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HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
From the Centre d'Oncologia Molecular, Institut de
Recerca Oncològica; the Unitat de Recerca de la Vall d'Hebron;
the Hospital de la Santa Creu i Sant Pau, Barcelona, Spain; the Center
for Transgene Technology, Flanders Interuniversity, Leuven, Belgium;
and the Randall Centre, New Hunt's House, Kings College, London,
United Kingdom.
Plasminogen activators urokinase-type plasminogen activator (uPA)
and tissue-type plasminogen activator (tPA) are extracellular proteases
involved in various tissue remodeling processes. A requirement for uPA
activity in skeletal myogenesis was recently demonstrated in vitro. The
role of plasminogen activators in skeletal muscle regeneration in vivo
in wild-type, uPA-deficient, and tPA-deficient mice is investigated
here. Wild-type and tPA Activation of the zymogen plasminogen (Plg) into
the active serine proteinase, plasmin, is a highly regulated and widely
used mechanism for the generation of extracellular proteolytic
activity. Activation of Plg is exerted by 2 distinct Plg activators,
tissue-type plasminogen activator (tPA) and urokinase-type plasminogen
activator (uPA). The plasminogen system is a key factor in the
dissolution of fibrin matrices and is critical in the maintenance of
hemostatic balance. By its action in concert with other proteolytic
systems, it is also thought to play a role in the degradation of
extracellular matrices in physiologic and pathologic tissue
remodeling and cell migration events, such as ovulation, trophoblast
invasion, post-lactational mammary involution, wound healing,
angiogenesis, and tumor cell invasion (reviewed in1).
Extracellular proteolysis takes place during skeletal muscle formation
and pathologic muscle regeneration, in which muscle precursor satellite
cells play a major role. In response to muscle injury, damaged tissue
is infiltrated by fibroblasts, inflammatory cells, and
macrophages.2 Necrotic tissue is removed,
revascularization starts, and proliferation of satellite cells is
initiated. Numerous proteolytic enzymes have been proposed to play a
role during muscle regeneration, either in the inflammatory response or
in the migration of myoblasts across the basal lamina and in their
further fusion to form the terminal muscle fiber.3,4
Metalloproteinases (MMPs) such as MMP-2 and MMP-9, meltrin- Animals
Induction of muscle regeneration
Histologic and immunohistochemical analysis At selected times, muscles of control and knockout mice were removed after cervical dislocation. They were carefully dissected, frozen in isopentane-chilled liquid nitrogen, and stored at 80°C before sectioning. Transverse cryostat sections (10 µm thick) were
stained with hematoxylin-eosin (H&E). Immunohistochemistry was
performed using the Vectastain Elite kit (Vector Laboratories, Burlingame, CA) and diaminobenzidine for single-labeling
experiments. The following antibodies were used: mouse monoclonal
antibody against myosin developmental-type heavy chain (MHCd)
(Novocastra, Newcastle, United Kingdom) 1:50, rat monoclonal anti-CD31
antibody (Pharmingen, San Diego, CA) 1:25, goat polyclonal M-cadherin
(sc-6470; Santa Cruz Biotechnology, Santa Cruz, CA) 1:25, and rabbit
antimouse fibrinogen (kindly provided by Dr Keld Dano, Finsenlab,
Denmark) 1:1000. Sections stained with antimouse fibrinogen were
co-stained with hematoxylin to visualize cellular structures. For
single-immunofluorescence staining, sections were incubated with rat
monoclonal antibodies anti-Mac-1 (1:30) or anti-Gr-1 (1:30;
Pharmingen) conjugated with fluorescein and mounted with Vectashield
(Vector Laboratories). For double-immunofluorescence staining, sections
were incubated overnight simultaneously with Mac-1/uPA, Gr-1/uPA, and
M-cadherin-uPA primary antibodies, washed, and overlaid with the
appropriate secondary antibodies conjugated with rhodamine (for
M-cadherin), Texas Red (for Mac-1 and Gr-1), or fluorescein (for uPA);
slides were then washed and mounted with Vectashield (Vector
Laboratories). Sections were photographed using an Olympus BX60
microscope (Tokyo, Japan). Control experiments without primary
antibody demonstrated that the immunofluorescence signals observed were
specific (data not shown).
RNA analysis Total RNA was extracted and purified from freshly isolated muscle tissue according to the procedure of Chomczynski and Sacchi.18 Three micrograms total RNA was subjected to Northern blot analysis, and blots were sequentially hybridized with radiolabeled complementary DNA (cDNA) probes corresponding to murine uPA, tPA, MyoD, and myogenin as previously described.13,19 To normalize signal intensity, blots were later rehybridized with a radiolabeled 18S oligonucleotide probe. For reverse transcription-polymerase chain reaction (RT-PCR) analysis, 2 µg total RNA was reverse transcribed using the first-strand cDNA synthesis kit (Pharmacia, Uppsala, Sweden) in a 35-µL reaction. Amplification parameters were denaturation at 95°C for 45 seconds; annealing for 2 minutes at 55°C (uPA), 60°C (MyoD), 60°C (myogenin), 55°C (glyceraldehyde phosphate dehydrogenase [GAPDH]); and extension at 72°C for 2 minutes. Aliquots of 10% of the reaction were removed from the assay after different cycles, as indicated, and analyzed on 2% agarose gels. Primers for the detection of reverse transcriptase products were derived from different exons to distinguish RT-PCR products from genomic DNA contaminations. Primer sequences were: uPA, 5'-GGCAGTGTACTTGGAGCTCCT-3' and 5'-TAGAGCCTTCTGGCCACACTG-3'; MyoD, 5'-AGGCTCTGCTGCGCGACC-3' and 5'-TGCAGTCGATCTCTCAAAGCACC-3'; myogenin, 5'-GAGCGCGATCTCCGCTACAGAGG-3' and 5'-CTGGCTTGTGGCAGGCCCAGG-3'; GAPDH, 5'-ACTCCCACTCTTCCACCTTC-3' and 5'-TCTTGCTCAGTGTCCTTGC-3'. Expected product sizes were: uPA, 450 base pairs (bp); MyoD, 471 bp; myogenin, 379 bp; GAPDH, 185 bp.Preparation of muscle extracts Gastrocnemius muscles were dissected, sectioned, weighed, and pounded in an ice-cold Potter tube with 0.1 mM Tris-HCl buffer, pH 7.6, containing 2 mM EDTA and 0.4% Triton X-100. The resultant extracts were centrifuged for 20 minutes at 4°C at 12 000g, and the supernatants were stored as aliquots at80°C until use. Protein concentration was determined in the supernatants using the Bio-Rad protein assay (Bio-Rad, Hercules, CA).
Plasmin kinetics Plasmin content in muscles was measured using the S-2251-based assay, which was performed in triplicate in microtiter plates at 37°C and calibrated with purified plasmin. The chromogenic substrate selective for plasmin, S-2251 (Chromogenix, Milan, Italy), was used to follow the initial rate of plasminogen activation by measuring p-nitroaniline generation. Forty micrograms muscle extract from wild-type mice (obtained on different days after injury; see Figure 7) or standard samples was mixed with a buffer containing 0.1 M Tris-HCl and 2 mM EDTA, pH 7.6, and 1.6 mM S-2251 as substrate. The generation of plasmin was detected by measuring p-nitroaniline release from the substrate, as indicated above.Zymography Three hundred micrograms muscle extracts was size-fractionated on a 10% nonreducing sodium dodecyl sulfate (SDS) acrylamide gel, which was washed for 30 minutes in 2.5% Triton X-100-phosphate-buffered saline and for 30 minutes in distilled water. The gel was subsequently placed in contact with a casein gel containing 2% (wt/vol) nonfat dry milk, 0.25 mM Tris-HCl, pH 7.6, 1% (wt/vol) agarose, 0.25 × phosphate-buffered saline, and 15 µg/mL plasminogen (Chromogenix), and incubated in a humid chamber at 37°C until caseinolytic bands were visualized and photographed.Systemic defibrinogenation Then uPA-deficient mice were anesthetized, and 14-day mini-osmotic pumps (model 1002; Alza, Palo Alto, CA) filled with a buffered solution of 500 U/mL ancrod (Sigma Chemical, St Louis, MO) were implanted subcutaneously into their backs (one mini-pump per animal). The insertion sites were sutured closed. The pumps deliver 0.25 µL/h, so the mice received 3 U ancrod/d. In control animals, saline-filled mini-pumps were implanted. On day 3 of ancrod or saline infusion, injury was induced by intramuscular injection of 50% glycerol. Nine days after injury, mice were killed and gastrocnemius muscles were dissected, frozen, and analyzed by H&E staining. Fibrinogen levels in citrated blood from ancrod- or saline-treated mice were analyzed by SDS-polyacrylamide gel electrophoresis (PAGE; 6% polyacrylamide gel) followed by Western blotting using an antimouse fibrinogen antibody (1:1000).
Expression of uPA messenger RNA and proteolytic activity are induced after skeletal muscle injury We have previously demonstrated a role for uPA in myogenesis in vitro because the inhibition of uPA activity abrogated both differentiation and fusion of C2C12 myoblasts in culture.13 Our aim in the current study was to decipher the role of uPA during skeletal muscle regeneration in vivo. Muscle regeneration was induced in mice either by freeze-crush16 or by intramuscular injection of 50% glycerol,17 and the expression of uPA was investigated in regenerating muscle after injury and compared with that of the contralateral control muscle.RNA was isolated from muscles on the third day after glycerol-induced
injury, and uPA, MyoD, and myogenin expression was analyzed by
semiquantitative RT-PCR. As shown in Figure
1A, significantly fewer amplification
cycles were needed with RNA from regenerating muscle to yield
comparable signals for uPA than with RNA from control muscle, whereas
no differences were observed when primers specific for GAPDH were used
(Figure 1A). As a control for muscle regeneration, the expression of
myogenic regulators MyoD and myogenin (up-regulated by satellite cells
during muscle regeneration in vivo)20,21 was also
examined. Detectable PCR signal for MyoD and myogenin was obtained
after fewer amplification cycles with RNA from glycerol-treated muscle
than with RNA from control muscle. Similarly, when muscle regeneration
was induced by traumatic freeze-crush, the expression of uPA, MyoD, and
myogenin mRNA was dramatically induced in regenerating muscle of
wild-type mice as assessed by Northern blot analysis (Figure 1B top;
see WT panel, control, and wound). We observed that, as with uPA, tPA
expression is induced in regenerating skeletal muscle, though to a
lesser extent, suggesting that uPA rather than tPA might may be
involved in skeletal muscle regeneration in vivo (Figure 1B bottom; see
WT panel, control, and wound). Thus, injured muscle up-regulates uPA
and tPA mRNA.
Next, we analyzed whether the induction of uPA and tPA messenger RNAs (mRNAs) caused by muscle injury was followed by an induction of their corresponding enzymatic activities. Tissue extracts were prepared from noninjured and injured muscles on days 2, 5, 7, and 9 after injury and were analyzed by zymography (Figure 1C top). This assay allows clear distinction between uPA and tPA, which migrate at 45 and 72 kd, respectively. In tissue extracts from noninjured muscle, uPA activity was not detectable. However, it was induced in the regenerating muscle samples, as demonstrated by the appearance of a casein degradation band of 45 kd, corresponding to murine uPA active enzyme. Under the same conditions, tPA activity was undetectable in all the tissue extracts analyzed. As expected, no uPA activity was detected in tissue extracts from muscle of uPA-deficient mice, demonstrating the specificity of the assay (Figure 1C bottom; see lanes 1-2). These results suggested that uPA rather than tPA might be involved in skeletal muscle regeneration in vivo. Expression of MyoD and myogenin mRNA is reduced in regenerating muscle tissue of uPA-deficient mice To evaluate the importance of uPA and tPA for the skeletal muscle regeneration process, we performed freeze-crush injury-induced regeneration experiments in uPA- and tPA-deficient mice.14 RNA levels for MyoD and myogenin from wild-type and uPA- or tPA-deficient mice were examined 3 days after injury by Northern blotting (Figure 1B). MyoD expression was below detectable levels in quiescent muscle from wild-type mice and was dramatically induced in injured muscle (Figure 1B; compare control and wound in WT). MyoD transcript levels were induced to a lesser extent in uPA-deficient mice than in wild-type or tPA-deficient mice after injury (Figure 1B; compare wound in WT, uPA/, and tPA/). Similarly, myogenin mRNA
expression was reduced in the regenerating muscle of uPA-deficient mice
compared to that of tPA-deficient and wild-type mice, whereas the 18S
RNA level was comparable in all lanes. As expected, uPA and tPA
transcripts were absent in muscle from uPA- and tPA-deficient mice,
respectively (Figure 1B). These results indicate that uPA, but not tPA,
is required for efficient skeletal muscle regeneration in vivo.
uPA deficiency exacerbates histologic features of skeletal muscle degeneration To determine the functional significance of the increased uPA expression in skeletal muscle after damage and to evaluate further the importance of uPA for skeletal muscle regeneration, we analyzed comparatively the histopathologic changes induced by intramuscular injection of glycerol in the muscle fibers of wild-type and uPA- and tPA-deficient mice. As shown in Figure 2A, skeletal muscle of uPA-deficient mice displayed a prominent regeneration defect after glycerol-induced injury, whereas regeneration in tPA-deficient and wild-type mice proceeded normally. This regeneration defect in uPA/ mice was
apparent by 5 days after injury, but it was most striking 9 to 20 days
after injury. Analysis of H&E-stained cross-sections of wild-type,
uPA-deficient, and tPA-deficient mice 2 days after injury showed that
the muscles of all mice were edematous and had fibrotic infiltrates
within the enlarged intercellular space separating the necrotic
myofibers. Analysis of cross-sections of muscle 5 days after injury
revealed well-advanced regeneration in wild-type and tPA-deficient
mice, with many new myofibers characterized by its small size and
single nuclei and with a reduction in fibrotic infiltrates (Figure 2A).
In contrast, in uPA-deficient mice at 5 days after injury, the muscle
appeared edematous, and no new uninucleated, small myofibers were
detected yet. Seven days after injury, most injured fibers regenerated
into groups of centrally nucleated myotubes, a clear sign of advanced
regeneration, and few necrotic fibers could occasionally be observed in
wild-type and tPA-deficient mice; in addition, the size of newly formed myofibers had augmented in the wild-type and tPA/ animals. In uPA-deficient mice, however, the muscle had a necrotic appearance, showing still extensive fibrosis 7 days after injury. Nine days after
injury, virtually no sign of previous damage was detectable in
wild-type and tPA/ mice, and centrally located nuclei were observed
inside the regenerated fibers, which also exhibited increases in the
cross-sectional areas, indicating complete regeneration (Figure 2A). In
uPA-deficient mice, however, a high number of degenerated myotubes was
still visible. Twenty days after injury, the lesion was no longer
noticeable in wild-type and tPA/ mice, except for the central
myonuclei. In uPA/ mice, however, the muscle showed extensive
fibrosis with high numbers of degenerated myotubes. Furthermore,
staining of wild-type and uPA-deficient muscle tissue at 5 and 9 days
after injury with a monoclonal antibody against developmental myosin
heavy chain (MHCd, an MHC isoform expressed only in embryonic and early
neonatal life, whose expression is re-induced on muscle regeneration in
adult life) showed regenerating myotubes expressing MHCd at 5 days after injury in both wild-type and uPA/ mice (Figure 2B). In
contrast, at 9 days after injury, MHCd expression was detected in
uPA/ mice but absent in wild-type mice. These results reveal the
persistence of ongoing muscle degeneration-regeneration in
uPA-deficient mice at stages at which muscle regeneration is more
advanced in wild-type mice.
Impact of uPA deficiency on the inflammatory response after muscle injury Blood-borne monocytes-macrophages are recruited after injury to skeletal muscle during the inflammatory phase. These cells play a major role in the phagocytosis of tissue debris after muscle injury.2 To analyze the distribution of macrophages in normal and regenerating muscle of wild-type and uPA-deficient mice, we performed immunohistochemical studies using an antibody against a well-characterized macrophage marker, Mac-1. A few resident Mac-1-positive cells were detected in the noninjured muscle sections of wild-type and uPA/ mice (Figure
3A; 0 days after injury). There was an
increase of Mac-1-positive cells at the injury site 2 days after
injury; however, the extent of Mac-1 staining in uPA-deficient mice was
reduced to almost 50% compared with that in wild-type mice (Figure 3A
[2 days after injury] and 3D), suggesting that macrophage recruitment
to the injured muscle was reduced in the absence of uPA. Although
macrophages have been identified as the predominant cell type in the
inflammatory infiltrate in experimentally induced muscle injury,
neutrophils have also been detected during the very early hours of the
muscle injury response.2 To investigate the presence of
neutrophil infiltrates in wild-type and uPA/ mice, we performed
immunohistochemical studies using an antibody against a neutrophil
marker, Gr-1. No Gr-1-positive cells were detected in noninjured
muscle sections of wild-type or uPA-deficient mice (Figure 3B; 0 days
after injury). Twelve hours after injury, there was an increase in
Gr-1-expressing cells at the injury site of both types of animals,
though the neutrophil response was more marked in wild-type than in
uPA/ mice at the site of injury (Figure 3B; 0.5 days after injury).
It is known that uPA has important functions in neovascularization and
capillary angiogenesis, such as after cardiac infarct.22
To assess whether angiogenesis in regenerating muscle of uPA/ mice
was impaired, immunohistochemistry with a vascular endothelial cell
marker, CD31/PECAM-1, was carried out.23 We found no
significant differences in the number of CD31-stained microvessels
between uPA/ and wild-type mice (Figure 3C). However, we cannot
completely rule out that small differences in the amount or diameter of
microvessels exist between uPA/ and wild-type mice. Taken together,
these data suggest that the reduced muscle regeneration of
uPA-deficient mice may be caused by a decreased inflammatory response
shortly after injury.
Macrophage and satellite cells contribute to the expression of uPA during skeletal muscle regeneration We have demonstrated that both uPA mRNA and proteolytic activity are induced after muscle injury in wild-type mice (Figure 1). To determine precisely where uPA is expressed during the regeneration process, we performed double-immunohistochemical staining on sections of regenerating muscle using an antibody against uPA together with antibodies against Mac-1, Gr-1, and M-cadherin, specific markers for macrophages, neutrophils, and satellite cells, respectively. As shown in Figure 4, uPA was detected in Mac-1-positive cells, but not in Gr-1-expressing cells, suggesting that uPA is expressed by macrophages but not by neutrophils during the inflammatory response to skeletal muscle injury. In addition, M-cadherin was found to be co-expressed with uPA, demonstrating that uPA was synthesized by skeletal muscle stem cells in vivo (Figure 4).
Increased fibrin deposition in damaged muscle of uPA-deficient mice Given that the established role of plasminogen activation is fibrinolysis (reviewed in24), we analyzed whether the loss of uPA resulted in increased fibrin deposition after muscle injury. Fibrin content in regenerating muscle from wild-type and uPA-deficient mice was analyzed by fibrin immunohistochemistry, using an antimouse fibrin(ogen) antibody (Figure 5). In 9-day-injured muscle of wild-type mice, minimal fibrin immunoreactivity was found (Figure 5; WT). In contrast, abundant deposits of fibrin were detected in muscles of uPA-deficient mice at the same time after injury (Figure 5; uPA/).
These results correlate with the persistence of muscle degeneration
features observed in uPA/ mice 9 days after injury (Figure 2;
uPA/, 9 days after injury).
Systemic fibrinogen depletion restores muscle regeneration in uPA-deficient mice Administration of ancrod (a viper venom) has been shown to lead to the consumption of systemic fibrinogen.25 Ancrod- or saline-delivering osmotic pumps (3 U/d) were implanted in uPA/ mice for 3 days before muscle injury and then throughout the
experimental period (up to 9 days after injury). As shown in Figure
6A, ancrod administration resulted in a
significant reduction of plasma fibrinogen, as assessed by Western
blotting, without any effect on survival (compare ancrod versus
saline). Moreover, signs of improved muscle regeneration were
detectable in ancrod-treated uPA-deficient mice 9 days after injury,
with centrally located nuclei inside the regenerated fibers. In
contrast, histologic features of muscle degeneration persisted in
saline-treated uPA-deficient mice at the same stage after injury,
including a high number of degenerated myotubes and fibrosis in the
intercellular space throughout the damaged muscle (Figure 6B). The
extent of muscle regeneration of ancrod-treated uPA-deficient mice 9 days after injury was comparable to the regeneration status of
wild-type mice at the same time after injury. This observation supports
the idea of a potential pathogenic role of fibrin(ogen) accumulation
during muscle regeneration in uPA-deficient mice.
Plasminogen-deficient mice show a severe regeneration defect with enhanced fibrosis and myotube degeneration The results shown above provide evidence of a role for uPA in skeletal muscle regeneration in vivo. To investigate whether the effect of uPA is plasminogen dependent (ie, whether the lack of uPA prevents muscle regeneration because of a failure of plasminogen processing), we measured plasmin activity in regenerating muscle tissues at different times after injury. Using a chromogenic substrate for the activity of plasmin, we observed that plasmin generation was increased during skeletal muscle regeneration in wild-type mice, with a peak of activity occurring 2 to 5 days after injury and decreasing by day 9 (Figure 7). To test the role of plasmin in skeletal muscle regeneration, we performed glycerol-induced muscle injury in wild-type mice and in mice deficient in plasminogen (Plg/) and comparatively analyzed skeletal muscle regeneration. By
day 9 after injury, virtually no sign of previous damage was detectable
in wild-type mice except for the presence of centrally located nuclei
inside the regenerating fiber, indicating complete regeneration. In
Plg-deficient mice, however, muscle presented histologic features of
degeneration (Figure 8A). The presence of
fibrin in cross-sections of regenerating muscle of wild-type and
Plg/ mice was also analyzed in both types of mice. Although no
fibrin immunoreactivity was detected in regenerating muscle of
wild-type mice, extensive fibrin deposition was observed in regenerating muscle of Plg-deficient mice 9 days after injury (Figure
8B). These results indicated that, like uPA-deficient mice,
Plg-deficient mice developed significant fibrin deposition on muscle
injury, leading to a defective muscle regeneration process.
In the current study, we describe the consequences of the inactivation of plasminogen activation system genes during skeletal muscle regeneration. Mice lacking uPA, but not tPA, show a pronounced regeneration defect after experimentally induced injury of muscle, suggesting a protective role for uPA in the muscle regeneration process. The persistence of muscle degeneration in uPA-deficient mice was reproduced in Plg-deficient mice. This indicates that between the 2 pathways of Plg activation (uPA- or tPA-mediated), uPA-mediated Plg activation is the major one in the muscle regeneration process. This conclusion is supported by zymographic analysis of regenerating muscles of wild-type mice, which only showed uPA activity. Interestingly, in C2C12 murine myoblasts, uPA activity was also predominant.13 Finally, the similar phenotypes of uPA- and Plg-deficient mice indicates, in this animal model, the predominance of uPA-dependent plasmin effects. We have demonstrated a significant accumulation of extravascular fibrin
in regenerating muscle of uPA Our finding that loss of Plg activation impedes muscle regeneration
raises the question of how the uPA-Plg system is involved in tissue
repair. Inflammation is a process frequently associated with tissue
repair because degenerating tissues are invaded by inflammatory cells.
Our study showed that in response to glycerol-induced muscle injury,
macrophages accumulated near the injury site 48 hours after injury.
Similar results in macrophage recruitment were reported in a study of
regeneration after bupivacaine-induced muscle injury in the
rat28 and after crush injury in the mouse.2 We have observed that mice with a specific deficit in uPA show a
reduced staining for Mac-1-positive cells 48 hours after injury, indicating that the number of macrophages reaching the injury site is
reduced in the absence of uPA. This suggests that uPA activity may have
a profound effect on inflammation and inflammation-related muscle
disease. Recent work29 with uPA-deficient mice has
demonstrated that uPA is required for the pulmonary inflammatory
response to Cryptococcus neoformans because a lack of uPA
resulted in inadequate macrophage recruitment, uncontrolled infection,
and death. In addition, endogenously produced uPA could amplify tumor
necrosis factor- The hypothesis that, in general, Plg activation facilitates cellular
penetration of fibrin-containing matrices is in accordance with a
putative scenario occurring in normal skeletal muscle regeneration, with local conversion of Plg to plasmin and subsequent fibrin degradation. Plasmin is probably one member of a team of carefully regulated and specialized matrix-degrading enzymes, including serine,
metallo, and other classes of proteases, that together serve in matrix
remodeling and cellular reorganization of wound fields. Our data
suggest that plasmin is particularly useful in fibrin solubilization;
without it, cellular reorganization of fibrin-rich matrices is severely
impeded. We conclude that the uPA-Plg system is required for the normal
regeneration and resolution of muscle damage in mice. The reduced
presence of macrophages at the injury site suggests that a key
factor slowing muscle repair in uPA-deficient animals may be the
diminished ability of responding macrophages to proteolytically dissect
their way through the extracellular matrix until they reach the injury
site. In this model, one major matrix component within damaged areas
that may represent a particular impediment to inflammatory cell
migration in the absence of uPA (or Plg) is fibrin. However, based on
the existing data, we cannot exclude that the uPA-Plg deficiency may
impede cell migration because of the lost contribution of plasmin to
degrade other matrix components or the lack of other key matrix
proteinases or of growth factors. Apart from its effects on
extracellular matrix proteins, uPA-plasmin can cleave and activate
latent forms of growth-angiogenic factors such as transforming growth
factor- In conclusion, our results demonstrate that uPA fulfills a beneficial role in skeletal muscle regeneration, mainly through uPA-mediated fibrinolytic activity. Compounds aimed at decreasing fibrin levels in muscle may be clinically useful in therapy for muscular dystrophy. Future experiments will be directed to further definition of the benefit of defibrinogenation, anticoagulant, or fibrinolytic agents in dystrophinopathy. The prevention of plasminogen activation has revealed the requirement of uPA-plasmin and the dispensability of tPA in the regeneration of skeletal muscle. This report constitutes the first demonstration for a role of a proteolytic system in skeletal muscle regeneration in vivo. Rescue experiments by injection of recombinant proteases or retroviruses will show whether exogenously supplied uPA can compensate for the mutation. If this is the case, uPA might prove to be useful for the treatment of muscle injuries or muscle dystrophies, particularly if uPA evokes the response preferentially in muscle.
We thank Dr Keld Dano for the antifibrinogen and Dr Lourdes Gombau, A. Martín, and D. Fernández for assistance.
Submitted August 22, 2000; accepted November 9, 2000.
Supported by DGES grant PM97-008 and by a grant from Fundació La Marató-TV3 (P.M.C., M.R.), by FISS94-0968, by the Medical Research Council and the British Council (S.M.H.), and by the European Union (P.M.C., S.M.H., P.C.).
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Pura Muñoz-Cánoves, Centre d'Oncologia Molecular, Institut de Recerca Oncològica, Aut Castelldefels, km 2.7, 08907 L'Hospitalet de LL, Barcelona, Spain; e-mail: pmunoz{at}iro.es.
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Top
Abstract
Introduction
, and
cathepsin B seem to be required for myotube formation in
vitro.
a single freeze-crush injury of the musculus tibialis anterior (TA), as described by McGeachie and
Grounds,
and
uPA-deficient 
mice. Muscles were analyzed for Mac-1
immunofluorescence at 0 and 2 days after injury. Mac-1-positive cells
were counted in 10 random fields per section, and data from 12 sections
per muscle were pooled. Mean values for both limbs were to provide a
mean value for each animal (3 animals were studied at each time point).
Histograms show the percentage of reduction (46%) of Mac-1-positive
cells in uPA
and hepatocyte growth factor-scatter
factor.