The signal transduction cascades that maintain muscle mass remain to be fully defined. Herein, we report that inhibition of extracellular signal-regulated kinase 1/2 (ERK1/2) signaling in vitro decreases myotube size and protein content after 3-day treatment with a MEK inhibitor. Neither p38 nor JNK inhibitors had any effect on myotube size or morphology. ERK1/2 inhibition also upregulated gene transcription of atrogin-1 and muscle-specific RING finger protein 1 and downregulated the phosphorylation of Akt and its downstream kinases. Forced expression of enhanced green fluorescent protein-tagged MAPK phosphatase 1 (MKP-1) in soleus and gastrocnemius muscles decreased both fiber size and reporter activity. This atrophic effect of MKP-1 was time dependent. Analysis of the reporter activity in vivo revealed that the activities of nuclear factor-κB and 26S proteasome were differentially activated in slow and fast muscles, suggesting muscle type-specific mechanisms may be utilized. Together, these findings suggest that MAPK signaling is necessary for the maintenance of skeletal muscle mass because inhibition of these signaling cascades elicits muscle atrophy in vitro and in vivo.
- signal transduction
- muscle atrophy
skeletal muscle mass is maintained by a delicate balance between protein synthesis and protein breakdown and experiences hypertrophy or atrophy in response to altered functional demands by adjusting either side of this equilibrium (10, 12, 38). Triggered by extracellular signals such as growth factors and mechanical overloading, muscle can increase in mass by changing the overall dimensions of its fibers (4, 34). On the other hand, cued by a variety of stimuli ranging from immobilization, clinical application of corticosteroids, cachexia, and microgravity, to normal aging process (29, 32), skeletal muscle undergoes significant loss of mass. Interestingly, fast-twitch muscle tends to experience greater hypertrophy in response to growth promotants such as β-adrenergic receptor agonist (37). Similarly, slow and fast muscles respond differentially in distinct atrophy models. For example, unloading (9) induces profound atrophy in slow-twitch muscles, whereas nutritional deprivation- (26), glucocorticoid- (13), and sepsis- (30) induced atrophy is preferentially restricted to fast-twitch muscles.
The signaling pathways that govern muscle hypertrophy and/or atrophy have yet to be fully defined. Recent research has identified Akt and its downstream signal cascades as pivotal regulators of muscle hypertrophy by enhancing protein synthesis and concomitant repression of protein breakdown (35, 40). Among all the atrophic signaling pathways, NF-κB and ubiquitin-proteasome pathways appear to play critical role in the “course adjustment” of muscle atrophy (7, 18, 22, 24, 43). Efforts to find common markers across diverse atrophy models have identified two muscle-specific E3 ubiquitin-protein ligases, muscle-specific RING finger protein 1 (MuRF-1) and atrogin-1 (3, 14). Both genes are markedly upregulated in all atrophy models studied, and knockout of these genes attenuates denervation-induced atrophy (3).
In mammals, there are at least four MAPK subfamilies: ERK1/2, p38α/β/γ/δ, JNK1/2/3, and ERK5 (5, 6, 46, 49). JNK and p38 MAPKs are associated with cytokine-induced muscle wasting. In contrast, ERK1/2 pathway is associated with enhanced protein synthesis by regulating ribosomal RNA gene expression (39) and phosphorylating eukaryotic initiation factor 4E through its downstream kinase MAPK-interacting kinase 1 (33). ERK signaling is also suggested to mediate the hypertrophic effects of IGF-I (16) and β-adrenergic receptor agonists (37). Nonetheless, some report that Ras-ERK had no effect on muscle fiber size in regenerating rat soleus muscle (31). Curiously, ERK1/2 phosphorylation increases concurrently with p38 activation in a hindlimb suspension-induced atrophy model (19). In this report, we examined the role of MAPK signaling in the maintenance of muscle mass in vitro and in vivo. Our findings suggest that MAPK signaling, especially ERK1/2 pathway, is necessary to regulate muscle growth because inhibition of MAPKs induced pronounced atrophy in both slow and fast muscles.
MATERIALS AND METHODS
Antibodies and reagents.
Antibody against green fluorescent protein (GFP) was purchased from Molecular Probes (Eugene, OR). Anti-dystrophin antibody was from Chemicon International (Temecula, CA). Anti-ERK1/2 and anti-phospho-ERK1/2 (Thr202/Tyr204), anti-phospho-GSK3α/β (Ser21/9), anti-phospho-p70 S6 protein kinase (p70S6K) (Thr389), anti-Akt, and anti-phospho-Akt (Ser473) antibodies were from Cell Signaling Technology (Beverly, MA). Anti-α-tubulin antibody was from Sigma (St. Louis, MO). Pharmacological inhibitors PD-98059, SB-203580, and SP-600125 were from EMD Biosciences (La Jolla, CA).
Myosin heavy chain (MyHC) IIb-luciferase (luc) reporter containing sequences from −2560 to +13 bp was obtained from S. Swoap (Williams College, Williamstown, MA) (42). Myoglobin-luc reporter was constructed by cloning human myoglobin sequence from −373 to +7 bp into SacI/HindIII cloning site of pGL3 vector (Promega, Madison, WI). NF-κB-luc activity reporter containing 5× NF-κB enhancer elements was from Stratagene (La Jolla, CA). 26S proteasome activity reporter plasmid UbFL-luc containing four tandem copies of ubiquitin at the NH2 terminus of firefly luciferase was obtained from H. Piwnica-Worms (Washington University in St. Louis, St. Louis, MO) (27). Plasmid expressing Renilla luciferase (pRL-SV40) was purchased from Promega. Enhanced GFP (EGFP)-only plasmid pWay21 was a generous gift from T. Hughes (Montana State University, Bozeman, MT). EGFP-tagged MKP-1 (pWayMKP1) was secured from A. Bennett (Yale University, New Haven, CT) (48).
Cell culture and myotube analysis.
C2C12 myoblasts were maintained in growth medium (10% FBS in DMEM) until confluence. Myoblast differentiation was induced by replacing the growth medium with differentiation medium (2% horse serum in DMEM), and cells were maintained for 5 days. Then pharmacological inhibitors PD-98059 (25 μM), SB-203580 (5 μM), and SP-600125 (5 μM) were applied. For fiber diameter measurement, myotubes were fixed in 4% paraformaldehyde for 10 min. Four fields were randomly chosen from each well, and the images were analyzed with Photoshop (Adobe Systems, San Jose, CA). Three points evenly distributed along the length of an individual myotube were chosen, and the diameter measured from these points was averaged to give rise to the diameter of individual fibers. For protein assay, myotubes were washed in ice-cold PBS and lysed in nonionic buffer (1% Nonidet P-40 in PBS, pH 7.6, in the presence of 1 mM PMSF, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 50 mM NaF, 1 mM Na3VO4, and 10 U/ml benzonase) for 30 min on ice. Cell lysate was centrifuged at 13,000 g for 10 min at 4°C. The pellets were redissolved in 2% SDS in PBS by heating in boiling water. Protein concentration was assayed by a Bio-Rad protein assay kit (Bio-Rad, Hercules, CA), and the total protein was calculated on the basis of the concentration and the volume of the lysates.
Rat surgery and electroporation.
Adult male Sprague-Dawley rats (∼130 g body wt) were purchased from Harlan (Indianapolis, IN). Rats were anesthetized by intraperitoneal injection of 100 mg/kg ketamine and 10 mg/kg xylazine. A minor incision was performed on rat hindlimb to expose soleus muscle, plasmid DNA purified using Qiagen Endofree plasmid mega kit (Qiagen, Valencia, CA) at the concentration of 0.5 mg/ml in 0.9% saline was injected into soleus. Meanwhile, plasmids were also injected into gastrocnemius muscle. For reporter experiments, Renilla reporter pRL-SV40 was coinjected with other reporters at 1:100 ratio in mass to normalize transfection efficiency. After the incision was closed, electroporation was applied using BTX ECM electroporator (Genetronics, San Diego, CA) as previously described (36, 37). Briefly, eight pulses of stimuli were introduced by two spatula electrodes (200 V/cm, 20 ms in length, 1 Hz). At indicated time postelectroporation, muscles were dissected and prepared for subsequent analysis. For reporter activity assay, muscles were lysed in passive lysis buffer (Promega), and firefly and Renilla luciferase was assayed with Dual-Luciferase assay kit (Promega). For in vivo fiber size assay, muscles were mounted on OCT-bedded cork and frozen in isopentane cooled in liquid nitrogen. All procedures were approved by the Purdue Animal Care and Use Committee.
Immunohistochemistry and fiber size measurement.
Soleus/gastrocnemius muscles were cut into 10 μm in thickness. Sections were fixed in ice-cold acetone for 6 min and briefly washed in PBS (pH 7.3). Five-percent goat serum was used to block the sections for 30 min at room temperature. Primary rabbit anti-GFP and mouse anti-dystrophin antibodies diluted in 5% goat serum were applied on the sections for 1 h at room temperature. After 3× 5-min washes in PBS, sections were incubated with Cy3-conjugated goat anti-rabbit and Cy2-conjugated goat anti-mouse secondary antibodies for 1 h at room temperature. After 3× 5-min washes in PBS, photomicrographs were taken using Leica DMI6000 B microscope (Leica Microsystems, Bannockburn, IL) at 23°C using a Photometrics Cool-SNAP camera (Roper Scientific, Tucson, AZ). Leica DMI6000 B software was used to take images at the magnification of ×100. Image colors were then reassigned as follows: GFP, green, and dystrophin, red using Adobe Photoshop CS2 (Adobe Systems). Fiber cross-sectional area of the fibers was analyzed using Photoshop, first expressed as pixels, and then these pixels were converted to micrometer square using a micrometer image taken at the same magnification as reference.
RNA was isolated using TRIzol reagent following the manufacturer's recommended protocol (Invitrogen, San Diego, CA). After the treatment, myotubes were briefly washed in ice-cold PBS, and 1 ml TRIzol reagent was applied on each 35-mm well. Total RNA was isolated and quantified. First-strand cDNA was synthesized from 2.5 μg of total RNA using random hexanucleotide-primed cDNA synthesis. Each PCR reaction contained 100 ng of cDNA, 10 pmol primers, and iQ SYBR Green Supermix (Bio-Rad) in a total volume of 15 μl. Amplicons were purified from the gel and used as a standard to quantify the transcripts. Assays were performed in duplicate using an iCycler Real-time PCR Detection System (Bio-Rad). Primers for atrogin-1 were as follows: forward 5′-CTC AGA GAG GCA GAT TCG CAA GC-3′ and reverse 5′-GGG TGA CCC CAT ACT GCT CTC TTC-3′. Primers for MuRF-1 were as follows: forward 5′-TCA AGA GCA ATG TAG AAG CCT CCA-3′ and reverse 5′-CCT CAA GGC CTC TGC TAT GTG TTC-3′. Primers for 18S rRNA were as follows: forward 5′-GAC ACG GAC AGG ATT GAC AGA TTG-3′ and reverse 5′-AAA TCG CTC CAC CAA CTA AGA ACG-3′. Quantitative PCR data were first normalized to 18S rRNA, and then data from treatment were expressed as percent control.
Western blot analysis.
Cytosolic protein was extracted in nonionic buffer as described above. A total of 80 μg protein was separated on a 10% SDS-PAGE and transferred to polyvinylidene difluoride membrane. The membrane was blocked in 7% nonfat milk overnight at 4°C. Primary antibodies were applied to the membrane for 1 h at room temperature. After washes in TBS containing 0.1% Tween 20, the blots were incubated with peroxidase-conjugated secondary antibodies for 1 h at room temperature. Images were developed using enhanced chemiluminescence (Amersham, Piscataway, NJ). Blots for pERK1/2 and pAkt were stripped and reprobed for total ERK1/2 and Akt. For quantification of the immunoblots, phosphorylated ERK1/2, pAkt were normalized to total ERK1/2 and Akt, respectively, and are expressed as percent control. pGSK3 and pp70S6K were normalized to tubulin and are expressed as percent control.
Values are expressed as means ± SE (represented as error bars). Statistical analysis was performed using Student's t-test or ANOVA, and the significance level was set as P < 0.05.
Inhibition of ERK1/2 pathway induced atrophic responses in cultured myocytes.
To test the role of MAPKs in the maintenance of muscle mass, we treated C2C12 myotubes with pharmacological inhibitors specific to the various MAPKs. We chose day 5 differentiated myotubes based on our observation that two differentiation markers, muscle creatine kinase (H. Shi, unpublished observation) and myosin heavy chain, reach a plateau at day 5 and remain elevated for at least 4 days (36). Neither the p38α/β inhibitor, SB-203580 (5 μM), or the JNK inhibitor, SP-600125 (5 μM), had detectable effects on fiber size 3 days after treatment (Fig. 1, B and C). In contrast, the MEK inhibitor, PD-98059 (25 μM), decreased myotube diameter by 35% after only 3 days of treatment. No noticeable differences in myotube morphology were observed at day 1 (Fig. 1, A, D, and E). Analysis of the cytosolic and contractile protein contents (40) indicated that the cytosolic and contractile proteins decreased by 22% and 30%, respectively, after 3 days of treatment (Fig. 1F). In contrast, we did not detect a noticeable effect of either p38 or JNK inhibitors (Fig. 1F). Together, these findings suggest that inhibition of ERK1/2 signaling induced myotube atrophy as reflected by the decrease in myotube size and protein content.
ERK1/2 inactivation-induced myocyte atrophy was accompanied by the upregulation of atrophic markers.
Muscle-specific E3 ubiquitin ligases atrogin-1 and MuRF-1 are upregulated in a variety of atrophy models (12). To investigate whether transcription of these genes was changed in our atrophy model, we quantified the transcripts of these two genes by quantitative RT-PCR. No significant changes were observed in the abundance of atrogin-1 and MuRF-1 mRNA in either SB-203580- or SP-600125-treated cells, whereas cells treated with PD-98059 experienced a marked increase in atrogin-1 and MuRF-1 expression by 39% and nearly 2-fold, respectively (Fig. 2, A and B).
Phosphorylation of Akt and its downstream kinases was downregulated in ERK1/2-inhibition-induced atrophy.
Because Akt and its downstream kinases are known to be involved in muscle hypertrophy (12) and are downregulated in a burn injury-induced muscle atrophy model (41), we examined whether Akt and its downstream kinases were affected in our atrophy model. Feeding myotubes with 25 μM PD-98059 for 3 days attenuated ERK1 and 2 phosphorylation by 45% and 38%, respectively (Fig. 3, A and B). This attenuation of ERK signaling decreased the phosphorylation of Akt by 55% (Fig. 3, A and C). Likewise, the phosphorylation of the downstream kinases, glycogen synthase kinase-3α and -β (GSK3), was reduced to 64% and 67%, respectively (Fig. 3, A and D). The other main downstream branch of the Akt signaling pathway is mammalian target of rapamycin (mTOR)/p70S6K signaling. The phosphorylation of p70S6K also decreased to 33% of control values (Fig. 3, A and E). Together, these findings indicate that the hypertrophic signaling cascades were downregulated in ERK1/2-inhibition-induced atrophy.
Overexpression of MKP-1 in slow-twitch soleus muscle induced profound atrophy.
To investigate the functional role of MAPKs in muscle mass maintenance in vivo, we used rats as our animal model. We chose rats because we wanted to test whether there was a muscle type-specific effect of MAPK inhibition, and in this regard, rat soleus is an ideal muscle because it is primarily composed of slow type I fibers. In addition, rat soleus is much larger than mouse soleus, which facilitates greater success when plasmids are delivered and electroporated into the muscle. Control EGFP did not change fiber cross-sectional area up to 10 days after electroporation (Fig. 4, A and B). Even though EGFP-only fibers were numerically larger than surrounding nontransfected fibers (Fig. 4C), differences were not statistically significant. The increase caused by overexpression of EGFP may be attributed to the fact that overproduction of the protein swelled the fibers. In contrast, forced expression of EGFP-tagged MKP-1 strikingly reduced fiber size by 63% by 10 days after electroporation (Fig. 4, A–C). To test whether gene transcription in soleus was affected and consistent with the observed morphological changes, we electroporated myoglobin-luc reporter into soleus muscle in the presence of either pWay21 or pWayMKP1. Seven days after electroporation, activity of myoglobin reporter was reduced by 62% as reflected by luciferase activity (Fig. 4D). Taken together, these findings indicate that inhibition of MAPK pathways by forced expression of MKP-1 induced profound muscle atrophy as demonstrated by decreases in fiber size and reduction of gene transcription.
MKP-1 overexpression induced time-dependent atrophy in fast-twitch gastrocnemius muscle.
To examine whether the atrophic responses observed in the slow-twitch soleus muscle could be reproduced in fast-twitch muscles, we electroporated the pWay21 and pWayMKP1 plasmids into the superficial gastrocnemius muscle where fibers are exclusively IIb/IIx fibers. This time we also investigated whether there was any time-dependent atrophic response. We chose 4, 7, and 13 days after electroporation and stained muscle sections with GFP antibodies and compared the size of GFP-positive fibers with their surrounding fibers. Again, there was no noticeable change in size of fibers overexpressing only EGFP (H. Shi, unpublished observation). Forced expression of EGFP-MKP-1 induced a reduction in fiber size similar to those observed in slow fibers in a time-dependent manner. MKP-1 decreased fiber size by 10%, 43%, and 63% at 4, 7, and 13 days after electroporation, respectively (Fig. 5, A and B). This time-dependent atrophic response was captured by frequency histograms. At day 4, the cross-sectional area of muscle fibers expressing EGFP-MKP-1 began to become smaller as compared with the surrounding fibers. At day 7, however, the size of EGFP-MKP-1 muscle fibers became even smaller, and this trend was more pronounced by day 13 (Fig. 5, A and B). Again, to determine whether gene transcription was affected by MKP-1 overexpression, we electroporated MyHC IIb-luc reporter into gastrocnemius muscle in the presence of either pWay21 or pWayMKP1. Seven days after electroporation, the reporter activity dropped by 53% (Fig. 5C), suggesting that fiber size reduction is also accompanied by the downregulation of gene transcription. Collectively, these data indicate that inhibition of MAPK signaling by MKP-1 induced muscle atrophy in a time-dependent manner.
NF-κB and proteasome activities were differentially activated in slow and fast muscles in response to MKP-1 overexpression.
To further investigate the underlying mechanism in MKP-1 overexpression-induced muscle atrophy, we chose two reporter constructs, NF-κB-luc and UbFL-luc. NF-κB plasmid contains five tandem NF-κB enhancer elements linked to luciferase; therefore this reporter provides an index of NF-κB activity. The other reporter, UbFL-luc, was used to monitor 26S proteasome activity in living animals (27). This reporter construct has four tandem repeats of ubiquitin fused in frame with luciferase. Thus degradation of the ubiquitin-tagged luciferase is inversely correlated with 26S proteasome activity; that is, reduced firefly luciferase activity implicates more involvement of 26S proteasome in the proteolytic degradation. Here, we showed that NF-κB activity increased by 4.8- and 1.8-fold in soleus and gastrocnemius, respectively, in response to forced expression of MKP-1 (Fig. 6, A and C). UbFL reporter luciferase increased by 25% in soleus muscle, although statistically there was no significant difference (Fig. 6B). In contrast, luciferase activity drastically decreased by 50% in gastrocnemius muscle in response to MKP-1 overexpression (Fig. 6D), suggesting that the activity of 26S proteasome is greatly enhanced during MAPK-inhibition-induced proteolysis in fast muscles. These findings indicate that NF-κB and UbFL reporters respond differentially to MKP-1 overexpression in slow and fast muscles.
In the present study, we first established an in vitro model in which attenuation of ERK1/2 signaling induced muscle atrophy. Mechanistically, upregulation of muscle-specific E3 ligase MuRF-1 and downregulation of signaling pathways involved in gene transcription and translation may account for the observed atrophic responses. We observed that MuRF-1 was preferentially upregulated over atrogin-1 (Fig. 2), a phenomenon that parallels the preferential degradation of contractile proteins over cytosolic proteins (Fig. 1F), implying that MuRF-1 plays a major role in degrading the contractile proteins. This notion is supported by studies of the substrates of these ligases. Atrogin-1 was reported to target the noncontractile proteins such as MyoD (44) and calcineurin (25) likely through eukaryotic initiation factor 3 (21), whereas MuRF-1 attacks proteins in the muscle contractile apparatus such as titin (28) and troponin I (20). Akt has been suggested to serve as a nodal point in the regulation of muscle hypertrophy and in the combat of muscle atrophy (11, 12). Yet in a burn injury-induced atrophy model, Akt and its downstream kinases are reported to be downregulated (41). Here, we report that attenuation of ERK1/2 pathway decreases the phosphorylation of Akt and its downstream kinases GSK3 and p70S6K. It appears that ERK1/2 inhibition-induced atrophy is caused by three distinct, yet related signaling pathways: first, reduction of ERK-mediated protein synthesis at the transcriptional (39) and translational levels (33); second, the indirect effects of the downregulation of the hypertrophic signaling such as Akt and it downstream kinases (12); finally, enhanced gene expression of atrophy markers such as MuRF-1 and atrogin-1.
Our in vivo studies using rat soleus and gastrocnemius muscles demonstrate that forced expression of MKP-1, a phosphatase that dephosphorylates and inactivates JNK, ERK1/2, and p38 MAPKs, induced profound muscle atrophy in a time-dependent manner. Phosphorylation of JNK, ERK2, and p38 MAPK increased by 5.4-, 3.1-, and 1.8-fold in mouse skeletal muscle deficient in MKP-1 (47), suggesting that MKP-1 dephosphorylates these three MAPKs in the order JNK > ERK1/2 >> p38 in skeletal muscle. Although we do not know exactly which MAPK pathway is responsible for MKP-1 overexpression-induced muscle atrophy, we speculate that this effect may be via inhibition of ERK1/2 signaling at least in fast gastrocnemius muscle for several reasons. First, our in vitro data demonstrate that inhibition of ERK1/2 pathway decreases myotube size and enhances protein degradation. Second, we have reported previously that the ERK1/2 pathway mediates β-adrenergic receptor agonist-induced muscle hypertrophy (37). Third, the ERK1/2 pathway is preferentially activated in fast skeletal muscles (36, 37). However, it is also likely that MKP-1-induced atrophy is through the combination effects of the inhibition of all three MAPKs by MKP-1. These MAPKs share some common substrates (45), which may provide a platform for these pathways to interplay to confer specific cellular responses.
Although forced expression of MKP-1 could induce profound atrophy in both slow and fast muscles, the mechanisms appear to be distinct. Among the pathways that are involved in muscle atrophy, NF-κB (7, 15) and ubiquitin-proteasome pathways (17, 23) have been suggested to mediate various maladies-induced loss of muscle mass. Interestingly, the basal level of NF-κB and UbFL reporter activities differs markedly in soleus and gastrocnemius muscles (Fig. 6), suggesting that distinct proteolytic pathways may be utilized to break down proteins by slow versus fast muscles in protein turnover in normal physiological status. In response to MKP-1 overexpression, NF-κB activity increased to a greater extent in soleus than in gastrocnemius, whereas the 26S proteasome activity was enhanced by twofold in gastrocnemius and had no significant change in soleus (Fig. 6). It appears that NF-κB, but not enhanced proteasome activity, plays a major role in mediating MAPK-inhibition-induced atrophy in slow muscle, whereas the enhanced activity of proteasome and, to a lesser extent, NF-κB may account for the atrophic responses in fast skeletal muscle.
In our recent study, we observed that overexpression of MKP-1 induced de novo synthesis of MyHC IIa/I in fast IIb/x fibers in mouse and rat fast gastrocnemius muscles (36). An interesting observation is that 13 days after MKP-1 overexpression, IIb/x fibers are about one third the size of normal IIb/x fibers in gastrocnemius (Fig. 5B), comparable to the size of normal type I/IIa fibers in the same muscle (H. Shi, unpublished observation). It seems that the conversion of IIb/x fibers to the slower IIa and I fibers is accompanied by both the switch of isoforms of contractile proteins, such as myosin heavy chain, and the decrease in fiber diameter. Along this line of reasoning, the increased 26S proteasome activity may account for the degradation of protein profiles characteristic of fast fibers to be replaced by slow fiber profiles, and the increased NF-κB activity may result from the slower phenotype because NF-κB activity is 3.5-fold greater in soleus than in gastrocnemius muscle (Fig. 6, A and C).
Nonetheless, in slow-contracting muscle, the atrophic effect of MKP-1 is more of a “pathological” implication. Although in a previous report (36) we observed increased activity of MyHC I-luc reporter by MKP-1 overexpression, in this study the slow fiber metabolic reporter myoglobin-luc decreased in response to MAPK inhibition. Similar phenotypical changes were observed in denervation-induced atrophy in mouse soleus muscle. Denervation increased the proportion of slow, type I fibers in soleus muscle (2, 8). Yet the fiber cross-sectional area and citrate synthase activity in the denervated soleus decreased (1). In our study, the increase in the activity of MyHC I-luc may reflect the compensatory mechanism that myofibers utilize in response to the enhanced protein breakdown. Yet, soleus muscle fibers expressing MKP-1 decreased in size 10 days after electroporation, a fact indicating that muscle atrophy was induced in soleus. In comparison with the phenomenon observed in fast gastrocnemius muscle, it is tempting to speculate that MAPK signaling may be tailored to fit different roles in slow and fast muscles in terms of muscle fiber specialization and muscle mass maintenance.
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