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GROWTH, DIFFERENTIATION, AND APOPTOSIS
1Department of Movement Sciences, University of Illinois, Chicago, Illinois; 2Department of Bioengineering, University of California, San Diego, California; and 3Department of Cell Biology, Vrije Universiteit, Amsterdam, The Netherlands
Submitted 16 May 2007 ; accepted in final form 19 July 2007
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ABSTRACT |
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skeletal muscle growth; inflammation; plasminogen system
Urokinase-type plasminogen activator (uPA) is an extracellular serine protease known to influence muscle regeneration following injury (23, 27). However, the role of uPA in muscle hypertrophy remains to be elucidated. uPA could contribute to hypertrophy through many different mechanisms. First, the classic function of uPA is to cleave plasminogen to form active plasmin, which, in turn, promotes extracellular matrix turnover either directly or through the activation of matrix metalloproteases (11, 41). Since muscle hypertrophy clearly requires matrix remodeling, uPA could contribute to hypertrophy in this manner. In addition, uPA binding to its membrane-bound receptor (uPAR) or to different integrin receptors can activate signaling pathways that may be important for hypertrophy (4, 10, 25, 35). Furthermore, uPA can regulate growth factor activity directly by proteolytic activation of HGF (31) and indirectly by activating plasmin, which can subsequently release growth factors from their extracellular matrix binding sites, increasing their bioactivity (37). Thus, the first purpose of this study was to determine whether uPA and its receptor uPAR are required for muscle hypertrophy.
Previous studies have indicated that skeletal muscle hypertrophy is associated with the accumulation of different cell types, some of which have been identified as inflammatory cells and fibroblasts by light and electron microscopy (1, 22). In addition, immunohistochemical analysis has demonstrated that different subpopulations of macrophages accumulate during mechanical load-induced muscle hypertrophy (39, 43). The classic functions of macrophages are to kill foreign organisms and to clear debris during tissue healing. However, macrophages also produce soluble factors that can enhance the proliferation of fibroblasts and muscle satellite cells (9, 12, 26, 29); satellite cells are thought to contribute to muscle growth by providing nuclei to growing muscle fibers. In a recent study (44) focused on recovery from muscle atrophy induced by hindlimb suspension, macrophage accumulation was reduced by repeated injections of an anti-F4/80 antibody, and this treatment appeared to impair recovery from atrophy. However, whether macrophages contribute to muscle hypertrophy remains to be determined. Thus, the second purpose of the present study was to determine whether macrophages are necessary for muscle hypertrophy.
For this study, we employed a widely used model of mechanical load-induced muscle growth involving surgical removal of all but one muscle of a synergist muscle group. This synergist ablation model produces a large and consistent compensatory increase in muscle mass, protein content, and force production (20, 21). Our data indicate that uPA and macrophages are indeed required for muscle growth.
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METHODS |
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C57BL/6 (wild-type) mice were obtained from Jackson or Harlan Laboratories, and uPA-null and uPAR-null mice were obtained from Jackson Laboratories on a C57BL/6 background and bred in our animal facility. No differences were found in responses to synergist ablation of wild-type mice obtained from the different suppliers or those bred in our facility. Mice were housed in a specific pathogen-free environment at a constant temperature and a 12:12-h light-dark cycle. Experiments were performed on 10- to 15-wk-old mice. All experimental procedures were approved by the Animal Care Committee of the University of Illinois (Chicago, IL).
Synergist Ablation
Bilateral synergist ablation surgery was performed on mice essentially as previously described (20, 21). Briefly, under ketamine (100 mg/kg) -xylazine (5 mg/kg) anesthesia and sterile conditions, gastrocnemius and soleus muscles were carefully removed, leaving the plantaris (PLT) muscle and its neurovascular supply intact. After recovery from surgery, mice remained ambulatory in cages for 1–14 days. Two hours before muscle collection, some mice were administered 30 mg/kg of 5-bromo-2'-deoxyuridine (BrdU) by an intraperitoneal injection for the assessment of cell proliferation. For sham operations, all surgical procedures were performed except the removal of muscles. Sham-operated mice were evaluated at 3 days postsurgery. For muscle collections, mice were killed by cervical dislocation while anesthetized with tribromoethanol (avertin; 400 mg/kg). PLT muscles were dissected, weighed, and either mounted in tissue-freezing medium and frozen in isopentane chilled with dry ice for histological assays (n = 6–12 muscles/group) or snap frozen in liquid nitrogen for biochemical assays (n = 6–10 muscles/group).
Clodronate Liposome Treatment
Clodronate liposomes have been widely used to deplete macrophages without affecting other cell types (46). Clodronate liposomes were prepared as previously described (46). Clodronate was a kind gift of Roche Diagnostics (Mannheim, Germany). Treated mice received 400-µl intraperitoneal injections immediately after synergist ablation and 2, 4, 6, 9, and 12 days postsurgery. A nonablated control group also received injections at the same time points.
Histology
Cross sections (10 µm thick) were cut from the midbelly of PLT muscles and either stained with hematoxylin and eosin for morphological analysis or processed for immunostaining for the quantification of inflammatory cells or proliferating cells.
Tissue edema. Images of two fields using a x40 lens objective were captured for two hematoxylin and eosin-stained sections per muscle (Labphot-2, Nikon; and SPOT software, Diagnostic Instruments). Each field was observed with an overlying 10 x 10 sampling grid. The percentage of intersection points that overlaid the extracellular space was used as an index of tissue edema. Sites containing sectioning artifacts were excluded.
Inflammatory cells. Immunohistochemistry was performed as previously described (34). Briefly, sections were air dried and then fixed in cold acetone. After being air dried again, sections were washed in PBS and quenched with 0.3% hydrogen peroxide. Sections were blocked with buffer containing 3% BSA and then incubated with primary antibodies for 2 h at room temperature. Macrophages were labeled with an F4/80 antibody (1:100, Serotec), and neutrophils were labeled with an Ly6G antibody (1:100, BD Biosciences). After being washed with PBS, sections were incubated with biotinylated mouse adsorbed anti-rat secondary antibody (1:200, Vector Laboratories). Sections were subsequently washed in PBS, incubated with avidin D-horseradish peroxidase (1:1,000), and developed using a 3-amino-9-ethylcarbazole kit (Vector Laboratories). Numbers of positively labeled cells were counted in three fields observed at x20 magnification under light microscopy for 2 sections/muscle and normalized to the volume of muscle sampled (area of section x section thickness).
Cell proliferation. After being fixed in cold acetone, sections were washed in PBS and incubated in 2 N HCl. Sections were neutralized with basic PBS (pH 8.5) and then washed with neutral PBS (pH 7.6). This was followed by an incubation in 0.1% IGEPAL and then in blocking buffer. Proliferating cells were labeled with a BrdU antibody (1:10, Roche Diagnostics) at room temperature. Sections were washed with PBS and subsequently incubated with FITC anti-mouse secondary antibody (1:200, Jackson ImmunoResearch). Numbers of positively labeled cells were counted in three fields observed at x20 magnification for 2 sections/muscle and normalized to the volume of muscle sampled.
Zymography
uPA activity in soluble fractions of PLT muscle homogenates was assessed using zymography as previously described (23). Muscles were homogenized in buffer [50 mM Tris·HCl (pH 7.6), 150 mM NaCl, and 0.5% Triton X-100] supplemented with protease inhibitors (5 mM EDTA, 1 mM PMSF, 1 µM leupeptin, and 0.3 µM aprotinin). Samples were centrifuged, and the soluble fraction was collected, mixed with loading buffer, and then separated on SDS-PAGE gels containing 
-casein (4 mg/ml) and human Glu-plasminogen (20 µg/ml). Gels were washed in 1% Tween 20 in PBS and incubated in 0.1% Tween 20 in PBS overnight at 37°C. Finally, gels were stained with Coomassie blue dye and subsequently destained overnight. uPA activity was identified by the position of its lytic band (45 kDa).
Myosin
Myosin concentration was determined from the insoluble fraction of the samples prepared for zymography. Samples were mixed with reducing SDS buffer and separated on SDS-PAGE gels. Gels were stained with Coomassie blue and then destained overnight. The optical density of myosin bands (
200 kDa) was measured and normalized to control values.
Cell Isolation
Cells were isolated from PLT muscles collected 5 days after ablation surgery using a protocol modified from the literature (3). Briefly, PLT muscles were dissected, minced, and then digested with 0.1% pronase (Calbiochem). After trituration to dissociate from fiber fragments, the suspension was filtered through 70-µm mesh, the filtrate was centrifuged, and cells were resuspended and counted. Cells were then separated into those that expressed the macrophage antigen Mac-1 and those that were negative for Mac-1 using microbeads conjugated to a Mac-1 antibody and a magnetic column (Miltenyi Biotec).
PCR
The expressions of uPA and IGF-I in Mac-1-positive and -negative cells isolated from PLT muscle were assessed at the mRNA level using RT-PCR. Total RNA was isolated from muscle or cells using the RNeasy kit (Qiagen) according to the manufacturer's instructions. RNA quantity was determined by UV absorption at 260 nm, and quality was verified by the 260-to-280-nm ratio and formaldehyde-agarose gel electrophoresis. mRNA (2 µg) was reverse transcribed using the Thermoscript RT-PCR kit (Invitrogen), and PCR was performed with the following primers: GAPDH, forward 5'-ACC ACA GTC CAT GCC ATC AC-3' and reverse 5'-TCC ACC ACC CTG TTG CTG GTA-3'; uPA, forward 5'-GCT CCT ATA ATC CTG GAG AGA TGA A-3' and reverse 5'-GGA GGG AAG AAG CTG AAA AGA CAG GT-3'; and IGF-I, forward 5'-ACA TCT CCC ATC TCT CTG GAT TTC CTT TTG C-3' and reverse 5'-CCC TCT ACT TGC GTT CTT CAA ATG TAC TTC C-3'.
Signaling
Western blots were used to analyze signaling pathways previously associated with muscle growth (19). PLT muscles were homogenized in buffer containing 40 mM Tris (pH 7.5), 1 mM EDTA, 5 mM EGTA, 0.5% Triton X-100, 25 mM 
-glycerophosphate, 25 mM NaF, 1 mM Na3VO4, 10 µg/ml leupeptin, and 1 mM PMSF. After homogenates had been centrifuged, the supernatant was mixed with reducing SDS buffer and separated on SDS-PAGE gels. Proteins were transferred to a polyvinylidene difluoride membrane, and membranes were blocked with 5% powdered milk and then incubated overnight at 4°C with phospho-specific primary antibodies against PKB (473), mTOR (2481) or p70 (389) (Upstate Biotechnology). Membranes were washed and then probed with horseradish peroxidase-conjugated secondary antibody at room temperature. Following another wash, blots were developed using an ECL kit (Amersham). Membranes were then stained with Coomassie blue to verify equal loading in all lanes. Densitometric measurements were carried out using the public domain NIH Image program (ImageJ).
Statistical Analysis
Values are reported as means ± SE. Data were compared across mouse strains and time points using one-way or two-way ANOVA, with Student-Newman-Keuls post hoc analysis. For groups that did not pass tests of normality and equal variance, the nonparametric Kruskal-Wallis one-way ANOVA on ranks was used with Dunn's post hoc method. Differences between groups were considered significant if P < 0.05.
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RESULTS |
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Hematoxylin and eosin-stained cryosections were used to assess changes in muscle edema in wild-type PLT muscle following synergist ablation (Fig. 1). There was a twofold increase in the amount of space between muscle fibers at 1 day postablation and a threefold increase at 3 days compared with control muscle, indicating tissue edema. Edema returned to control levels by 5 days postablation. There was no difference in edema at 3 days after sham surgery compared with control muscle (Fig. 1), suggesting that changes in edema were the result of increased mechanical load rather than surgical procedures. Of note, synergist ablation resulted in little overt damage to muscle fibers at all time points examined (e.g., little or no evidence of inflammatory cell invasion into muscle fibers, pale and/or discontinuous staining of the cytoplasm, or substantially swollen appearance of muscle fibers).
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uPA Activity Is Increased in Wild-Type Mice During Compensatory Hypertrophy
Casein zymograms were generated to evaluate the time course of uPA activity in PLT muscle of wild-type mice following synergist ablation (Fig. 1). Although uPA activity was barely detectable in control muscle, proteolytic activity increased at 1–14 days postablation. uPA activity appeared to peak at 5 days postablation but remained markedly elevated at 14 days. These data indicate that uPA activity is upregulated during compensatory hypertrophy and that uPA may play a role in both early and late events involved in hypertrophy.
Compensatory Hypertrophy Is Abrogated in uPA-Null Mice
In wild-type mice, synergist ablation resulted in a twofold increase in PLT muscle mass and a 75% increase in total protein at 14 days postablation compared with control muscle (Fig. 2). Myosin concentration in muscle collected at 14 days postablation was 90% of that in control muscle (data not shown). These data verify that there was substantial muscle hypertrophy in wild-type mice, with perhaps some deterioration in muscle quality, as noted in a previous study (21).
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Macrophage Accumulation Is Reduced in uPA-Null Mice
Next, we determined whether uPA influences inflammatory cell accumulation during compensatory hypertrophy. The numbers of macrophages were reduced by 1/3 in PLT muscle of uPA-null mice compared with wild-type mice at 3 and 5 days postablation, respectively (Fig. 3). Neutrophil accumulation demonstrated a delayed, transient reduction in PLT muscle of uPA-null mice compared with wild-type mice. Neutrophil accumulation was not reduced in uPA-null mice compared with wild-type mice at 1 and 5 days postablation but was reduced at 3 days. These data indicate that impaired muscle hypertrophy in uPA-null mice was associated with a sustained impairment of macrophage accumulation and transient reduction of neutrophils following synergist ablation.
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Since impaired muscle hypertrophy in uPA-null mice was associated with reduced macrophage accumulation, liposome-encapsulated clodronate was used to deplete macrophages following synergist ablation to determine whether macrophages are required for compensatory hypertrophy. To provide evidence that clodronate liposomes do not influence muscle cells directly, preliminary in vitro experiments were performed. C2C12 myoblasts or myotubes cultured as previously described (13) or bone marrow-derived macrophages cultured as previously described (47) were incubated with or without clodronate liposomes (200–600 µM) for 24 h. Clodronate liposomes induced death of all macrophages but had no effect on muscle cell numbers or protein content (not shown), indicating that the drug was indeed specific for macrophages.
For in vivo experiments, the accumulation of macrophages after synergist ablation was significantly reduced in clodronate liposome-treated mice compared with untreated mice (Fig. 3). Clodronate liposome treatment decreased macrophage accumulation by more than half at 3 and 5 days postablation. Although clodronate liposomes are thought to be specific for macrophages, there was also a delayed, transient reduction in neutrophil accumulation in treated mice. Neutrophil accumulation was not reduced in clodronate liposome-treated mice at 1 or 5 days postablation compared with untreated muscle but was reduced at 3 days in treated mice. These data indicate that clodronate liposomes were effective in reducing macrophage accumulation but also transiently reduced neutrophils following synergist ablation. This influence on neutrophils may have been secondary to decreased macrophage accumulation, since macrophage depletion has previously been found to reduce neutrophil accumulation during peritoneal inflammation (8), and it has been demonstrated that clodronate liposomes do not directly affect neutrophils (36).
Clodronate Liposomes Reduce Compensatory Hypertrophy in Wild-Type Mice
The increase in PLT muscle mass at 14 days postablation in clodronate liposome-treated mice was half of that in untreated control mice (Fig. 2). The increase in total protein was reduced by 2/3 in treated mice compared with untreated mice. The magnitude of the reduction in muscle hypertrophy was similar to the magnitude of reduction in macrophage accumulation with clodronate liposome treatment, indicating that macrophages are required for muscle hypertrophy following synergist ablation.
Macrophage Expression of uPA and IGF-I During Compensatory Hypertrophy
To determine whether macrophages contribute to the expression of uPA and other factors that may promote hypertrophy, macrophages were isolated from wild-type PLT muscle at 5 days postablation using microbeads coupled to an antibody against Mac-1. PCR analysis indicated that both Mac-1-positive and Mac-1-negative cells expressed uPA and IGF-I, two factors that have the potential to influence hypertrophy (Fig. 4). These data indicate that macrophages and other cells, possibly satellite cells, contribute to uPA and IGF-I expression following synergist ablation.
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BrdU incorporation was used to evaluate cell proliferation following synergist ablation (Fig. 5). BrdU incorporation peaked at 5 days postablation in PLT muscle of untreated wild-type mice (time course not shown), indicating the presence of a substantial number of proliferating cells. Clodronate liposome treatment resulted in 2/3 fewer BrdU-positive cells compared with muscle of untreated wild-type mice. Although there were 1/2 fewer proliferating cells in muscle of uPA-null mice, only a trend toward statistical significance was observed (P = 0.081). These data indicate that impaired cell proliferation may contribute to the reduced muscle growth that was observed with reduced macrophage accumulation.
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uPA has been shown to stimulate intracellular signaling through different molecules, including phosphatidylinositol 3-kinase (PI3K) and PKB (10, 15, 25). Since mTOR is thought to contribute to muscle hypertrophy and is considered to be downstream of PI3K and PKB (19, 20, 24), Western blots with phospho-specific antibodies were used to analyze whether uPA and/or macrophages influence the PKB-mTOR-p70 signaling pathway during compensatory muscle growth (Fig. 6). In nontreated wild-type mice, synergist ablation resulted in strong increases in phosphorylation of PKB-473, mTOR-2481, and p70–389. However, there were similar levels of phosphorylation of these signaling molecules in both uPA-null mice and clodronate-treated wild-type mice. These data indicate that the mechanism(s) by which macrophages and uPA influence muscle growth does not appear to include the mTOR signaling pathway.
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DISCUSSION |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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uPA could influence muscle hypertrophy through a number of different mechanisms. uPA has been demonstrated to stimulate the proliferation, migration, and fusion of satellite cells in vitro (5, 14, 30). Thus, uPA could stimulate satellite cell activity that may be required for compensatory muscle hypertrophy. uPA may stimulate activity of satellite cells and other cells by binding to cell surface receptors and activating intracellular signaling pathways (e.g., PI3K) and/or by increasing the activity of growth factors (e.g., HGF). Although our data indicate that uPAR is not required for muscle hypertrophy, uPA can bind to other cell surface receptors (e.g., integrins), and such binding could also influence intracellular signaling (10, 25). In addition, although our data indicate that uPA does not influence signaling through mTOR, other signaling pathways remain to be tested. Finally, uPA could regulate muscle hypertrophy through its influence on matrix remodeling. A well-known function of plasmin activation is turnover of the extracellular matrix both through direct proteolysis of a variety of matrix components and through the activation of matrix metalloproteases (11, 41). Whether matrix remodeling is an important function of uPA during muscle hypertrophy remains a question for further investigation.
In previous studies (23, 27) using different types of chemical injury, macrophage accumulation in damaged muscle was impaired in uPA-null mice, and this was associated with impaired muscle healing. In addition, in a recent study (7), we have found that neither inflammatory cell accumulation nor muscle regeneration is impaired in uPAR-null mice. The present study extends these findings, as our data demonstrate that macrophage accumulation during compensatory hypertrophy was impaired in uPA-null mice, and this was associated with impaired muscle hypertrophy. uPA could influence macrophage accumulation during muscle hypertrophy through effects on matrix proteolysis or signaling for migration (10, 11, 25).
Previous studies have reported evidence of inflammatory cell accumulation during compensatory muscle hypertrophy using light and electron microscopy, enzymatic assays, and immunohistochemistry. Compensatory growth has been associated with the accumulation of nuclei in the muscle interstitium, epimysium, and interfasicular spaces, along with an increase in edema, as observed using light microscopy of hematoxylin and eosin-stained sections (1). Although the authors suggested that these nuclei predominantly belong to neutrophils, other cell types could not be excluded. Compensatory muscle growth has also been associated with a large increase in macrophages and fibroblasts observed using electron microscopy, although the increase in these cells was not quantified (22). In both of these previous studies, growth was also associated with an increase in enzymatic markers of inflammatory cell activity. Finally, immunohistochemical analysis indicated that both ED1+ and ED2+ macrophages, different subpopulations of rat macrophages, were increased during compensatory muscle growth (39, 43). There are differences in opinion about whether the accumulation of inflammatory cells is due to surgical procedures or to increased mechanical loading during compensatory growth (1, 22, 39, 43). In the present study, immunohistochemical techniques were used to quantify the accumulation of both neutrophils, which peaked at 3 days postablation, and macrophages, which peaked at 5 days. In our experiments, sham surgery did not induce the accumulation of inflammatory cells (Fig. 1), indicating that increased mechanical load was likely responsible for the accumulation of inflammatory cells.
Previous studies using anti-inflammatory drugs have supported a role for inflammatory cells in muscle hypertrophy. The administration of ibuprofen resulted in reduced compensatory muscle growth in rats (39), and a specific cyclooxygenase-2 inhibitor reduced muscle recovery following atrophy in mice (6). In addition, ibuprofen reduced the increase in protein synthesis normally observed following increased mechanical loading of human skeletal muscle (45). Although these studies suggested that inflammatory cells may be involved in muscle growth, the drugs utilized may affect many different cells, including muscle cells (33, 38).
Other studies have specifically targeted macrophages with different approaches to determine the influence of these inflammatory cells in muscle remodeling. In a study (44) focused on recovery from muscle atrophy induced by hindlimb suspension, macrophage accumulation was reduced by repeated injections of an anti-F4/80 antibody, and this treatment reduced the formation of regenerating muscle fibers and impaired recovery of the fiber area to control levels. Following freeze injury, macrophage depletion with clodronate liposomes resulted in impaired muscle regeneration associated with delayed clearance of necrotic muscle fibers and interstitial fat accumulation (42). In the latter study, although the formation of regenerating fibers appeared to be impaired, the areas of the regenerating fibers that were formed did not differ between control and macrophage-depleted mice.
Our data extend the findings of these previous studies; we used clodronate liposomes to deplete macrophages during compensatory hypertrophy, and our results demonstrate that macrophage depletion resulted in decreased muscle growth. Clodronate liposomes had the intended effect of a sustained reduction in macrophage accumulation but also produced a delayed, transient reduction in neutrophil accumulation. This delayed reduction in neutrophils could have been a result of decreased macrophage accumulation, since macrophage depletion has previously been found to reduce neutrophil accumulation during peritoneal inflammation (8), and it has been demonstrated that clodronate liposomes do not directly affect neutrophils (36). Whether neutrophils can directly influence muscle hypertrophy remains to be determined; however, these data indicate that macrophages are required for compensatory hypertrophy.
Macrophages produce a variety of factors that have the potential to influence muscle growth, including proteases, growth factors, and cytokines (18, 32). These factors could influence growth through the regulation of the activity of muscle satellite cells and fibroblasts, growth of mature muscle fibers, and/or remodeling of connective tissue. Our findings indicate that factors expressed by macrophages during compensatory muscle hypertrophy include uPA and IGF-I. IGF-I has been previously implicated as an important factor in muscle hypertrophy (17, 28), and our data indicate that macrophages are a source of IGF-I during compensatory hypertrophy. Macrophage-derived factors have been shown previously to stimulate proliferation of fibroblasts and satellite cells in vitro (9, 12, 26, 29). In the present study, macrophage depletion in vivo resulted in reduced cell proliferation during muscle hypertrophy (Fig. 5). Although the affected cells have not yet been identified, candidate cell types include satellite cells and fibroblasts as well as macrophages. Future studies will be focused on identifying the proliferating cell types in muscle during hypertrophy and probing the mechanisms by which macrophages and uPA regulates the proliferation of these cells.
In conclusion, uPA and macrophages are required for muscle hypertrophy in response to mechanical loading. These results enhance our understanding of the role of nonmuscle cells and extracellular processes in muscle growth. The findings also indicate that manipulating macrophage function and/or uPA activity could represent potential therapeutic approaches for improving muscle growth.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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