Mice deficient in plasminogen activator inhibitor-1 have improved skeletal muscle regeneration

Timothy J. Koh,1 Scott C. Bryer,1 Augustina M. Pucci,1 and Thomas H. Sisson2

1Department of Movement Sciences, University of Illinois at Chicago, Chicago, Illinois; and 2Department of Internal Medicine, University of Michigan, Ann Arbor, Michigan

Submitted 17 November 2004 ; accepted in final form 11 February 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Skeletal muscle possesses a remarkable capacity for regeneration. Although the regulation of this process at the molecular level remains largely undefined, the plasminogen system appears to play a critical role. Specifically, mice deficient in either urokinase-type plasminogen activator (uPA–/– mice) or plasminogen demonstrate markedly impaired muscle regeneration after injury. In the present study, we tested the hypothesis that loss of the primary inhibitor of uPA, plasminogen activator inhibitor-1 (PAI-1), would improve muscle regeneration. Repair of the extensor digitorum longus muscle was assessed after cardiotoxin injury in wild-type, uPA–/–, and PAI-1-deficient (PAI-1–/–) mice. As expected, there was no uPA activity in the injured muscles of uPA–/– mice, and muscles from these transgenic animals demonstrated impaired regeneration. On the other hand, uPA activity was increased in injured muscle from PAI-1–/– mice to a greater extent than in wild-type controls. Furthermore, PAI-1–/– mice demonstrated increased expression of MyoD and developmental myosin after injury as well as accelerated recovery of muscle morphology, protein levels, and muscle force compared with wild-type animals. The injured muscles of PAI-1-null mice also demonstrated increased macrophage accumulation, contrasting with impaired macrophage accumulation in uPA-deficient mice. The extent of macrophage accumulation correlated with both the clearance of protein after injury and the efficiency of regeneration. Taken together, these results indicate that PAI-1 deficiency promotes muscle regeneration, and this protease inhibitor represents a therapeutic target for enhancing muscle regeneration.

muscle injury; muscle repair; urokinase-type plasminogen activator; muscle inflammation; macrophage


SKELETAL MUSCLE POSSESSES an extraordinary capacity for regeneration after injury caused by disease, trauma, or intense exercise. The process of regeneration can be separated into three overlapping phases. The first phase is marked by the accumulation of inflammatory cells that remove tissue debris and may play further critical roles in regeneration (10, 18, 24, 40). The second phase involves the activation and proliferation of quiescent muscle precursor cells called satellite cells, which migrate and fuse to replace damaged fibers (7, 16, 32). The third phase involves the growth of newly formed fibers and remodeling of damaged fibers, a process highlighted by the synthesis of proteins to form new sarcomeres and associated structures. Although much has been learned about the events involved in muscle regeneration, the molecular mechanisms that regulate this process remain largely undefined.

Many studies have demonstrated that the plasminogen system plays an important role in the repair of skeletal muscle as well as other tissues (19, 31, 3335, 43). The plasminogen system is a serine protease cascade that includes the urokinase-type plasminogen activator (uPA). The proteolytic activity of uPA is tightly regulated by its primary inhibitor, plasminogen activator inhibitor-1 (PAI-1). The classic function of uPA is to activate plasminogen to plasmin, and plasmin possesses broad spectrum protease activity against a variety of extracellular matrix molecules, including fibrin, fibronectin, and proteoglycans (37). Plasmin activity may be required to remove fibrin clots after injury and clear a path to allow the migration of different cells to the site of injury. uPA and PAI-1 may also contribute to tissue repair through pathways that do not involve plasminogen, including the regulation of growth factor activity and cell migration (6, 27, 29, 36, 41).

Recent studies support the idea that uPA and PAI-1 play critical roles in muscle regeneration. In vitro uPA appears to stimulate satellite cell proliferation, migration, and fusion (2, 12, 13, 25). In vivo muscle injury results in increased expression of uPA (11, 19), and mice with a targeted deletion of uPA or plasminogen show impaired expression of myogenic factors and markedly deficient muscle regeneration (19, 38). Muscle injury also results in increased expression of PAI-1 (9, 11, 44), which, through the inhibition of uPA activity, could limit the efficiency of muscle regeneration. Thus the main hypothesis of this study was that PAI-1 deficiency results in increased uPA activity and improved muscle regeneration. On the other hand, antibodies that block PAI-1's interaction with uPA were found to impair satellite cell migration and fusion in vitro (2, 8). On the basis of this result, one could speculate that the presence of PAI-1 in the injured muscle is required for efficient in vivo regeneration. Thus the alternative hypothesis for this study was that PAI-1 deficiency results in impaired muscle regeneration. The available data make it difficult to predict whether PAI-1 deficiency would promote or impair muscle regeneration. To explore the effect of PAI-1 on muscle repair, we injured the extensor digitorum longus (EDL) muscle in wild-type, uPA-deficient (uPA–/–), and PAI-1-deficient (PAI-1–/–) mice and assessed the efficiency of repair with measurements of muscle function, morphology, and protein levels. We also assessed inflammatory cell accumulation to determine whether PAI-1 regulates the inflammatory response after injury.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Mice. uPA- and PAI-1-null mice were originally generated by Dr. Peter Carmeliet (4, 5). Both knockout strains had been bred into a C57BL/6 background for a minimum of eight generations. C57BL/6 (wild-type) mice were obtained from Harlan (Indianapolis, IN). Experiments were performed on mice aged 10–12 wk. All experimental procedures were approved by the University Animal Care and Use Committee of the University of Michigan and the Animal Care Committee at the University of Illinois at Chicago.

Cardiotoxin injection and muscle function assay. EDL muscle force measurements and cardiotoxin injuries were performed using standard procedures (17, 44). Briefly, mice were anesthetized with an intraperitoneal injection of tribromoethanol (avertin; 400 mg/kg) and fixed on a heated Plexiglas platform (37°C). Preinjury in situ maximal isometric force was measured as described previously (17), and then 10 µl of cardiotoxin (10 µM; Calbiochem, San Diego, CA) were injected into the EDL muscle in three locations to ensure distribution of cardiotoxin throughout the muscle. After injection, the skin incision was closed with 7-0 nylon suture, and the procedures were repeated in the contralateral muscle. Mice were allowed to recover and then were subjected to a postinjury muscle function test using the same muscle-testing procedures at 1–20 days postinjury.

Sample preparation. After the terminal muscle function assay, EDL muscles were dissected, blotted dry, weighed, and processed for histological or biochemical analysis. For histological assays, muscles were mounted in tissue-freezing medium and frozen in isopentane chilled with dry ice. For biochemical assays, muscles were homogenized in buffer A (50 mM Tris·HCl, pH 7.4, 150 mM NaCl, 0.5% Triton X-100, and 5 mM EDTA) supplemented with protease inhibitors (1 mM phenylmethylsulfonyl fluoride, 1 µM leupeptin, and 0.3 µM aprotinin). Samples were centrifuged at 15,000 g for 10 min at 4°C, and then the soluble fraction was collected and the insoluble fraction was resuspended in buffer A.

Histology. Cross sections were cut 10 µm from the midbelly of each EDL muscle and either stained with hematoxylin and eosin for morphological analysis or processed for immunohistochemistry for inflammatory cell analysis. For morphological analysis, images of two fields using a x20 lens objective were captured for each muscle section (Labphot-2, Nikon; and SPOT software, Diagnostic Instruments, Sterling Heights, MI). For each field, fibers were classified as normal, damaged, or regenerating, and the number and area of these fibers were recorded. Normal fibers were identified as those demonstrating no clear evidence of damage, damaged fibers were identified as those demonstrating overt damage (e.g., infiltration of inflammatory cells, pale and/or discontinuous staining of the cytoplasm, or substantially swollen appearance), and regenerating fibers were identified as those containing centrally located nuclei without evidence of damage. The damaged area of each muscle section was estimated by subtracting the summed area of normal and regenerating fibers from the total area of each field.

Analysis of inflammatory cells was performed using immunohistochemical methods described previously (28). Neutrophils were labeled with a Ly6G antibody (1:100 dilution; Pharmingen, San Diego, CA), and macrophages were labeled with an F4/80 antibody (1:100 dilution; Serotec, Oxford, UK), followed by incubation with biotinylated mouse adsorbed anti-rat IgG (1:200 dilution; Vector Laboratories, Burlingame, CA) and then with avidin D horseradish peroxidase (1:1,000 dilution). Sections were then developed using the AEC kit (Vector Laboratories). The number of positive cells were counted in two entire sections for each muscle with the aid of an eyepiece grid and normalized to the volume of muscle sampled (area of section x section thickness; 10 µm).

Zymography. uPA activity in soluble fractions of EDL muscle homogenates was assessed using zymography as described previously (34). Briefly, protein content of the soluble fraction was determined using a method described previously (23), and then 10 µg of each sample were separated on 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels containing {alpha}-casein (4 mg/ml) and human Glu-plasminogen (20 µg/ml). Gels were then washed in 1% Tween 20 in PBS for 1 h at 37°C and then incubated in 0.1% Tween 20 in PBS overnight at room temperature. Finally, gels were stained with Coomassie blue dye and then destained. Murine uPA activity was identified by the position of its lytic band (45 kDa) relative to that of a recombinant human uPA standard (55 kDa).

Fibrinolysis assay. uPA activity was also measured quantitatively in EDL muscle homogenates using a microplate fibrinolysis assay. Samples (10 µg of protein) were mixed with plasminogen (0.5 µM final concentration) and fibrinogen (8 µM final concentration). Thrombin was added to a final concentration of 0.4 U/ml to initiate fibrin clot formation. Clot formation was followed by the increase in absorbance at 405 nm, and fibrinolysis resulted in the subsequent decrease in optical density (OD) 405. The time required for the OD 405 to fall to one-half its maximal value was taken as a measure of uPA activity (t1/2). Control experiments were performed without the addition of thrombin and without the addition of sample.

Protein levels. Aliquots of both soluble and insoluble fractions of EDL muscle homogenates were mixed with concentrated reducing sample buffer, boiled, and protein concentrations and total muscle protein content were determined (23). Equal amounts of protein (1 µg for fast myosin heavy-chain isoform; 10 µg for developmental myosin heavy-chain isoform) were separated on 8% SDS-PAGE gels and transferred onto nitrocellulose membranes. After transfer, membranes were stained with Ponceau S to confirm equal loading and then blocked in 5% milk overnight. Membranes were incubated with primary antibodies against MyoD (1:500 dilution; Novocastra, Newcastle upon Tyne, UK), developmental myosin heavy chain (1:500 dilution; Novocastra), or fast myosin heavy chain (1:5,000 dilution; Sigma, St. Louis, MO); washed; and then incubated with secondary antibody conjugated to horseradish peroxidase (1:25,000 dilution; Pierce Biotechnology, Rockford, IL). After another wash, protein bands were detected using enhanced chemiluminescence (Amersham, Little Chalfont, UK) and band densities were determined using image analysis (Fluor-S; Bio-Rad, Hercules, CA). Individual band densities were normalized to the average of the control lanes for wild-type mice and multiplied by 100%.

Data analysis. Values are reported as means ± SE. Data were compared across different mouse strains and time points using two-way ANOVA. Post hoc tests were performed using the Student-Newman-Keuls test. The 0.05 level was used to indicate statistical significance.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
uPA activity. Casein zymograms were obtained to assess the time course of uPA activity in injured muscle (Fig. 1). Although uPA was barely detectable in noninjured muscle, proteolytic activity was markedly increased on days 1, 3, and 5 postinjury in both wild-type and PAI-1-null mice. Thereafter uPA activity decreased nearly to control levels by day 20 postinjury. As expected, muscle homogenates from uPA-null mice showed no uPA activity at any time point. Of note, muscle injury also resulted in an increase in activity of tissue-type plasminogen activator (tPA) in all mouse types, with a time course that lagged behind that of uPA. A microplate fibrinolysis assay was performed to quantify uPA activity in muscles from wild-type and PAI-1-null mice at day 1 postinjury. Injured muscle from PAI-1-null mice showed greater fibrinolytic activity (fibrin clot dissolution t = 211 ± 15 min) than muscle from wild-type mice (t = 374 ± 54 min). Increased fibrinolytic activity in injured muscle of PAI-1-null mice is consistent with loss of uPA inhibition.



View larger version (52K):
[in this window]
[in a new window]
 
Fig. 1. Urokinase-type plasminogen activator (uPA) activity is upregulated after muscle injury. Top: gel zymographic analysis of uPA and tissue-type plasminogen activator (tPA) activities in muscles of wild-type (WT), uPA-null (uPA–/–), and plasminogen activator inhibitor-1 (PAI-1)-null (PAI-1–/–) mice at different time points after cardiotoxin injury. Plasminogen activator activity forms clearing bands in zymogram gels. Note the increase in uPA activity after muscle injury in wild-type and PAI-1–/– mice and the lack of clearing bands for uPA–/– mice. Note also the increase in tPA activity that lags the increase in uPA. The identity of uPA and tPA bands was determined using protein standards (data not shown).

 
Muscle function. Muscle force production was measured after injury to assess recovery of muscle function (Fig. 2). Before injury, specific tension (muscle force divided by physiological cross-sectional area) was not different between muscles of wild-type, uPA-null, and PAI-1-null mice (~22 N/cm2), indicating that muscle function did not differ between genotypes before injury. On days 1 and 3 postinjury, maximal isometric muscle force for each mouse strain was reduced to ~20% of control values, indicating that cardiotoxin-induced injury to the EDL muscle was independent of genotype. On day 5 postinjury, only muscles from PAI-1-null mice showed a significant recovery of force. By days 10 and 20 postinjury, wild-type mice showed substantial improvement, and PAI-1-null mice continued to exhibit significantly greater recovery of force than wild-type mice. On the other hand, uPA-null mice showed little, if any, recovery over the time course of the experiment.



View larger version (26K):
[in this window]
[in a new window]
 
Fig. 2. Recovery of muscle function after injury is accelerated in PAI-1-null mice and impaired in uPA-null mice. Maximal isometric force was measured before and at different time points after cardiotoxin injury of the extensor digitorum longus (EDL) muscle. Muscle force is expressed as %preinjury value. Data are means ± SE (bars represent SE; n = 4–10 no. of muscle samples studied). Two-way ANOVA showed a significant interaction effect of mouse strain by number of days after injury. 1, mean value significantly greater than that for 3 days (within strain); 2, mean value significantly greater than that for wild-type mice (within day); 3, mean value significantly smaller than that for wild-type mice (within day).

 
Muscle morphology. Hematoxylin and eosin-stained cryosections were used to assess changes in muscle morphology after injury (Fig. 3). In noninjured muscle, muscle fiber morphology showed no overt differences between wild-type, uPA-null, and PAI-1-null mice. Furthermore, fiber area was not different between genotypes (923 ± 36 µm), suggesting no differences in muscle fiber development. On day 3 after cardiotoxin injection, cross sections of EDL muscles from each mouse type showed substantial fiber damage and edema. The amount of damage (~90% of total area in cryosections) was indistinguishable between the three genotypes. On day 5 postinjury, wild-type mice demonstrated a significant reduction in damaged area. At the same time point, PAI-1-null mice showed a significantly greater reduction in damaged area compared with wild-type mice, consistent with greater force production. By days 10 and 20 postinjury, damaged area in both wild-type and PAI-1-null mice approached zero. In contrast, uPA-null mice showed little, if any, reduction in the damaged area up to 20 days postinjury.



View larger version (69K):
[in this window]
[in a new window]
 
Fig. 3. Recovery of normal morphology after injury is accelerated in muscles of PAI-1-null mice and impaired in muscles of uPA-null mice. Top: hematoxylin and eosin-stained cryosections from uninjured control muscle (C) and from muscle at different time points after injury in wild-type (WT), uPA-null (uPA–/–), and PAI-1-null mice (PAI-1–/–). Note the robust regenerative response in muscle of wild-type and PAI-1-null mice and the absence of regeneration in muscle of uPA-null mice. Bottom: damaged area estimated by subtracting area of normal and regenerating fibers from total area of two fields per section. Damaged area is expressed as %total area. Data are means ± SE (bars represent SE; n = 4–8 no. of muscle samples studied). Two-way ANOVA showed significant interaction effect of mouse strain by day after injury. 1, mean value significantly smaller than that for 3 days (within strain); 2, mean value significantly greater than that for wild-type mice (within day); 3, mean value significantly less than that for wild-type mice (within day).

 
Protein levels. Levels of total protein and fast myosin heavy chain were measured to assess muscle protein turnover after muscle injury (Fig. 4). In muscles from both wild-type and PAI-1-null mice, total protein and fast myosin heavy chain levels decreased rapidly after injury, reaching a nadir on day 3. Muscles from uPA-null mice demonstrated little change in protein levels during this period, suggesting impaired clearance of damaged tissue. During the subsequent phase of regeneration, muscles of wild-type mice demonstrated increases in total protein and fast myosin heavy chain to near-normal levels by day 20 postinjury. PAI-1-null mice showed accelerated recovery of muscle protein levels compared with their wild-type counterparts, consistent with accelerated recovery of function and morphology. In contrast, uPA-null mice exhibited a gradual loss of muscle protein up to day 20. Together, these data show that protein turnover was impaired in injured muscles of uPA-null mice and that protein accretion was accelerated during the regenerative phase in PAI-1-null mice.



View larger version (61K):
[in this window]
[in a new window]
 
Fig. 4. Recovery of total protein and adult myosin heavy-chain levels after injury is accelerated in muscles of PAI-1-null mice and impaired in muscles of uPA-null mice. Left: total protein levels in injured muscle of wild-type, uPA-null, and PAI-1-null mice at different time points after injury were normalized to protein level in noninjured control muscle. Right: fast myosin heavy-chain (MHCf) protein levels measured using densitometry of Western blots (top row) and expressed relative to levels in noninjured control muscle. Data are means ± SE (bars represent SE; n = 4–8 no. muscle samples studied). Two-way ANOVA showed significant interaction effect of mouse strain by day after injury for both total protein and fast myosin heavy-chain levels. 1, mean value significantly different from control (within strain); 2, mean value significantly greater than that for wild-type mice (within day); 3, mean value significantly smaller than that for wild-type mice (within day).

 
Myogenesis. Myogenesis after injury was assessed by measuring the expression of the myogenic transcription factor MyoD and the developmental isoform of the myosin heavy chain using Western blot analysis and by counting the number of central nucleated fibers in muscle sections (Fig. 5). On day 1 postinjury, protein expression of MyoD was significantly greater in injured muscle of PAI-1-null mice than in wild-type mice. Similarly, the expression of developmental myosin heavy chain was significantly greater in injured muscle of PAI-1-null mice than in that of wild-type mice on day 5 postinjury. In contrast, developmental myosin protein expression was significantly lower in injured muscles of uPA-null mice than in those of wild-type mice. Also on day 5 postinjury, muscle sections from wild-type and PAI-1-null mice showed substantial numbers of central nucleated fibers. Mean values for PAI-1-null mice were greater than those of wild-type mice, although the difference did not reach statistical significance (P = 0.08). On the other hand, muscle sections from uPA-null mice showed significantly fewer central nucleated fibers after injury than those from wild-type mice.



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 5. Expression of MyoD and developmental myosin heavy chain isoform in muscles of wild-type, uPA-null, and PAI-1-null mice. Top: MyoD protein levels at 1 day postinjury were measured using densitometry of Western blots and expressed relative to levels in muscles of wild-type mice. Middle: developmental myosin heavy-chain (MHCd) protein levels at 5 days postinjury measured using densitometry of Western blots and expressed relative to levels in muscles of wild-type mice. Bottom: number of central nucleated fibers counted per cryosection for muscles of WT, uPA–/–, and PAI-1–/– mice at 5 days postinjury normalized to total area of two fields per section. Central nucleated fibers were considered to be regenerating fibers. Data are means ± SE (bars represent SE; n = 4–8 no. of muscle samples studied). One-way ANOVA showed effects of mouse strain for both central nucleated fibers and developmental myosin heavy-chain expression. 1, mean value significantly smaller than that for wild-type mice; 2, mean value significantly greater than that for wild-type mice.

 
Inflammatory cells. Immunohistochemical methods were used to quantify inflammatory cell accumulation in injured muscle (Fig. 6). Neutrophil accumulation peaked on day 1 postinjury in each mouse strain, and the number of neutrophils did not differ significantly between genotypes. In the muscles of wild-type mice, macrophage accumulation peaked on day 3 and then declined by day 5 postinjury. In muscles of PAI-1-null mice, there was a trend toward increased macrophage accumulation on day 3 postinjury (P = 0.08), and there was significant accumulation at day 5 postinjury compared with wild-type mice. In contrast, the accumulation of macrophages was significantly impaired in injured muscles of uPA-null mice compared with wild-type mice; the number of macrophages was <5% of those found in wild-type mice on day 3 postinjury.



View larger version (29K):
[in this window]
[in a new window]
 
Fig. 6. Macrophage accumulation is impaired in injured muscle of uPA-null mice and increased in injured muscle of PAI-1-null mice. Top: neutrophil accumulation assessed at different time points after muscle injury in wild-type, uPA-null, and PAI-1-null mice using immunostaining for the neutrophil-specific Ly6G antigen, counting the number of Ly6G+ cells per cryosection, and normalizing to cryosection volume. Bottom: macrophage accumulation assessed using immunostaining for the macrophage-specific F4/80 antigen, counting the number of F4/80+ cells per cryosection, and normalizing to cryosection volume. Data are means ± SE (bars represent SE; n = 4–8 no. of muscle samples studied). Two-way ANOVA showed a significant effect of day for neutrophils (all time points significantly greater than controls) and a significant interaction effect of mouse strain by day after injury for macrophages. 1, mean value significantly different from control (within strain); 2, mean value significantly greater than that for wild-type mice (within day); 3, mean value significantly smaller than that for wild-type mice (within day).

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Although skeletal muscle regeneration typically is efficient, we have found that manipulation of the plasminogen system can further enhance muscle repair. Specifically, mice deficient in PAI-1 demonstrated increased MyoD and developmental myosin expression after injury, as well as accelerated recovery of muscle morphology, protein levels, and function, compared with wild-type mice. The more rapid muscle regeneration in PAI-1-null mice was also associated with an increased accumulation of macrophages after injury. As anticipated on the basis of previous studies (19, 38), we also found that mice deficient in uPA had nearly complete abrogation of muscle fiber regeneration and macrophage accumulation after injury. Taken together, our data indicate that PAI-1 deficiency accentuates plasminogen activator activity and promotes muscle regeneration.

The balance of uPA and PAI-1 may influence muscle regeneration through many pathways. Previous in vitro studies have demonstrated that uPA and PAI-1 can directly influence myogenesis. Exogenous uPA stimulated proliferation of human satellite cells, and antibodies that prevented interaction between uPA and its receptor (uPAR) blocked this increase in proliferation (12). These same antibodies also blocked a fibroblast growth factor (FGF)-stimulated increase in satellite cell proliferation, indicating that the uPA system may play a role in growth factor-mediated proliferation (13). In addition, exogenous uPA stimulated migration of human satellite cells through a matrix composed of basement membrane components (Matrigel), and antibodies that blocked binding of uPA to uPAR inhibited both exogenous uPA-dependent and exogenous FGF-dependent migration (12, 13). Finally, uPA activity may be required for satellite cell fusion, because in vitro studies have shown that uPA activity is increased during this process and antibodies that block the catalytic activity of uPA or the interaction of uPA with uPAR have been found to inhibit fusion (2, 25). Taken together, these in vitro results demonstrate that uPA is capable of modulating satellite cell proliferation, migration, and fusion. Furthermore, these processes appear to depend on both the proteolytic (e.g., plasmin formation, growth factor activation) and the nonproteolytic (e.g., uPAR binding, matrix binding) functions of uPA. In the present study, injured muscle of PAI-1–/– mice demonstrated increased uPA activity and increased expression of developmental muscle proteins compared with that observed in wild-type mice. Whether the enhanced expression of these proteins resulted from a direct effect of uPA on muscle cells, and whether it required proteolytic or nonproteolytic uPA functions, remains to be determined.

In addition to influencing myogenesis directly, uPA and PAI-1 may modulate regeneration by regulating the inflammatory response. uPA, PAI-1, and uPAR have been reported to be important in the regulation of inflammatory cell activation and infiltration into injured tissues through both proteolytic and nonproteolytic mechanisms (1, 6, 14, 15). In the present study, macrophage accumulation was largely abrogated in uPA–/– mice, whereas PAI-1 deficiency augmented this process. Impaired macrophage accumulation in damaged muscle was previously reported in uPA–/– and plasminogen–/– mice (19, 38). The mechanism by which macrophage accumulation is altered in these transgenic animals is unclear, but differences in the level of activation, the ability to migrate, and/or the levels of chemotactic factors could be important. After muscle injury, macrophages appear to be critical for both clearance of damaged tissue and promotion of muscle regeneration (10, 18, 24, 40). In support of the latter role, isolated macrophages, presumably through the release of soluble factors, have been found to stimulate satellite cell proliferation and migration in vitro (3, 22, 30).

uPA and PAI-1 may also modulate muscle regeneration through the regulation of extracellular matrix turnover. Through the activation of plasminogen, uPA promotes the degradation of several extracellular matrix proteins, including fibrin, fibronectin, and proteoglycans (37). Furthermore, plasmin can also activate a subset of matrix metalloproteinases, which can degrade other matrix components, including collagen, elastin, and laminin (6, 26). Previous investigators (19, 38) have reported the persistence of intramuscular fibrin deposition after injury in both uPA–/– and plasminogen–/– mice. Furthermore, systemic defibrinogenation with ancrod limited fibrin deposition and, in turn, improved muscle regeneration in these transgenic mice. The removal of extracellular matrix barriers may be necessary for efficient macrophage and satellite cell migration during muscle regeneration. Regardless of whether it is impaired, fibrin removal as the primary cause of delayed regeneration in uPA–/– mice requires further study.

In addition to matrix turnover, uPA and PAI-1 may influence muscle repair by regulating the bioactivity of a variety of growth factors. For example, plasmin can activate latent transforming growth factor-{beta} (TGF-{beta}) and enhance the bioactivity of FGF-2 by releasing it from its extracellular matrix reservoir (20, 29). uPA is also capable of activating growth factors directly, without the involvement of plasminogen. Specifically, uPA can cleave inactive single-chain hepatocyte growth factor (HGF) to generate the active two-chain form (21, 27). Because HGF and FGF are thought to play key roles in the regulation of satellite cell activity (39, 42), regulation of their activity could be an important mechanism by which uPA and PAI-1 influence muscle repair.

tPA activity was also increased after cardiotoxin injection in wild-type mice with a time course that lagged behind the increase in uPA activity. On the basis of this finding, it is possible that tPA also participates in the regenerative response after muscle injury. However, a previous study (19) demonstrated that muscle repair is not qualitatively different in tPA-deficient mice compared with wild-type controls, demonstrating that tPA is not required for efficient muscle regeneration.

Efficient muscle regeneration helps to restore muscle function after injury, but repair is inadequate to prevent progressive degeneration in the setting of muscle diseases such as Duchenne muscular dystrophy. Furthermore, the efficiency of muscle repair declines with age. Our results suggest that PAI-1 may be an attractive therapeutic target in patients with insufficient regenerative capacity in their skeletal muscles. The mechanism by which PAI-1 deficiency accelerates muscle regeneration remains unclear and requires further study. Because macrophages appear to be critical to this process, we speculate that PAI-1 deficiency may promote muscle repair, at least in part, by modulating the inflammatory response after injury. This hypothesis is currently being addressed in ongoing experiments. A further definition of the mechanisms by which uPA and PAI-1 influence muscle regeneration may provide insight into improving regeneration in patients with muscle diseases and in older adults. Furthermore, because uPA and PAI-1 have been implicated in the repair of other tissues, elucidating the mechanisms of action of this system in muscle regeneration may provide insights into improving the repair of other tissues that do not efficiently regenerate after injury (e.g., heart).


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This project was supported in part by National Heart, Lung, and Blood Institute Grant HL-04434 (to T. H. Sisson). S. C. Bryer was supported by a National Aeronautics and Space Administration Graduate Student Research Fellowship.


    ACKNOWLEDGMENTS
 
We thank Kerstin Hanson and Bi Yu for technical assistance.


    FOOTNOTES
 

Address for reprint requests and other correspondence: T. J. Koh, Dept. of Movement Sciences (M/C 194), Univ. of Illinois at Chicago, 901 W. Roosevelt Rd., Chicago, IL 60608 (e-mail: tjkoh{at}uic.edu)

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


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Abraham E, Gyetko MR, Kuhn K, Arcaroli J, Strassheim D, Park JS, Shetty S, and Idell S. Urokinase-type plasminogen activator potentiates lipopolysaccharide-induced neutrophil activation. J Immunol 170: 5644–5651, 2003.[Abstract/Free Full Text]

2. Bonavaud S, Charrière-Bertrand C, Rey C, Leibovitch MP, Pedersen N, Frisdal E, Planus E, Blasi F, Gherardi R, and Barlovatz-Meimon G. Evidence of a non-conventional role for the urokinase tripartite complex (uPAR/uPA/PAI-1) in myogenic cell fusion. J Cell Sci 110: 1083–1089, 1997.[Abstract/Free Full Text]

3. Cantini M, Massimino ML, Bruson A, Catani C, Dalla Libera L, and Carraro U. Macrophages regulate proliferation and differentiation of satellite cells. Biochem Biophys Res Commun 202: 1688–1696, 1994.[CrossRef][ISI][Medline]

4. Carmeliet P, Kieckens L, Schoonjans L, Ream B, van Nuffelen A, Prendergast G, Cole M, Bronson R, Collen D, and Mulligan RC. Plasminogen activator inhibitor-1 gene-deficient mice. I. generation by homologous recombination and characterization. J Clin Invest 92: 2746–2755, 1993.[ISI][Medline]

5. Carmeliet P, Schoonjans L, Kieckens L, Ream B, Degen J, Bronson R, De Vos R, van den Oord JJ, Collen D, and Mulligan RC. Physiological consequences of loss of plasminogen activator gene function in mice. Nature 368: 419–424, 1994.[CrossRef][ISI][Medline]

6. Chapman HA Jr, Reilly JJ Jr, and Kobzik L. Role of plasminogen activator in degradation of extracellular matrix protein by live human alveolar macrophages. Am Rev Respir Dis 137: 412–419, 1988.[ISI][Medline]

7. Chargé SBP and Rudnicki MA. Cellular and molecular regulation of muscle regeneration. Physiol Rev 84: 209–238, 2004.[Abstract/Free Full Text]

8. Chazaud B, Bonavaud S, Plonquet A, Pouchelet M, Gherardi RK, and Barlovatz-Meimon G. Involvement of the [uPAR:uPA:PAI-1:LRP] complex in human myogenic cell motility. Exp Cell Res 258: 237–244, 2000.[CrossRef][ISI][Medline]

9. Chen YW, Nader GA, Baar KR, Fedele MJ, Hoffman EP, and Esser KA. Response of rat muscle to acute resistance exercise defined by transcriptional and translational profiling. J Physiol 545: 27–41, 2002.[Abstract/Free Full Text]

10. Farges MC, Balcerzak D, Fisher BD, Attaix D, Béchet D, Ferrara M, and Baracos VE. Increased muscle proteolysis after local trauma mainly reflects macrophage-associated lysosomal proteolysis. Am J Physiol Endocrinol Metab 282: E326–E335, 2002.[Abstract/Free Full Text]

11. Festoff BW, Reddy RB, VanBecelaere M, Smirnova I, and Chao J. Activation of serpins and their cognate proteases in muscle after crush injury. J Cell Physiol 159: 11–18, 1994.[CrossRef][ISI][Medline]

12. Fibbi G, Barletta E, Dini G, Del Rosso A, Pucci M, Cerletti M, and Del Rosso M. Cell invasion is affected by differential expression of the urokinase plasminogen activator/urokinase plasminogen activator receptor system in muscle satellite cells from normal and dystrophic patients. Lab Invest 81: 27–39, 2001.[ISI][Medline]

13. Fibbi G, D'Alessio S, Pucci M, Cerletti M, and Del Rosso M. Growth factor-dependent proliferation and invasion of muscle satellite cells require the cell-associated fibrinolytic system. Biol Chem 383: 127–136, 2002.[CrossRef][ISI][Medline]

14. Gyetko MR, Chen GH, McDonald RA, Goodman R, Huffnagle GB, Wilkinson CC, Fuller JA, and Toews GB. Urokinase is required for the pulmonary inflammatory response to Cryptococcus neoformans: a murine transgenic model. J Clin Invest 97: 1818–1826, 1996.[Abstract/Free Full Text]

15. Gyetko MR, Sud S, Kendall T, Fuller JA, Newstead MW, and Standiford TJ. Urokinase receptor-deficient mice have impaired neutrophil recruitment in response to pulmonary Pseudomonas aeruginosa infection. J Immunol 165: 1513–1519, 2000.[Abstract/Free Full Text]

16. Hawke TJ and Garry DJ. Myogenic satellite cells: physiology to molecular biology. J Appl Physiol 91: 534–551, 2001.[Abstract/Free Full Text]

17. Koh TJ and Escobedo J. Cytoskeletal disruption and small heat shock protein translocation immediately after lengthening contractions. Am J Physiol Cell Physiol 286: C713–C722, 2004.[Abstract/Free Full Text]

18. Lescaudron L, Peltékian E, Fontaine-Pérus J, Paulin D, Zampieri M, Garcia L, and Parrish E. Blood borne macrophages are essential for the triggering of muscle regeneration following muscle transplant. Neuromuscul Disord 9: 72–80, 1999.[CrossRef][ISI][Medline]

19. Lluís F, Roma J, Suelves M, Parra M, Aniorte G, Gallardo E, Illa I, Rodríguez L, Hughes SM, Carmeliet P, Roig M, and Muñoz-Cánoves P. Urokinase-dependent plasminogen activation is required for efficient skeletal muscle regeneration in vivo. Blood 97: 1703–1711, 2001.[Abstract/Free Full Text]

20. Lyons RM, Keski-Oja J, and Moses HL. Proteolytic activation of latent transforming growth factor-beta from fibroblast-conditioned medium. J Cell Biol 106: 1659–1665, 1988.[Abstract]

21. Mars WM, Zarnegar R, and Michalopoulos GK. Activation of hepatocyte growth factor by the plasminogen activators uPA and tPA. Am J Pathol 143: 949–958, 1993.[Abstract]

22. Merly F, Lescaudron L, Rouaud T, Crossin F, and Gardahaut MF. Macrophages enhance muscle satellite cell proliferation and delay their differentiation. Muscle Nerve 22: 724–732, 1999.[CrossRef][ISI][Medline]

23. Minamide LS and Bamburg JR. A filter paper dye-binding assay for quantitative determination of protein without interference from reducing agents or detergents. Anal Biochem 190: 66–70, 1990.[CrossRef][ISI][Medline]

24. Mitchell CA, McGeachie JK, and Grounds MD. Cellular differences in the regeneration of murine skeletal muscle: a quantitative histological study in SJL/J and BALB/c mice. Cell Tissue Res 269: 159–166, 1992.[CrossRef][ISI][Medline]

25. Muñoz-Cánoves P, Miralles F, Baiget M, and Félez J. Inhibition of urokinase-type plasminogen activator (uPA) abrogates myogenesis in vitro. Thromb Haemost 77: 526–534, 1997.[ISI][Medline]

26. Murphy G, Ward R, Gavrilovic J, and Atkinson S. Physiological mechanisms for metalloproteinase activation. Matrix Suppl 1: 224–230, 1992.[Medline]

27. Naldini L, Tamagnone L, Vigna E, Sachs M, Hartmann G, Birchmeier W, Daikuhara Y, Tsubouchi H, Blasi F, and Comoglio PM. Extracellular proteolytic cleavage by urokinase is required for activation of hepatocyte growth factor/scatter factor. EMBO J 11: 4825–4833, 1992.[Abstract]

28. Pizza FX, Koh TJ, McGregor SJ, and Brooks SV. Muscle inflammatory cells after passive stretches, isometric contractions, and lengthening contractions. J Appl Physiol 92: 1873–1878, 2002.[Abstract/Free Full Text]

29. Rifkin DB, Mazzieri R, Munger JS, Noguera I, and Sung J. Proteolytic control of growth factor availability. APMIS 107: 80–85, 1999.[ISI][Medline]

30. Robertson TA, Maley MA, Grounds MD, and Papadimitriou JM. The role of macrophages in skeletal muscle regeneration with particular reference to chemotaxis. Exp Cell Res 207: 321–331, 1993.[CrossRef][ISI][Medline]

31. Romer J, Bugge TH, Pyke C, Lund LR, Flick MJ, Degen JL, and Dano K. Impaired wound healing in mice with a disrupted plasminogen gene. Nat Med 2: 287–292, 1996.[CrossRef][ISI][Medline]

32. Schultz E and McCormick KM. Skeletal muscle satellite cells. Rev Physiol Biochem Pharmacol 123: 213–257, 1994.[ISI][Medline]

33. Shimizu M, Hara A, Okuno M, Matsuno H, Okada K, Ueshima S, Matsuo O, Niwa M, Akita K, Yamada Y, Yoshimi N, Uematsu T, Kojima S, Friedman SL, Moriwaki H, and Mori H. Mechanism of retarded liver regeneration in plasminogen activator-deficient mice: impaired activation of hepatocyte growth factor after Fas-mediated massive hepatic apoptosis. Hepatology 33: 569–576, 2001.[CrossRef][ISI][Medline]

34. Sisson TH, Hanson KE, Subbotina N, Patwardhan A, Hattori N, and Simon RH. Inducible lung-specific urokinase expression reduces fibrosis and mortality after lung injury in mice. Am J Physiol Lung Cell Mol Physiol 283: L1023–L1032, 2002.[Abstract/Free Full Text]

35. Sisson TH, Hattori N, Xu Y, and Simon RH. Treatment of bleomycin-induced pulmonary fibrosis by transfer of urokinase-type plasminogen activator genes. Hum Gene Ther 10: 2315–2323, 1999.[CrossRef][ISI][Medline]

36. Stefansson S and Lawrence DA. The serpin PAI-1 inhibits cell migration by blocking integrin {alpha}v{beta}3 binding to vitronectin. Nature 383: 441–443, 1996.[CrossRef][ISI][Medline]

37. Stepanova VV and Tkachuk VA. Urokinase as a multidomain protein and polyfunctional cell regulator. Biochemistry (Mosc) 67: 109–118, 2002.[CrossRef][ISI][Medline]

38. Suelves M, López-Alemany R, Lluís F, Aniorte G, Serrano E, Parra M, Carmeliet P, and Muñoz-Cánoves P. Plasmin activity is required for myogenesis in vitro and skeletal muscle regeneration in vivo. Blood 99: 2835–2844, 2002.[Abstract/Free Full Text]

39. Tatsumi R, Anderson JE, Nevoret CJ, Halevy O, and Allen RE. HGF/SF is present in normal adult skeletal muscle and is capable of activating satellite cells. Dev Biol 194: 114–128, 1998.[CrossRef][ISI][Medline]

40. Teixeira CFP, Zamunér SR, Zuliani JP, Fernandes CM, Cruz-Hofling MA, Fernandes I, Chaves F, and Gutiérrez JM. Neutrophils do not contribute to local tissue damage, but play a key role in skeletal muscle regeneration, in mice injected with Bothrops asper snake venom. Muscle Nerve 28: 449–459, 2003.[CrossRef][ISI][Medline]

41. Waltz DA, Natkin LR, Fujita RM, Wei Y, and Chapman HA. Plasmin and plasminogen activator inhibitor type 1 promote cellular motility by regulating the interaction between the urokinase receptor and vitronectin. J Clin Invest 100: 58–67, 1997.[Abstract/Free Full Text]

42. Yablonka-Reuveni Z, Seger R, and Rivera AJ. Fibroblast growth factor promotes recruitment of skeletal muscle satellite cells in young and old rats. J Histochem Cytochem 47: 23–42, 1999.[Abstract/Free Full Text]

43. Zhang G, Kim H, Cai X, López-Guisa JM, Alpers CE, Liu Y, Carmeliet P, and Eddy AA. Urokinase receptor deficiency accelerates renal fibrosis in obstructive nephropathy. J Am Soc Nephrol 14: 1254–1271, 2003.[Abstract/Free Full Text]

44. Zhao P, Iezzi S, Carver E, Dressman D, Gridley T, Sartorelli V, and Hoffman EP. Slug is a novel downstream target of MyoD: temporal profiling in muscle regeneration. J Biol Chem 277: 30091–30101, 2002.[Abstract/Free Full Text]





This Article
Abstract
Full Text (PDF)
All Versions of this Article:
289/1/C217    most recent
00555.2004v1
Alert me when this article is cited
Alert me if a correction is posted
Citation Map
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Download to citation manager
Google Scholar
Articles by Koh, T. J.
Articles by Sisson, T. H.
Articles citing this Article
PubMed
PubMed Citation
Articles by Koh, T. J.
Articles by Sisson, T. H.


HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2005 by the American Physiological Society.