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. 2010 Aug;24(8):2985–2997. doi: 10.1096/fj.09-150045

MAP kinase phosphatase-1 deficiency impairs skeletal muscle regeneration and exacerbates muscular dystrophy

Hao Shi *, Emmanuel Boadu *,1, Fatih Mercan *, Annie M Le *, Rachel J Roth Flach *,2, Lei Zhang *, Kristina J Tyner †, Bradley B Olwin †, Anton M Bennett *,3
PMCID: PMC2909286  PMID: 20371627

Abstract

In skeletal muscle, the mitogen-activated protein kinase (MAPK) phosphatase-1 (MKP-1) is a critical negative regulator of the MAPKs. Since the MAPKs have been reported to be both positive and negative for myogenesis, the physiological role of MKP-1 in skeletal muscle repair and regeneration has remained unclear. Here, we show that MKP-1 plays an essential role in adult regenerative myogenesis. In a cardiotoxin-induced muscle injury model, lack of MKP-1 impaired muscle regeneration. In mdx mice, MKP-1 deficiency reduced body weight, muscle mass, and muscle fiber cross-sectional area. In addition, MKP-1-deficient muscles exhibit exacerbated myopathy accompanied by increased inflammation. Lack of MKP-1 compromised myoblast proliferation and induced precocious differentiation, phenotypes that were rescued by pharmacological inhibition of p38α/β MAPK. MKP-1 coordinates both myoblast proliferation and differentiation. Mechanistically, MyoD bound to the MKP-1 promoter and activated MKP-1 expression in proliferating myoblasts. Later, during myogenesis, MyoD uncoupled from the MKP-1 promoter leading to the down-regulation of MKP-1 and facilitation of promyogenic p38α/β MAPK signaling. Hence, MKP-1 plays a critical role in muscle stem cells and in the immune response to coordinate muscle repair and regeneration.—Shi, H., Boadu, E., Mercan, F., Le, A. M., Roth Flach, R. J., Zhang, L., Tyner, K. J., Olwin, B. B., Bennett, A. M. MAP kinase phosphatase-1 deficiency impairs skeletal muscle regeneration and exacerbates muscular dystrophy.

Keywords: myogenesis, muscle damage, MAPK, protein tyrosine phosphatase, muscle stem cells

The generation, maintenance, and repair of adult skeletal muscle are dependent on the activation of quiescent muscle precursor cells known as satellite cells (1,2,3). Located between the basal lamina and the sarcolemma of mature myofibers, satellite cells are isolated from the existing myofiber and are poised in a state of quiescence, exhibiting limited metabolic and gene expression activity. In response to skeletal muscle injury, in a process that remains poorly defined, satellite cells become activated and initiate a complex but coordinated series of events that involves proliferation, migration, and differentiation (1, 2).

The mitogen-activated protein kinases (MAPKs) constitute a family of serine/threonine kinases that control a multitude of cellular processes (4). The MAPK family includes the growth factor-responsive, extracellular signal-regulated kinases 1 and 2 (ERK1/2) and the stress-responsive MAPKs, p38 MAPK, and the c-Jun NH2-terminal kinase (JNK) (4). ERK1/2 have been shown to exhibit both positive and negative regulatory roles in myogenesis (5,6,7,8,9), whereas JNK has been shown to be either dispensable or negative for myogenesis (10,11,12). On the other hand, p38 MAPK, specifically p38α MAPK, has emerged as an essential positive regulator of myogenesis. p38α MAPK is required to mediate differentiation of myogenic cell lines (13,14,15). However, p38α/β MAPK has been suggested to be required for myoblast proliferation and differentiation (16), whereas others have proposed a role for p38α/β MAPK in cell cycle exit (15).

MKP-1 is a ubiquitously expressed, nuclear dual-specificity phosphatase, whose substrates include primarily p38 MAPK and JNK, with ERK1/2 exhibiting reduced substrate selectivity (17, 18). MKP-1 is an immediate-early gene and is induced by numerous stresses (18). Mice lacking MKP-1 exhibit enhanced levels of ERK1/2, JNK, and p38 MAPK in skeletal muscle, demonstrating that MKP-1 plays an essential physiological role as a negative regulator of the MAPKs in this tissue (19). MKP-1 has also been reported to play a role in skeletal muscle fiber specialization (20, 21) and muscle mass maintenance (22). The first suggestion that MKP-1, and hence the MAPKs, participate in myogenic regulation emerged from studies in which conditional overexpression of MKP-1 was shown to stimulate precocious myogenesis in the context of the inhibitory actions of growth factors (23). MKP-1 expression levels in proliferating myoblasts are initially high and decline upon the onset of myogenesis, suggesting that extinguishing the expression of MKP-1 might be a prerequisite, or at least a permissive event, that is required for myogenic progression and/or maintenance (15, 23, 24). In addition, overexpression of MKP-1 at the onset of myogenesis inhibits multinucleated myotube formation (23, 24). Hence, MKP-1 may play both positive and negative roles in myogenesis in a temporal manner by selectively regulating one or more MAPKs. As such, the complexity of the outcome through which MKP-1 integrates multiple MAPK activities cannot be simply inferred.

Growth factors such as hepatocyte growth factor, fibroblast growth factor (FGF), and tumor necrosis factor-α have been suggested to play important initiating roles in satellite cell activation. It is thought that physical trauma to the myofiber releases these growth factors and cytokines, which then stimulate satellite cell proliferation, migration, and differentiation, by activating the MAPK signaling cascade (1, 6, 25, 26). Given the complexity of how integration of both positive and negative MAPK-dependent signals by MKP-1 might influence skeletal muscle function, we attempted to resolve the issue of whether MKP-1 plays a critical role in skeletal muscle repair and regeneration using a genetic approach. The studies presented here have revealed that MKP-1 is an essential positive mediator of muscle repair and regeneration in vivo.

MATERIALS AND METHODS

Animals and reagents

mkp-1−/− mice (C57BL/6) were generated and maintained as described previously (19). To generate mkp-1−/− mice on an mdx (C57BL/10 ScSn DMDmdx) background, mkp-1−/− male mice were intercrossed with female mdx mice (the Jackson Laboratory, Bar Harbor, ME, USA). The F1 males were backcrossed with female mdx mice to establish a F2 progeny on an mdx background. F2 mdx/mkp-1+/− male and female mice were intercrossed. PCR primers for genotyping for mdx and mkp-1 mice have been previously described elsewhere (19, 27). All animal experiments were approved by Yale University Institutional Animal Care and Use Committee. MAPK inhibitors PD98059, SP600125, and SB203580 were purchased from EMD Biosciences (La Jolla, CA, USA).

Cardiotoxin (CTX)-induced skeletal muscle regeneration

Eight-week old mkp-1+/+ and mkp-1−/− mice were used. Mice were anesthetized by i.p. injection of 10 mg/kg ketamine and 1 mg/kg xylazine. CTX from Naja mossambica mossambica (Sigma Chemical, St. Louis, MO, USA) was injected into the right tibialis anterior (TA) muscle at the dosage of 50 μl of 0.1 mg/ml CTX in PBS per muscle. The left TA muscle was injected with 50 μl of PBS as control.

Analysis of skeletal muscle structure and function

TA and gastrocnemius muscles were isolated, weighed, and fixed in 10% neutral buffered formalin (3.7% formaldehyde) overnight; washed in 70% ethanol; and paraffin-embedded for hemotoxylin and eosin (H&E) staining. To analyze the cross-sectional area, random fields were taken from muscle sections; images were analyzed using the U.S. National Institutes of Health’s Image J software. The percentage of degenerating muscle area in mdx gastrocnemius was calculated as a ratio of the degenerating area to total area of the muscle section. To assess muscle damage in mdx/mkp-1+/+ and mdx/mkp-1−/− mice, sera were collected, and creatine kinase activity was analyzed using a creatine kinase assay kit (BioAssay Systems, Hayward, CA, USA), according to the manufacturer’s instructions. To evaluate muscle functionality, mice were allowed to walk on the iron cage cover, and then the cage was inverted so that the mice used their 4 legs to hold on to the cage. The time from the onset of the test to when mice fell was recorded as a gross measure of muscle strength.

Analysis of inflammatory responses

To stain macrophages and neutrophils infiltrated into the damaged area in CTX-injured muscle, paraffin-embedded TA muscle sections were incubated in sodium citrate buffer (10 mM sodium citrate with 0.05% Tween 20, pH 6.0) to retrieve antigens before immunostaining. mkp-1−/− mice on a mdx background were injected with 1% Evans Blue dye at the dosage of 1% of body weight. Muscles were collected 24 h later and snap-frozen in isopentane precooled in liquid nitrogen. Muscles were cut into 10-μm sections and fixed in ice-cold acetone for 6 min. Sections were incubated with 5% goat serum at room temperature for 1 h, followed by incubation with anti-CD11b (clone M1/70.15.11.5.2; Developmental Studies Hybridoma Bank, Iowa City, IA, USA) and 7/4 (clone 7/4; AbD Serotec, Raleigh, NC, USA) antibodies. Alexa Fluor 488 goat anti-rat secondary antibodies were applied, and images were taken using a LSM 510 meta confocal microscope (Zeiss, Jena, Germany). To quantify macrophage and neutrophil infiltration into CTX-injured muscle, TA, gastrocnemius, and quadriceps muscles were injected with CTX; 42 h later, muscles were pooled and enzyme-digested. Isolated cells were equally divided and blocked with 5% goat serum for 20 min. Cells were stained for 15 min on ice with anti-CD11b and anti-7/4 antibodies in 5% goat serum, respectively. After a brief wash, cells were stained with FITC-conjugated goat anti-rat secondary antibody for 15 min on ice before FACS analysis. Rat IgG isotype was used as a control. mkp-1/mdx muscles were processed similarly.

Myoblast proliferation and differentiation analysis

Satellite cell-derived myoblasts were isolated as described previously (28) from neonatal muscles and cultured at a concentration of 104 cells/ml in growth medium (20% FBS in F-10 medium containing 5 ng/ml FGF-2) for the indicated time. The number of cells within each clone was calculated, and the average cell number per clone was determined. For single-fiber explant cultures, extensor digitorum longus muscle was digested with 0.2% type II collagenase (Worthington Biochemical, Lakewood, NJ, USA) in DMEM at 37°C for 60 min with occasional agitation. Muscle fibers were dissociated by repeated titration with a sterile pipette, washed twice with DMEM, and plated onto 24-well plates. Satellite cells at this point become activated, migrate to the surface of the culture dish, and proliferate. Cell numbers were assessed per clone after culturing in F-10 medium containing 20% FBS with FGF-2 (5 ng/ml). Primary myoblasts were cultured in growth medium or differentiation medium (DMEM containing 2% horse serum) for the indicated times. Cells were washed once in PBS, fixed in 4% paraformaldehyde for 10 min at room temperature, and permeabilized with 0.25% Triton X-100 in PBS for 5 min. Cells were blocked in 5% goat serum for 30 min and incubated with embryonic myosin heavy chain (MyHC; Developmental Studies Hybridoma Bank) for 1 h at room temperature, washed 3 times for 5 min, and incubated with Alexa Fluor 546-conjugated goat-anti-mouse IgG for 1 h at room temperature. Nuclei were counterstained with DAPI. Where appropriate, cell lysates were subject to immunoblotting to detect the expression of MyHC.

BrdU incorporation assay

Single-fiber explants were prepared by digesting EDL and TA muscles with 0.2% collagenase II, washed twice with F-10 medium, and cultured in F-10 containing 20% FBS. Myoblasts derived this way are >95% pure as stained by either Pax7 or MyoD. BrdU (10 μM; Sigma Chemical) was applied to the medium for 2 h. Cells were then stained for FITC-BrdU and 7-AAD using the FITC BrdU Flow Kit (BD Biosciences, San Jose, CA, USA). MyoD was detected using anti-MyoD antibodies (Santa Cruz Biotechnology, Santa Cruz, CA, USA) and PE-conjugated goat anti-rabbit IgG (Jackson Immunoresearch, West Grove, PA, USA). Only MyoD-positive cells were analyzed for BrdU incorporation. For in vivo BrdU incorporation, gastrocnemius, and quadriceps muscles were injected with CTX, 24 h later, BrdU was injected intraperitoneally at 100 mg/kg body weight. Muscles were collected 18 h later, minced and digested, and cells isolated were subjected to FACS analysis.

Cell culture and transient transfection assays

C2C12 myoblasts were cultured in growth medium consisting of DMEM (Invitrogen, Carlsbad, CA, USA) supplemented with 10% FBS, 1% sodium pyruvate (Invitrogen), 50 U/ml penicillin, and 50 ng/ml streptomycin at 37°C in a humidified atmosphere of 95% air and 5% CO2. Subconfluent cultures of C2C12 myoblasts were transiently cotransfected in triplicate with a plasmid containing the MKP-1 promoter fused to the luciferase reporter gene (29), pRL-Renilla, and an expression plasmid for MyoD with or without an expression plasmid for the constitutively active mutant of MKK6 (MKK6-EE) using Lipofectamine 2000, according to the manufacturer’s instructions (Invitrogen). Transiently transfected C2C12 cells were harvested after 48 h and the luciferase activity was measured using the Dual-luciferase reporter system (Promega, Madison, WI, USA) according to the manufacturer’s instructions. MKP-1 luciferase activity was normalized to the Renilla values. RNA was isolated from C2C12 cells using TRIzol reagent (Invitrogen) according to manufacturer’s instructions. RNA was reverse transcribed with Applied Biosystems Reverse Transcription Reagents and RT-PCR was performed on ABI Prism 7500 (Applied Biosystems, Framingham, MA, USA); data were quantified using the ΔΔCT method as compared to 18S. For MyoD knockdown, C2C12 cells were transfected with control and MyoD siRNA (Santa Cruz Biotechnology) using Lipofectamine 2000; 48 h later, cells were shifted to differentiation medium for 24 h before cell lysates were collected and subjected to immunoblotting with MyoD antibody (Santa Cruz Biotechnology) or quantitative RT-PCR to analyze MKP-1 mRNA abundance using TaqMan MKP-1 primers (Applied Biosystems).

Chromatin immunoprecipitation assays

C2C12 myoblasts that were cultured in growth medium (GM) or in differentiation medium (DM) for 72 h were fixed in 1% formaldehyde for 10 min at room temperature followed by incubation in 0.125 M glycine for 5 min to stop cross-linking. Cells were then washed in phosphate-buffered saline (PBS) and recovered by scraping on ice and centrifugation, followed by lysis in SDS buffer (50 mM Tris-HCl at pH 8.1, 10 mM EDTA, 1% SDS, and protease inhibitors). Lysates were sonicated to yield genomic DNA fragments with a bulk size of ∼200–1,000 bp followed by centrifugation at 14,000 g for 15 min at 4°C. Supernatants were diluted in immunoprecipitation dilution buffer (0.01% SDS, 1.1% Triton X-100, 16.7 mM Tris-HCl at pH 8.1, 167 mM NaCl, and protease inhibitors). The diluted chromatin lysates were precleared by the addition of 50 μl salmon sperm DNA/protein A-agarose and incubation at 4°C for 1 h with gentle agitation. Following a brief centrifugation, lysates were separated into 1-ml aliquots and immunoprecipitated with 3 μg of anti-MyoD antibody (Santa Cruz Biotechnology) or a preimmune rabbit serum. Antibody nucleoprotein complexes were incubated overnight at 4°C and recovered by incubation with 50 μl salmon sperm DNA/protein A-agarose for 1 h at 4°C. 100 μl aliquots were reserved from the negative control (preimmune antibody) samples prior to washes, and these aliquots were processed in parallel with eluted samples and used as input DNA. Beads were washed once each with 900 μl of low-salt buffer (0.1% SDS, 1% Triton X-100, 20 mM Tris-HCl at pH 8.1, 2 mM EDTA, and 150 mM NaCl), high salt buffer (0.1% SDS, 1% Triton X-100, 20 mM Tris-HCl at pH 8.1, 2 mM EDTA, and 0.5 M NaCl), immunoprecipitation buffer 3 (1% Nonidet P-40, 1% sodium deoxycholate, 100 mM Tris-HCl at pH 8.1, and 500 mM LiCl), and twice with Tris-EDTA buffer (10 mM Tris-HCl at pH 8.1 and 1 mM EDTA). Nucleoprotein complexes were eluted from protein A-agarose beads by 5-min incubation at room temperature in 100 μl of immunoprecipitation elution buffer (1% SDS and 50 mM sodium bicarbonate). Cross-links were reversed by the addition of 4 μl 5 M NaCl and incubation at 65°C for 4.5 h. DNA fragments were recovered using QIAquick PCR purification columns (Qiagen, Valencia, CA, USA), and samples were eluted in 60 μl dH2O, 5 μl of these elutes were used in PCR reactions. The following primers were used in the ChIP experiments: MKP-1 (−1 kb) forward 5′-GAAGCATTCCCTTGTTCAGC-3′, reverse 5′-GCCACTCCTGGTATCACGTT-3′; MKP-1 (−3 kb) upstream forward 5′-CATCAGGGTGAGCCTTAAAG-3′, reverse 5′-TGGGAGTTGTCAATGCAGAC-3′; glucose-6-phosphate dehydrogenase forward 5′-AAGCCAAACTAGCAGCTAGG-3′, reverse 5′-GGGCTAGTCTATCATTGCAG-3′.

Statistical analysis

Results in this study are presented as means ± se. Statistical analyses were calculated using Student’s t test, assuming normal distribution and equal variance between different genotypes. Two-tailed distributions for P-value calculations were employed.

RESULTS

Lack of MKP-1 disrupts skeletal muscle regeneration and repair

We tested whether injury, induced by an intramuscular injection of CTX, in mice lacking MKP-1 (mkp-1−/−) impairs skeletal muscle regeneration. Following an intramuscular injection of CTX, reconstruction of the musculature rapidly ensues through a process whereby satellite cells become activated, proliferate, and fuse to form multinucleated fibers (30). We injected CTX into the TA muscle of 8-wk-old mkp-1+/+ and mkp-1−/− mice and monitored skeletal muscle repair. As a control, the contralateral TA muscle from mkp-1+/+ and mkp-1−/− mice was injected with PBS. We found that 5 d after muscle injury, nascent myofibers from mkp-1+/+ mice contained centrally localized nuclei, and myofiber bundles began forming (Fig. 1A). By 10 d following muscle injury, nascent TA myofibers in mkp-1+/+ mice were enlarged with prominent centrally located nuclei (Fig. 1B). There was no damage in the contralateral TA muscle, where only PBS was injected (Fig. 1A, control). In contrast, mkp-1−/− mice exhibited dramatic histological differences in muscle architecture (Fig. 1A, B). Notably, myofibers were visibly smaller as compared with those from mkp-1+/+ muscle sections at 5 and 10 d following injury (Fig. 1A, B). To quantify the observed phenotypic difference between injured mkp-1+/+ and mkp-1−/− mice, we measured the cross-sectional area of myofibers at 5 and 10 d postinjury. The cross-sectional area of nascent TA muscle fibers in mkp-1−/− mice was markedly smaller at both 5 and 10 d postinjury as compared with injured muscles from mkp-1+/+ mice (Fig. 1C). These findings indicate that lack of MKP-1 impaired, or at least delayed, skeletal muscle regeneration.

Figure 1.

Figure 1

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MKP-1 deficiency impairs muscle regeneration. TA muscle from 8-wk-old mice was injected with CTX. Contralateral TA was injected with PBS as a control. A, B) H&E muscle sections of control treatment and 5 d (A) and 10 d postinjury (B). Scale bars = 50 μm. C) Histograms of myofiber cross-sectional area in 5- and 10-d regenerating TA fibers. Data are derived from ∼800 myofibers from 4 mice/genotype counted at each time point.

Reduced body weight and muscle mass in mdx/mkp−/− mice

Duchenne muscular dystrophy (DMD) is a common X-linked disease caused by mutations in the gene encoding for dystrophin (31). DMD can be modeled in the mdx mouse strain, which lacks the dystrophin gene, and its skeletal muscle exhibits cycles of progressive degeneration and regeneration (31). To test the contribution of MKP-1 in the mdx mouse model, we intercrossed mkp-1−/− mice into the mdx background to generate double mutant mice. The resultant intercrosses yielded mdx/mkp-1+/+ and mdx/mkp-1−/− genotypes at the expected Mendelian frequency. At 8 wk of age, we noted that mdx/mkp-1−/− mice were significantly smaller as compared with mdx/mkp-1+/+ mice (Fig. 2A). The body weight of both female and male mdx/mkp-1−/− mice was significantly reduced as compared with mdx/mkp-1+/+ littermates (Fig. 2A). Whereas, at a similar age, the weights of mkp-1+/+ and mkp-1−/− are equivalent (19, 21), implying that the reduction in weight was due to the loss of MKP-1 in the mdx background. Consistent with this, the relative muscle weight (muscle weight/body weight) of the TA and gastrocnemius muscles from female and male mdx/mkp-1−/− mice was significantly reduced as compared with mdx/mkp-1+/+ mice (Fig. 2B, C). The cross-sectional area of both TA and soleus myofibers was 28% (mdx/mkp-1+/+; 1975±30 μm2vs. mdx/mkp-1−/−; 1,423±21 μm2, P<0.0005) and 15% (mdx/mkp-1+/+; 943±24 μm2vs. mdx/mkp-1−/−; 802±20 μm2, P<0.005) smaller, respectively in diameter in mdx/mkp-1−/− mice as compared with mdx/mkp-1+/+ mice. The change in the distribution of myofiber size in TA and soleus myofibers from mdx/mkp-1−/− mice was shifted to the left (smaller size) as compared with mdx/mkp-1+/+ mice (Fig. 2D, E).

Figure 2.

Figure 2

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Lack of MKP-1 reduces body weight and muscle mass in mdx mice. A) Phenotype and body mass of mdx/mkp-1+/+ and mdx/mkp-1−/− mice. B, C) Relative muscle mass of tibialis anterior (B) and gastrocnemius (C). D, E) Histogram distribution of myofiber size in tibialis anterior (D) and soleus (E). Data represent means ± se from ≥6 littermates in each genotype and gender. Data are means ± se. *P < 0.05, **P < 0.01 vs. mdx/mkp-1+/+ mice.

Exacerbation of the dystrophic phenotype in mdx/mkp−/− mice

Examination of muscle sections from the diaphragm, TA, gastrocnemius, and plantaris muscle groups revealed degenerating areas of muscle that were more widespread in mdx/mkp-1−/− mice as compared with mdx/mkp-1+/+ mice (Fig. 3A). To quantify these observations, we measured the area of degeneration from the gastrocnemius muscle. The degeneration area as a percentage of total muscle area was significantly higher in the mdx/mkp-1−/− muscles as compared with mdx/mkp-1+/+ mice (Fig. 3B). To further confirm the severity of the myopathy in mdx/mkp-1−/− mice, we injected Evans Blue dye into mice and harvested their muscles 24 h later. Myofibers that have compromised membrane integrity are susceptible to the uptake of the dye, and this serves as a measure of myofiber damage. We found that the percentage of Evans Blue-positive areas was higher in mdx/mkp-1−/− mice as compared with mdx/mkp-1+/+ mice (Fig. 3C). The serum level of creatine kinase also serves as a marker of muscle damage, and was found to be significantly higher in the mdx/mkp-1−/− mice as compared with mdx/mkp-1+/+ mice (Fig. 3D). To test whether gross skeletal muscle function was compromised in mdx/mkp-1−/− mice, we performed a grip test. When inverted, female and male mdx/mkp-1+/+mice maintained their grip for 175 and 156 s, respectively. Whereas female and male mdx/mkp-1−/− mice maintained their grip for only 19 and 46 s, respectively (Fig. 3E). Taken together, these data support the conclusion that loss of MKP-1 dramatically exacerbates the dystrophic phenotype in mdx mice, suggesting that MKP-1 functions to promote skeletal muscle repair and regeneration.

Figure 3.

Figure 3

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Lack of MKP-1 exacerbates muscular dystrophy in mdx mice. A) H&E staining of muscle cross-sections of the diaphragm (Dia.), TA, gastrocnemius (Gast.), and plantaris (PL) muscles in mdx/mkp-1+/+ and mdx/mkp-1−/− mice. Scale bar = 100 μm. B) Percentage of degenerating area (n=6 littermates/genotype). C) Percentage of Evans blue positive area in gastrocnemius muscle (n=5 littermates/genotype. D) Serum creatine kinase activity (n=8 mice/genotype). E) Grip test (n=6 mice/genotype and gender). Data are means ± se. *P < 0.05, **P < 0.01 vs. mdx/mkp-1+/+ mice.

Enhanced inflammatory response in skeletal muscles of mice lacking MKP-1

MKP-1 functions as a critical negative regulator of innate immunity (32,33,34,35). Therefore, we tested whether MKP-1-deficiency would elicit an enhanced immune response locally in response to skeletal muscle repair and regeneration. We found that both macrophage and neutrophil infiltration was greatly enhanced in the TA muscle of mkp-1−/− mice at 5 and 10 d post-CTX-induced injury (Fig. 4A). In mdx/mkp-1−/− mice, Evans Blue-positive myofibers were surrounded by a greater number of macrophages and neutrophils as compared with mdx/mkp-1+/+ mice (Fig. 4B). Quantitative assessment of the inflammatory response confirmed that in skeletal muscle from both mkp-1−/− and mdx/mkp-1−/− mice neutrophil and macrophage infiltration was significantly higher as compared with their respective wild-type counterparts (Fig. 4C, D). These findings suggest that an enhanced inflammatory response may contribute to the impaired skeletal muscle regeneration in mice lacking MKP-1.

Figure 4.

Figure 4

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MKP-1 deficiency enhances macrophage and neutrophil infiltration into degenerating muscles. A) At 5 and 10 d post-CTX-injury, TA muscle sections were stained for CD11b for macrophages (Mac.) and 7/4 for neutrophils (NF). B) mdx mice were injected with Evans Blue dye; 24 h later, gastrocnemius muscles were harvested and sectioned. CD11b and 7/4 antibodies were applied to detect for macrophages and neutrophils, respectively. C) FACS analysis of macrophages and neutrophils in CTX-damaged muscles. At 42 h after injury, quadriceps, gastrocnemius, and TA muscles were pooled for enzyme digestion. Isolated cells were then stained for FITC-CD11b and FITC-7/4 (n=5 mice/genotype). *P < 0.05, **P<0.01 vs. mkp-1+/+ mice. D) Muscles from mdx mouse hind limb were pooled for FACS analysis as in C (n=5/genotype). *P < 0.05 vs. mdx/mkp-1+/+ mice. Scale bars = 50 μm.

MKP-1 is required for myoblast proliferation

Satellite cells are essential for skeletal muscle regeneration after damage (30, 33, 36). Dysfunctional satellite cell proliferation and differentiation either alone, or in combination, contribute to impaired muscle regeneration. Therefore, we asked whether satellite cell-derived myoblasts from mice lacking MKP-1 were defective either in proliferation and/or differentiation. MKP-1 is an inducible immediate-early gene and behaves as such when myoblasts are stimulated with serum and, as expected, MKP-1 is absent in myoblasts derived from mkp-1−/− mice (Fig. 5A). When myoblasts were stimulated with FGF-2, the phosphorylation of the ERK1/2 and p38 MAPK was enhanced, both in magnitude and duration, in mkp-1−/− myoblasts as compared with mkp-1+/+ myoblasts (Fig. 5A). JNK phosphorylation in myoblasts in response to FGF-2 was undetectable. These results indicate that in myoblasts, MKP-1 is an essential negative regulator of at least ERK1/2 and p38 MAPK activity.

Figure 5.

Figure 5

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Satellite cells deficient in MKP-1 have impaired proliferation. A) Primary myoblasts were stimulated with either FCS or FGF-2 for the indicated times. Western blots for MKP-1 and phosphorylated and total ERK and p38 MAPK were measured. B) Myoblasts in growth medium were pulsated with 10 μM BrdU for 2 h; cells were stained for BrdU FITC and 7-AAD for FACS analysis (A, apoptotic cells; G0/G1, S, and G2+M). Data are expressed as means ± se from n = 4 littermates/genotype. C) Myoblasts were cultured at low density (104 cells/ml) for 3 d, and number of cells per clone was counted. Data are means ± se from ≥3 independent experiments. D) Individual myofibers were cultured for 4 d; number of myoblasts proliferating from these fibers on the dish was counted. Data are means ± se from ≥3 independent experiments. E) At 24 h after CTX injury, BrdU was injected into muscles of mkp-1+/+ and mkp-1−/− mice. Muscles were isolated 18 h later for FACS analysis. Data are means ± se from n = 4 littermates. *P < 0.05, **P < 0.01 vs. mkp-1+/+ mice.

When myoblasts were pulsed with BrdU for 2 h, the percentage of myoblasts entering into S phase of the cell cycle was 19.8% in mkp-1+/+ myoblasts, whereas only 11.6% of mkp-1−/− myoblasts entered S phase (Fig. 5B). In addition, the reduction of myoblasts in S phase was not a result of increased apoptosis, since this parameter was unaffected in mkp-1−/− myoblasts (Fig. 5B). Clonal proliferation analysis of myoblasts isolated either from muscles, or those derived ex vivo from single myofiber explants, showed that the number of myoblasts within a given clone was significantly lower in either those isolated from muscles (Fig. 5C), or those derived ex vivo from single myofibers (Fig. 5D), from mkp-1−/− mice as compared with those from mkp-1+/+ mice. We determined whether satellite cells in vivo also exhibited a similar proliferative defect. Mice were injured with CTX, and 24 h following injury, they were injected with BrdU for an additional 18 h. MyoD-positive satellite cells were analyzed by FACS for BrdU incorporation. The percentage of myoblasts that incorporated BrdU was significantly lower in CTX-injured muscle in mkp-1−/− mice as compared with mkp-1+/+ mice (Fig. 5E). Collectively, these data show that MKP-1 is required for myoblast proliferation, and its expression is essential for satellite cell proliferation in response to muscle injury.

Promiscuous myogenesis in mkp-1−/− myoblasts

Our findings that MKP-1 is required for myoblast proliferation (Fig. 5) suggested that mkp-1−/− myoblasts may have an increased propensity to differentiate. We stained primary myoblasts cultured for 6 d in growth medium with the terminal differentiation marker myosin heavy chain (MyHC) antibody and found that even in growth medium containing high serum (20% FBS), mkp-1−/− myoblasts began to differentiate into myotubes (Fig. 6A). We calculated the percentage of nuclei within MyHC-positive cells to the total number of nuclei (fusion index) in order to quantitatively assess the extent of myoblast differentiation. In mkp-1+/+ myoblasts, ∼12% of nuclei were found within MyHC-positive myotubes, indicating these myoblasts exited the cell cycle and underwent myogenesis. In contrast, mkp-1−/− primary myoblasts underwent precocious differentiation when cultured under high-serum conditions since ∼40% of myoblasts were found within MyHC-positive myotubes (Fig. 6B). The enhanced differentiative capacity of mkp-1−/− myoblasts was also observed when myoblasts were switched from growth medium to low-serum medium to induce differentiation. MyHC expression was detectable as early as 24 h of differentiation in mkp-1−/− myoblasts, whereas in mkp-1+/+ myoblasts, the expression of this marker was barely detectable (Fig. 6C) at this time point. These observations show that loss of MKP-1 results in aberrant myogenesis. Therefore, the inability to coordinate myoblast differentiation likely contributes to the impaired regenerative response in MKP-1-deficient mice.

Figure 6.

Figure 6

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MKP-1 deficiency induces precocious differentiation. A) Primary myoblasts were cultured in growth medium for 6 d; cells were stained for myosin heavy chain (MyHC) to visualize the differentiated cells. Arrows indicate differentiated myotubes. Scale bar = 100 μm. B) Percentage of nuclei within MyHC-positive cells (MyHC+nuclei) of total nuclei were calculated. **P < 0.01 vs. mkp-1+/+ mice. Data are expressed as means ± se from n = 4 littermates. C) Primary myoblasts were immunoblotted for MyHC expression following 24 h in differentiation medium.

p38α/β MAPK conveys MKP-1 effects on cell proliferation and myogenesis

We showed that the lack of MKP-1 expression increased ERK1/2 and p38 MAPK activities in myoblasts (Fig. 5A) and in skeletal muscle in vivo p38 MAPK, ERK1/2 and JNK activities are enhanced as compared with wild-type mice (19). MKP-1 dephosphorylates these MAPKs, not in isolation, but affects its actions on multiple MAPKs simultaneously. Despite this complexity of MKP-1 signaling we attempted to determine which of the MAPK (or MAPKs) is responsible for transmitting the MKP-1-deficient phenotype at the level of both myoblast proliferation and differentiation. To determine this, we used the inhibitors PD098059, SB203580, and SP600125 to specifically interfere with MEK/ERK1/2, p38α/β MAPK, and JNK, respectively, in order to determine which of these when inhibited would rescue the mkp-1−/− phenotype. We selected concentrations of these inhibitors that affected MAPK without causing deleterious effects on myoblast viability. Primary myoblasts isolated from myofibers were treated with these inhibitors for 4 d, and the number of cells per clone was counted as an indicator of the rate of proliferation. As shown previously, mkp-1−/− myoblasts are significantly impaired in their ability to proliferate as compared with mkp-1+/+ myoblasts (Fig. 7A–C). PD098059 was found to have a dose-dependent inhibition of myoblast proliferation in mkp-1+/+ myoblasts but was unable to rescue the inhibition of mkp-1−/− myoblast proliferation (Fig. 7A). SP600125 was also unable to rescue the proliferative defect in mkp-1−/− myoblasts (Fig. 7B). When myoblasts were treated with 5 μM SB203580, myoblast proliferation in mkp-1+/+ myoblasts was inhibited (Fig. 7C), supporting the notion that p38α/β MAPK is required not only for differentiation but also for proliferation (15, 16). In contrast, mkp-1−/− myoblasts when treated with 5 μM SB203580 increased myoblast proliferation to a level comparable to that of the mkp-1+/+ myoblasts (Fig. 7C).

Figure 7.

Figure 7

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Blockade of p38α/β MAPK rescues MKP-1-deficient myoblast phenotypes. Primary myoblasts from adult muscles were cultured in growth medium with different dosage of MEK inhibitor (PD98059) (A), JNK inhibitor (SP600125) (B), and p38α/β MAPK inhibitor (SB203580) (C, D) for 4 d (A–C), and 6 d (D). A–C) After 4 d in culture, number of cells per clone was counted. D) After 6 d in growth medium, cells were fixed and stained for MyHC; MyHC-positive nuclei were calculated as a percentage of total nuclei. *P < 0.05, **P<0.01 vs. vehicle treatment within each genotype; ‡P < 0.01 vs. mkp-1+/+ vehicle treatment. Data are expressed as means ± se from n = 4 littermates.

Because the p38α/β MAPK inhibitor rescued the impaired rate of proliferation in mkp-1−/− myoblasts, we asked whether the precocious differentiation observed in growth medium (Fig. 6A, B) could also be rescued by inhibition of p38α/β MAPK. To assess this quantitatively, the percentage of nuclei contained within myosin heavy chain-positive myotubes was determined. Inhibition of p38 MAPK had no detectable effect on the differentiation of mkp-1+/+ myoblasts in growth medium (Fig. 7D). In contrast, SB203580 inhibited the precocious differentiation of mkp-1−/− myoblasts where it significantly reduced the percentage of MyHC-positive nuclei (Fig. 7D). Taken together, these findings suggest that the effects of MKP-1 on myoblast proliferation and differentiation occur through its ability to antagonize p38α/β MAPK activity.

Regulation of MKP-1 expression by MyoD during myogenesis

During myogenesis MKP-1 protein expression in terminally differentiated myotubes is inversely correlated with the promyogenic actions of p38α/β MAPK (13, 15, 23, 24). Moreover, knockdown of MKP-1 during myogenesis results in enhanced p38 MAPK activity (24). In agreement with our previous findings, and those of others, MKP-1 mRNA and protein expression are detected in proliferating myoblasts, and these levels decline later on during myogenesis (Fig. 8A, B). p38α/β MAPK has been shown to be required for both satellite cell proliferation and differentiation (15, 16). We hypothesized that regulation of MKP-1 expression levels might serve to control the threshold of p38 MAPK (and/or other MAPKs) activities through which proliferative and differentiative cues are tightly regulated. We hypothesized that basal levels of p38α/β MAPK activity (high levels of MKP-1) may be sufficient to support myoblast proliferation; whereas higher levels of p38α/β MAPK activity (low levels of MKP-1) are required to promote myogenesis. Further, we speculated that transcription factors that are directly coupled to myogenesis might control the regulation of MKP-1. We tested whether MKP-1 could be a target for regulation by myogenic regulatory factors (MRFs) such as MyoD. Using a MKP-1 luciferase reporter containing 1.1 kb of the proximal MKP-1 promoter (29), we found that MKP-1 reporter activity in either proliferating C2C12 myoblasts (containing endogenous MyoD) or in a heterologous system in COS7 cells (lacking MyoD), was enhanced by either exogenous expression of MyoD, p38 MAPK activation by a constitutively active mutant of the upstream p38α/β MAPK kinase (MKK6) or both (Fig. 8C, D).

Figure 8.

Figure 8

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MKP-1 is a target for transcriptional activation by MyoD. A, B) C2C12 myoblasts were cultured in either growth medium (GM) or varying times in differentiation medium (DM). Expression of MKP-1 was determined by quantitative RT-PCR (A) and Western blotting (B). C) C2C12 myoblasts were transiently transfected with a −1.1 kb MKP-1 proximal promoter fused to luciferase along with MyoD, MKK6-EE, or both and pRL-Renilla. Cells were cultured in GM for 48 h, and luciferase activities were normalized to Renilla. Results shown are representative of means ± se of 4 independent experiments. *P < 0.05, **P < 0.01 vs. vector control. D) Experiments were performed in COS7 cells as described in C. E) Chromatin immunoprecipitation assays on the endogenous MKP-1 promoter were performed on C2C12 myoblasts cultured in GM or DM for 72 h using either preimmune (Pre-Imm) or anti-MyoD antibodies. Bottom panel shows PCR amplification of the G6PD promoter as control. F) Schematic representation of the wild-type proximal MKP-1 promoter containing the putative wild-type E-box (WT MKP-1-luc) and the mutated E-box (Mut MKP-1-luc) at position −341 to −346. G, H) C2C12 myoblasts (G) and COS7 cells (H) were cultured under proliferative conditions and were transfected with either WT MKP-1-luc or Mut MKP-1-luc, as indicated, in absence or presence of MyoD. Luciferase values were normalized to pRL-Renilla; results represent means ± se from 3 independent experiments. *P < 0.05; **P < 0.01. I, J) C2C12 cells were transiently transfected with control or MyoD siRNA, 48 h later, cells were allowed to differentiate for 24 h. I) Western blotting for MyoD 72 h post-transfection. J) MKP-1 mRNA levels were determined by quantitative RT-PCR and expressed as percentage of control.

Consistent with the differential expression of MKP-1 in proliferating myoblasts and terminally differentiated myotubes (Fig. 8A, B), we found that MyoD formed a complex with the −1-kb region of the MKP-1 promoter to a much greater extent in proliferating myoblasts as compared with differentiated myotubes by chromatin immunoprecipitation experiments with antibodies to MyoD (Fig. 8E). The specificity for MyoD interacting with this region of the MKP-1 promoter was demonstrated by showing that MyoD did not bind to a region of the MKP-1 promoter that was located at −3 kb (data not shown). These data suggest that MyoD preferentially drives the expression of MKP-1 in proliferating myoblasts but not in terminally differentiated myotubes. To further define the mechanism of MyoD-induced transcriptional activation of MKP-1, we inspected the proximal MKP-1 promoter for putative E-box elements, which are transcriptional target sequences for DNA binding of MyoD. We identified a single E-box element within the −1-kb promoter fragment of MKP-1 (Fig. 8F). When this E-box was mutated, MyoD-dependent up-regulation of MKP-1 transcription in both C2C12 myoblasts and in COS7 cells was abrogated (Fig. 8G, H). To further characterize the regulation of MyoD on MKP-1 gene expression, we knocked down MyoD using siRNA transfection and tested whether MKP-1 gene expression was altered. Knockdown of MyoD (Fig. 8I) reduced MKP-1 mRNA by 46% (Fig. 8J). Collectively, these data demonstrate that MyoD stimulates MKP-1 transcriptional activation through the E-box element of the MKP-1 promoter in proliferating myoblasts but to a much lesser extent in terminally differentiated myotubes.

DISCUSSION

Although it has been proposed that MKP-1 plays a role in myogenesis in cultured muscle cell lines (15, 23, 24), a role for MKP-1 in context of muscle repair and regeneration in vivo, is unknown. The complexity of MKP-1-mediated inactivation of multiple MAPK family members, implicated as both positive and/or negative regulators of myogenesis, makes it unclear as to whether MKP-1 positively or negatively regulates muscle repair and regeneration. In this study, we show that MKP-1 is required for muscle repair and regeneration in response to injury, as well as in a pathophysiological model of muscle repair and regeneration in the mdx mouse. Furthermore, these results provide genetic evidence that dysfunctional MAPK signaling, either downstream of the dystrophin-glycoprotein complex, and/or through a parallel MAPK-dependent pathway, plays an important role in adult regenerative myogenesis and in the pathogenesis of muscular dystrophy.

A role for MKP-1 in muscle repair and regeneration can be attributed, at least in part, to a cell autonomous effect in myoblasts. Our results show that MKP-1-deficient myoblasts exhibited reduced proliferation as compared with wild-type myoblasts demonstrating that MKP-1 is required for myoblast proliferation. Previous studies in fibroblasts lacking MKP-1 also showed similar results (37), suggesting that MKP-1’s positive role in cell proliferation may be more general. Reducing the elevated levels of p38 MAPK activity, but not those of either ERK1/2 or JNK activities, in mkp-1−/− myoblasts by using pharmacological inhibitors rescued this phenotype. These results are consistent with those in which myoblasts derived from p38α MAPK-deficient mice showed increased proliferation (15). Interestingly, we also observe reduced Pax7-positive cells in uninjured skeletal muscles of mkp-1−/− mice, which supports the observation that MKP-1 is required for satellite cell proliferation (unpublished results). Provocatively, these results appear to be at odds with those in which overexpression of MKP-1 in proliferating myoblasts also inhibited cell proliferation and led to ectopic differentiation (23). One possible explanation for this difference is that when overexpressed, MKP-1 dephosphorylates not only p38 MAPK, but also ERK1/2 and JNK. The outcome of the effects of ERK1/2 and JNK inhibition in this scenario was proposed to lead to an inactivation of cyclin D1 resulting in cell cycle exit and entry into myogenesis, even in the presence of serum (23). These overexpression experiments are likely to be distinct in their downstream effects as compared to the loss of MKP-1, where presumably the consequences of enhanced p38 MAPK predominates over ERK1/2 and/or JNK, such that myoblasts are driven into differentiation rather than enhancing their proliferation. Although our results strongly suggest that MKP-1 acts through p38 MAPK, we could not distinguish the contribution of either p38α MAPK or p38β MAPK due to the indiscriminate effect of SB203580 on both isoforms. Nevertheless, it seems likely that p38α MAPK plays a major role (15), in light of reports that deletion of other p38 isoforms have no effect on myogenesis (38). We cannot exclude the possibility that the skeletal muscle phenotype of mkp-1−/− mice is a more subtle but complex blend of other MAPKs, which are simultaneously dephosphorylated by MKP-1. Nonetheless, it is reasonable to propose that MKP-1 exerts its positive effects on proliferation via repression of p38α MAPK activity.

We found that isolated myoblasts from MKP-1-deficient mice were impaired in their ability to proliferate and showed enhanced differentiative capacities, suggesting that miscoordination of these events, in part, contributes to a loss of fidelity in the repair and regenerative response. The major promyogenic MAPK, p38α MAPK, is active in proliferating myoblasts and undergoes further increases in activity concomitant with the decline of MKP-1 levels during myogenesis. MKP-1 is a major negative regulator of p38 MAPK activity in skeletal muscle in vivo and during myogenesis (19, 24). However, how p38 MAPK activity is regulated such that it is required for both myoblast proliferation and differentiation is unclear. We propose that MyoD supports the expression of MKP-1 following satellite cell activation, which retains myoblasts in a proliferative state, and restricts premature differentiation. Although MyoD is considered to be transcriptionally inactive in proliferating myoblasts, genome-wide ChIP-on-ChIP analyses for MyoD targets have demonstrated that MyoD, as well as other MRFs, have an array of both myogenic and nonmyogenic targets in proliferating and differentiating myoblasts (39). Therefore, our data are consistent with a role for MyoD in regulating MKP-1 in proliferating myoblasts. With the onset of myogenesis, MyoD disengages from the MKP-1 promoter, correlating with reduced MKP-1 expression during differentiation, thereby relieving the negative effects of MKP-1 on p38 MAPK activity. Given that p38α MAPK is required for myogenesis, our data support previous observations whereby overexpression of MKP-1 during myogenesis inhibited differentiation, and loss of MKP-1 during myogenesis, results in enhanced p38 MAPK activation and differentiation (23, 24). The regulation of p38 MAPK phosphorylation by MKP-1 may be through distinct mechanisms at different stages of myogenesis, given the fact that other MKPs that inactivate p38 MAPK may work synergistically with MKP-1 to fine tune its regulation in a temporal and spatial manner. Indeed, we have shown recently that in skeletal muscle, MKP-1 regulates the nuclear pool of p38 MAPK to control phosphorylation of a transcriptional coactivator protein (21). Hence, the actions of MyoD on MKP-1 may serve to control this local pool of activated p38 MAPK. Once the MKP-1-regulated p38 MAPK-dependent nuclear substrates involved in myogenesis have been identified, it would then become possible to assess what is presumably a discrete mode of both spatial and temporal regulation of MKP-1/p38 MAPK-mediated signaling.

We found that MKP-1 deficiency alone in a mouse model of local muscle damage, and in mdx mice, resulted in increased levels of neutrophil and macrophage infiltration around areas of damaged myofibers. The cause of this enhanced inflammatory infiltration into damaged muscles of mkp-1−/− and mkp-1−/−/mdx mice is likely due to the fact that MKP-1 is a critical negative regulator of innate immune responsiveness (32,33,34,35, 40). Chronic inflammation is likely causal to the progressive deterioration of muscle function following muscle injury and in muscle diseases such as DMD (41). Similarly, because of the inhibitory role of MKP-1 in the innate immune response, it is likely that the enhanced inflammatory response seen in MKP-1-deficient backgrounds exacerbates muscle damage. Hence, MKP-1 may serve to promote muscle repair and regeneration, and reduce muscle deterioration in DMD by attenuating chronic inflammation. Yet to build a causal relationship between the enhanced macrophage and neutrophil accumulation and deteriorating regeneration observed in MKP-1-deficient mice, tissue-specific knockouts of MKP-1 in skeletal muscles and macrophages will be required to dissect satellite cell-autonomous and immune response contributions on regenerative myogenesis.

In summary, we define MKP-1 as a critical component in the signaling pathway controlling skeletal muscle repair and regeneration. MKP-1 may also play dual roles, not only in controlling satellite cell proliferation and differentiation, but also in limiting the inflammatory response in the damaged areas. Together, both processes require close coordination in order to restore functional skeletal muscle. Finally, our results define MKP-1 as playing a role in the pathogenesis of DMD, which may open new avenues of investigation into potential therapeutic strategies targeting the MKPs for the treatment of skeletal muscle diseases.

Acknowledgments

This work was supported by National Institutes of Health grants AR46504 to A.M.B. and AR039467 to B.B.O.

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