as means ± SE. Data were run in duplicate on different gels for each gene. *P < 0.05, **P < 0.01: data are significantly different from those of Con animals. B: representative PCR result from soleus muscles isolated from control (Con) and trained (TR) animals. PCR products were visualized with ethidium bromide staining. Quantification of the PCR signal was obtained by densitometric analysis of the signal product optical density (OD) x resulting band area. The expressions of genes were normalized to the ribosomal 18S signal (315 bp) from the same RT product.

 
In contrast, soleus muscle MyoD mRNA levels, as determined by analysis of RT-PCR, were similar between TR and Con groups (P > 0.05). We found that the ratio of myogenin to MyoD mRNA estimated by RT-PCR in TR animals was 28% higher than that in Con animals (Con: 1.39 ± 0.1; TR: 1.77 ± 0.1; P < 0.05) (Fig. 2).



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  		Fig. 2. Ratio of myogenin to MyoD mRNA contents determined by RT-PCR. Data are presented as means ± SE. *P < 0.05: data are significantly different from those of Con animals.

 
MLC expression was similar in samples obtained from Con and TR animals (P > 0.05) (data not shown). mRNA content of myogenin determined by dot-blot hybridization in muscles from the TR group was 16% higher than that in the Con group (P < 0.05) (Fig. 3). These dot-blot data using a riboprobe to myogenin confirmed the increase in myogenin mRNA that had been estimated by RT-PCR.



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  		Fig. 3. A: myogenin mRNA content was estimated by dot-blot hybridization to a myogenin riboprobe. Normalized data are presented as means ± SE. *P < 0.05: data are significantly different from those of Con animals. B: representative result of myogenin dot-blot hybridization of RNA from soleus muscles isolated from Con and TR animals. Quantification of dot-blot signals was obtained by densitometric analysis of the signal product OD x resulting band area. Expressions of genes are normalized to the {beta}-actin signal from the same RNA sample.

 
Myogenin and MyoD protein levels.   The extracted total protein content was similar in Con and TR groups (Con: 3.94 ± 0.24 µg/µl; TR: 3.94 ± 0.23 µg/µl; P > 0.05). An immunoreactive band of ~35 kDa corresponding to the predicted molecular mass of the rat myogenin protein and ~45-kDa immunoreactive band corresponding to the MyoD protein were detected in the Western blots. In the soleus muscles from the TR animals, the protein level of myogenin, as estimated by Western blot analysis, was 24% higher (P < 0.01) than that from the Con animals (Fig. 4). However, no difference was observed in the MyoD protein level between Con and TR groups (P > 0.05). The ratio of myogenin to MyoD protein levels in TR animals was not different from that in Con animals (P > 0.05).



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  		Fig. 4. Myogenin and MyoD protein levels were determined by Western blot analyses. Data are expressed as OD x resulting band area and are expressed in arbitrary units x 108. Inset: example of a Western blot for myogenin and MyoD from soleus muscles isolated from Con and TR animals. Data are presented as means ± SE. **P < 0.01: data are significantly different from those of Con animals.

 
CS activity.   CS activity was measured in soleus muscle samples from Con and TR animals. CS activity increased by 25% in muscles from the TR animals relative to the Con animals (TR vs. Con: 411 ± 11.2 vs. 329 ± 7.0 nmol·mg protein–1·min–1; P < 0.01) (Fig. 5).



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  		Fig. 5. CS activities of soleus muscles in Con and TR animals. CS activity was normalized to the total protein content of the sample used in the assay. Normalized data are presented as means ± SE. prot, Protein. **P < 0.01: data are significantly different from those of Con animals.

 
Relationships between myogenin and metabolic enzymes.   The relationships of myogenin and metabolic enzymes, including CS, COX II, and COX VI, were analyzed by examining the corresponding Pearson's correlation coefficient (r). Data from soleus muscles of both groups were collapsed and treated as a single group. The RT-PCR data showed that the myogenin mRNA content was significantly correlated with the CS activity (r = 0.648, P < 0.01), CS mRNA content (r = 0.715, P < 0.01), COX II mRNA content (r = 0.660, P < 0.01), and COX VI mRNA content (r = 0.515, P < 0.05) (Fig. 6).



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  		Fig. 6. Relationship between myogenin mRNA content and CS activity (A), CS mRNA (B), COX II mRNA (C), and COX VI mRNA contents (D) in soleus muscle samples obtained from Con and TR rats. mRNA contents were estimated by using RT-PCR, normalized to 18S mRNA, and expressed as relative OD x band area. Con and TR animals were collapsed and treated as a single pooled group.

 
Additionally, positive relationships existed between the ratio of myogenin and MyoD mRNA content and CS mRNA content (r = 0.559, P < 0.05), as well as COX II mRNA content (r = 0.529, P < 0.05) (Fig. 7). Moreover, the mRNA content of myogenin determined by dot-blot hybridization to a riboprobe was significantly correlated with CS activity (r = 0.503, P < 0.05) (Fig. 8).



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  		Fig. 7. Relationship between the ratio of myogenin and MyoD mRNA contents and CS activity (A) and COX II mRNA contents in soleus muscle samples (B) from Con and TR rats. mRNA contents were estimated by using RT-PCR, normalized to 18S mRNA, and expressed as relative OD x band area. Con and TR animals were collapsed and treated as a single pooled group.

 


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  		Fig. 8. Relationship between mRNA content of myogenin as estimated by dot-blot hybridization and CS activity in muscle samples from Con and TR animals. Myogenin mRNA was normalized to {beta}-actin mRNA and expressed as relative OD x band area. Con and TR animals were collapsed and treated as a single pooled group.

 
We found that the myogenin protein content was positively correlated with the CS activity (r = 0.573, P < 0.05), CS mRNA content (r = 0.662, P < 0.01), COX II mRNA content (r = 0.679, P < 0.01), and COX VI mRNA content (r = 0.566, P < 0.05) (Fig. 9). A positive linear relationship was also found between the ratio of myogenin and MyoD protein content and CS activity (r = 0.604, P < 0.05) (Fig. 10).



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  		Fig. 9. Relationship between myogenin protein expression and CS activity (A), CS mRNA (B), COX II mRNA (C), and COX VI mRNA contents (D) in soleus muscles from Con and TR animals. Con and TR animals were collapsed and treated as a single pooled group.

 


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  		Fig. 10. Relationship between the ratio of myogenin and MyoD protein contents and CS activity in Con and TR muscle samples. Con and TR animals were collapsed and treated as a single pooled group.

 
Immunocytochemistry localization of myogenin.   The quality and morphology of the muscle sections was less than optimal for immunocytochemistry. This was likely the result of freezing the tissue directly in liquid nitrogen, rather than in isopentane cooled in liquid nitrogen, and the freeze-and-thaw tissue damage that resulted from obtaining samples for Western blotting. Nevertheless, we observed myogenin-positive nuclei along the periphery of the muscle sections obtained from Con and TR animals (data not shown). This is the location at which satellite cells and myofiber nuclei should reside. However, the poor tissue quality did not permit us to determine whether the number of myogenin-positive nuclei differed in Con and TR muscles.


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Upregulation of mitochondrial oxidative enzymes is well recognized as a metabolic improvement after chronic endurance exercise training. However, the molecular mechanism of this specific metabolic improvement is not presently well understood. To the best of our knowledge, this is the first study to directly assess the effect of endurance exercise training on metabolic enzymes and MRFs. The major findings of this study are that 1) 8 wk of endurance training elevates mRNA and protein levels of myogenin by a similar magnitude as the increase in metabolic enzymes (i.e., CS, COX II, and COX VI), and 2) significant positive linear relationships exist among transcript and protein contents of myogenin and CS activity and transcript contents of CS, COX II, and COX VI after the 8-wk endurance treadmill training in the rat soleus muscles. These findings support the hypothesis that myogenin may have a role in the pathway leading to exercise-induced changes in metabolic enzymes.

MRF and endurance training.   Myogenin and MyoD are MRFs that act as key regulatory molecules during early muscle differentiation, and it has been suggested that they may play a more extended role because their expression also persists in postmitotic mature muscles (22). Several studies suggest that MRFs are involved in regulating the metabolic processes intrinsic to muscle catabolism or anabolism (1, 22, 26, 29, 32), and they are important regulators of muscle-specific gene expression during conditions of muscle loading, unloading, and regeneration (5, 29). MRFs have been reported to increase in response to various perturbations, including increased muscle loading (27, 41), disuse (6), and spinal cord injury (15). Indeed, they have been shown to be influenced by electrical activity (16, 31) and have been thought to provide a link in the pathway between electrical activity and acetylcholine receptor gene expression during disuse supersensitivity (18). Although the molecular mechanism by which exercise training induces metabolic and biochemical adaptations in skeletal muscles is largely unknown, it has generally been suggested that physical activity regulates muscle gene expression by altering the neural electrical activity provided to the muscle. This view has been generally accepted, although mechanical stretch and hormone modulation also contribute to altered MRF expression (17). Nevertheless, it is possible that MRFs may play a role in the training-induced metabolic adaptation to endurance training.

Taken together, we hypothesize that myogenin may be involved in the regulatory pathway linking neural-electrical activity that originates from endurance exercise to adaptations leading to altered oxidative gene expression and metabolic enzyme activity. Our findings demonstrate that myogenin transcriptional and protein expressions are increased in parallel with the increases in the transcripts for oxidative enzymes (CS, COX II, and COX VI), as well as the enzymatic activity of CS after 8 wk of endurance exercise training in rat soleus muscles. No apparent difference was found in mRNA and protein contents of MyoD and mRNA of MLC (a protein regulated by MyoD) between TR and Con animals. We found that myogenin mRNA and protein contents are positively correlated to CS enzyme activity and to the mRNA contents of CS, COX II, and COX VI. Significant positive linear relationships still exist when myogenin mRNA or protein levels were normalized to the corresponding MyoD mRNA or protein contents. From these findings, we speculate that myogenin may contribute to the metabolic enzyme adaptation after endurance exercise training, perhaps as a result of exercise-induced modification of the muscle's electrical (i.e., neural) activity.

Although denervation (3, 13), disuse (6), and spinal cord injury (15) have been shown to increase myogenin expression, it is unlikely that the rats in our study had exercise-induced atrophy in their soleus muscles. Our data are consistent with previous studies that have also reported that body weight, but not soleus muscle weight, is slightly suppressed in rats that were endurance trained compared with age-matched untrained animals (23, 43). Similarly, rats trained by running to exhaustion show a decrease in body mass without changes in heart or soleus muscle weight relative to control rats (44). Furthermore, we would have anticipated a decrease in MLC expression in muscles of TR rats compared with Con rats, if the trained muscles had been atrophic, but this was not the case. Thus it is very unlikely that the soleus muscles of TR rats were atrophic relative to the Con rats, even though the body weight of the Con rats was greater than that of the TR rats. Additionally, the finding of unaltered MLC transcript content following the 8-wk endurance training suggested that MLC, a key constituent protein of muscle, was not affected by the 8-wk training protocol. This was consistent with the endurance training-induced adaptations, which typically show little or no muscle hypertrophy. Nonetheless, the changes in myogenin transcript and protein levels were similar in direction and magnitude to the changes in the metabolic enzymes (CS, COX II, and COX VI) following the 8-wk endurance training.

MRFs are expressed in activated satellite cells or interstitial myogenic cells, such as muscle side-population cells in vitro (8, 25, 42). However, localization of myogenin to myofiber nuclei and satellite cells has been previously shown in rat muscles (14, 15). Clearly, activated satellite cells are not the only source of myogenin production, because irradiation of quail muscles has been shown to eliminate satellite cell proliferation, yet there is a significant (although suppressed) myogenin expression when irradiated muscles are loaded compared with nonirradiated muscles (27). It has been shown that proliferating satellite cells, as well as nonmitotic myonuclei, are capable of expressing MyoD and myogenin in vivo (27, 41), and muscle-derived hematopoietic cells may also participate in regulation of muscle transcription (24, 30). Myogenin protein levels were 24% greater in soleus muscles obtained from TR compared with Con rats. Although immunocytochemical analysis in our study showed localization of myogenin to peripheral nuclei in the location occupied by satellite cells and myonuclei, and this has also been seen by others (13, 15, 41), our observations did not permit us to conclude whether the frequency of myogenin-positive nuclei increased in parallel to the training-induced increase in myogenin that we observed via RT-PCR and Western blot analyses.

In the present study, we have demonstrated that myogenin is linearly related to the adaptations of oxidative metabolic enzymes (i.e., CS, COX II, and COX VI) from 8 wk of endurance training. Our previous study has shown that CS (a common oxidative marker for aerobic exercise training adaptations) is not affected by the acute response of exercise if the measurement is taken 48 h after the last exercise session of 8 wk of exercise training (38). In contrast, samples obtained 1 h postexercise were influenced by the last exercise session (38). If myogenin responds similarly, it would be unlikely that the alteration in myogenin that we have measured in TR muscles 48 h postexercise is due to the acute effect of the last exercise session. Nonetheless, we recognize that the interpretation of the linear relationship between the oxidative enzymes and myogenin has been based on measurements taken 48 h after the last exercise session of the 8-wk exercise-training program. This does not allow us to rule out any changes that may have occurred at other time points that were not examined in this study.

In summary, 8 wk of endurance exercise training increases mRNA contents of CS, COX II, COX VI, and LDH and CS enzymatic activity in soleus muscle of rats. Myogenin mRNA and protein levels increase with a similar magnitude as the increase in mRNA and enzymatic activity of the measured metabolic enzymes. The mRNA and protein contents of MyoD and the mRNA of MLC do not change after endurance training. In the present study, we have shown that significant positive linear relationships exist among mRNA and protein contents of myogenin, CS enzyme activity, and transcript contents of CS, COX II, and COX VI. Our findings have demonstrated that endurance exercise training regulates increases in metabolic enzyme mRNA (CS, COX II, and COX VI) and enzymatic activity (CS) that parallel upregulations of myogenin mRNA and protein in soleus muscle of rats. Although we could not rule out the other possible factors (e.g., mechanical stretching and hormones) that could influence the metabolic profile of skeletal muscle after exercise training, we have provided evidence showing that myogenin may be involved in the regulatory process of the exercise-induced metabolic adaptation. Our data are in agreement with the hypothesis that endurance exercise training upregulates the metabolic enzymes by modifying the muscle's neural-electrical activity, and myogenin acts as a link between electrical activity and skeletal muscle gene expression. Nonetheless, the molecular mechanism that regulates the training-induced metabolic adaptation of skeletal muscle is still unclear. Additional research is required to fully define the intracellular pathway and the molecular mechanisms that control the metabolic adaptations of skeletal muscle to endurance exercise.


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 ABSTRACT
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REFERENCES
 
This study was supported by National Institute on Aging Grant AG-021530 and Research Development Funds from West Virginia University School of Medicine Grant 10005611 (to S. E. Alway).


   FOOTNOTES
 

Address for reprint requests and other correspondence: S. E. Alway, Div. of Exercise Physiology, School of Medicine, Robert C. Byrd Health Science Center, West Virginia Univ., Morgantown WV 26506-9227 (E-mail: salway{at}hsc.wvu.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.


   REFERENCES
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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