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HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
From the Centre d'Oncologia Molecular, Institut de
Recerca Oncològica, Barcelona, Spain; and the Center for
Transgene Technology, Flanders Interuniversity, Leuven, Belgium.
Plasmin, the primary fibrinolytic enzyme, has a broad substrate
spectrum and is implicated in biologic processes dependent upon
proteolytic activity, such as tissue remodeling and cell migration.
Active plasmin is generated from proteolytic cleavage of the zymogen
plasminogen (Plg) by urokinase-type plasminogen activator (uPA) and
tissue-type plasminogen activator (tPA). Here, we have investigated the
role of plasmin in C2C12 myoblast fusion and differentiation in vitro,
as well as in skeletal muscle regeneration in vivo, in wild-type and
Plg-deficient mice. Wild-type mice completely repaired experimentally
damaged skeletal muscle. In contrast, Plg The proteolytic conversion of the ubiquitous
zymogen plasminogen (Plg) to the active protease plasmin is an
extensively used mechanism for the generation of extracellular
proteolytic activity. This conversion is exerted by 2 physiologic
plasminogen activators (PAs), tissue-type plasminogen activator (tPA)
and urokinase-type plasminogen activator (uPA). Because unrestrained
generation of the broad-spectrum protease plasmin may be hazardous to
the cells, plasmin activity is tightly controlled at the levels of PAs
by plasminogen activator inhibitors (PAI-1 and PAI-2) and at the level
of plasmin by Extracellular proteolysis takes place during skeletal muscle formation
and pathological muscle regeneration, in which satellite cells play a
major role. In response to muscle injury, damaged tissue is infiltrated
by fibroblasts, inflammatory cells, and macrophages.3
Necrotic tissue is removed, revascularization starts, and proliferation
of satellite cells is initiated. A number of proteolytic enzymes have
been proposed to play a role during muscle regeneration, either in the
inflammatory response and/or in the migration of myoblasts across the
basal lamina and in their further fusion to form the terminal muscle
fiber.4,5 Metalloproteinases such as MMP-2 and MMP-9,
meltrin- Cell culture
Animals
Induction of muscle regeneration Regeneration of skeletal muscle was induced by intramuscular injection of 50 µL of 50% glycerol (vol/vol) in the gastrocnemius muscle group of the mice, as described by Kawai et al.17 The experiments were performed in right hind-limb muscles, and contralateral intact muscles were used as control. Morphologic and biochemical examinations were performed at 2, 4, 7, 10, 30, and 50 days after injury (AI). Four animals were used for each time point.Histologic and immunohistochemical analysis At selected times, muscles of control and Plg /
mice were removed after cervical dislocation. They were carefully
dissected, frozen in isopentane-chilled liquid nitrogen, and stored at
 80°C prior to 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) 1:50 (Novocastra, Newcastle, United Kingdom) and rabbit
antimouse fibrinogen 1:1000 (kindly provided by Dr K. Dano, Finsenlab,
Denmark). For immunofluorescence staining, sections were incubated with
rat monoclonal antibodies anti-Mac-1 1:30 (Pharmingen, San Diego, CA),
anti-Gr-1 1:30 (Pharmingen), and anti-T11 1:50 (Coulter Immunology,
Hialeah, FL) conjugated with fluorescein and mounted with Vectashield
(Vector Laboratories). Sections were photographed using an Olympus BX60
microscope (Tokyo, Japan). Control experiments without primary antibody
demonstrated that immunofluorescence signals observed were specific
(not shown).
RNA analysis Total RNA was extracted and purified from either C2C12 cells or freshly isolated muscle tissue according to the procedure of Chomczynski and Sacci.18 Five micrograms of total RNA was subjected to Northern analysis, and blots were hybridized with radiolabeled complementary DNA probe corresponding to murine myogenin, as previously described.14,19 To normalize signal intensity, blots were later rehybridized with a radiolabeled 18S oligonucleotide probe. For reverse transcription (RT)-PCR analysis, 2 µg total RNA was reverse transcribed using the first-strand complementary DNA 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 58°C, and extension at 72°C for 1 minute. Primers for detection of Plg reverse transcriptase product were derived from murine Plg complementary DNA sequence: forward primer: 5'-TCAGCAGGGAATGTCACGG-3'; reverse primer: 5'-CTCTCTGTCGCCTTCCATGG-3'. Expected RT-PCR product size was 410 bp.Preparation of muscle extracts Gastrocnemius muscles were dissected out and sectioned, weighed, and pounded in an ice-cold Potter tube with 0.1 mM Tris-HCl buffer, pH 7.6, containing 2 mM ethylenediaminetetraacetic acid (EDTA) and 0.4% Triton X-100. The resulting 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 (Herts, United Kingdom) protein assay.
Analysis of myogenin protein expression Tissue extracts from noninjured or injured muscle were prepared as described above, and myogenin expression was analyzed by Western blotting using a rabbit polyclonal antimyogenin antibody 1:200 (sc-576; Santa Cruz Biotechnology, Santa Cruz, CA). Immunoblots were performed and developed with the ECL detection system (Amersham, Buckinghamshire, United Kingdom).Plasmin kinetics Plasmin content in 20 µL conditioned media (5-fold concentrated, using Centricon, Pall Filtron, Ann Arbor, MI) from C2C12 cells in DMEM or DMEM supplemented with insulin was measured using the S-2251-based assay, which was done in triplicate in microtitration plates at 37°C and calibrated with purified plasmin (Chromogenix, Milan, Italy). The chromogenic substrate selective for plasmin, S-2251 (Chromogenix), was used to follow the initial rate of Plg activation by measuring p-nitroaniline generation. Conditioned media were mixed with a buffer containing 0.1 M Tris-HCl and 2 mM EDTA, pH7.6, and 1.6 mM S-2251 as substrate. The generation of plasmin was detected by measuring the p-nitroaniline release from the substrate as indicated above.Zymography Sixty micrograms of muscle extracts or 40 µL C2C12 conditioned media were 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 Plg (Chromogenix) and incubated in a humid chamber at 37°C until caseinolytic bands were visualized and photographed.Systemic defibrinogenation Plg-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), were implanted subcutaneously into their backs (one minipump per animal). The insertion sites were then closed by sutures. The pumps deliver 0.25 µL/h, so the mice received 3 U ancrod/d. In control animals, saline-filled minipumps were implanted. On day 3 of ancrod or saline infusion, injury was induced by intramuscular injection of 50% glycerol. Ten 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.Statistical analysis The Student t test was used to determine whether there were significant (P < .05) differences between WT and Plg-deficient mice.
C2C12 muscle cells produce uPA-dependent plasmin activity Previous studies have reported that myogenic cells express Plg activators in vitro.11,12,14 However, the presence of the downstream protease plasmin in muscle cells has never been documented. Thus, we investigated whether plasmin activity was generated by C2C12 cells (a myoblast cell line derived from murine satellite cells), which have been extensively used as a model for myogenesis in vitro.20 First, we demonstrated by RT-PCR that Plg messenger RNA was synthesized in C2C12 cells (Figure 1A). Using a specific chromogenic substrate for the activity of plasmin, we observed that plasmin activity was detected both in conditioned medium of proliferating preconfluent C2C12 myoblasts (lane 1) (Figure 1B) and in conditioned medium of fusing myotubes (lane 2) (Figure 1B). To analyze whether the generation of plasmin by C2C12 cells was uPA- or tPA-dependent, we analyzed the activity of both PAs in these cells by zymography. This assay allowed clear distinction between uPA and tPA, which migrated at 45 and 72 kd, respectively. As shown in Figure 1C, uPA proteolytic activity was clearly observed in C2C12-conditioned medium of both myoblasts (lane 1) and myotubes (lane 2), whereas tPA activity was practically undetectable. These results suggested that the generation of plasmin in C2C12 muscle cells was mediated mainly by uPA.
Inhibition of plasmin activity reduces myotube formation and muscle differentiation in vitro To investigate the role of plasmin in C2C12 myogenesis, we analyzed whether inhibition of plasmin activity affected myoblast fusion and differentiation in vitro. C2C12 cells cultured in DMEM plus 10% FBS were transferred to DMEM containing insulin for 3 days, to induce myoblast fusion and differentiation, in the absence or presence of the physiologic plasmin inhibitor, 2-antiplasmin, and
myotube formation and differentiation were analyzed consequently. As
shown in Figure 2A,B, in the absence of
2-antiplasmin, C2C12 cells fused extensively; in contrast, fewer
myotubes formed in cultures that had been treated with
2-antiplasmin; as an experimental control, we observed that SB203580
(a specific inhibitor of p38 mitogen-activated protein kinase)
significantly reduced myoblast fusion, as previously
reported,21,22 while no decrease in myotube formation was
observed in the presence of BSA. The effect of 2-antiplasmin on
myotube formation was determined microscopically by counting the
fraction of nuclei present in myotubes. The results represented in the
graph (Figure 2A) show that 2-antiplasmin reduced myotube formation
by 45%. SB203580 reduced myotube formation by 80%, while treatment of
cells with BSA had practically no effect. To establish whether the
inhibition of plasmin activity also affected myogenic differentiation,
RNA was obtained from cells treated or not with 2-antiplasmin,
SB203580, or BSA and analyzed for the expression of the myogenic
differentiation marker myogenin. As shown in Figure 2C, myogenin
transcripts were absent in preconfluent myoblasts grown in DMEM plus
10% FBS (lane 1), while their accumulation was induced when C2C12
cells were switched to DMEM plus insulin for 3 days, coinciding with
extensive myotube formation (lane 2). When differentiating C2C12 cells
were incubated with 2-antiplasmin (lane 3) or SB203580 (lane 4) for
3 days, the levels of myogenin messenger RNA were significantly
decreased; in contrast, no alteration in the levels of these
transcripts was observed when BSA was added to the cells (lane 5).
Furthermore, expression of myogenin was already detected 18 hours
following incubation of C2C12 cells in differentiation medium, at a
stage when no myotubes had been formed (Figure 2D, lane 1), while in
the presence of 2-antiplasmin or SB203580 the levels of myogenin
were reduced (lanes 2 and 3, respectively). Altogether, these results
indicate that plasmin activity is required for complete myoblast fusion
and differentiation in vitro.
Plasmin production in regenerating muscle is dependent on uPA activity We next investigated the functional relevance of plasmin during skeletal muscle regeneration in vivo. Muscle regeneration was induced in mice by intramuscular injection of 50% glycerol.17 Using the chromogenic substrate S-2251 for the activity of plasmin, we had shown previously that plasmin generation was increased during skeletal muscle regeneration in WT mice, with a peak of activity occurring 2 to 4 days AI and decreasing by day 10.15To investigate whether the generation of plasmin from Plg is uPA- or
tPA-dependent, we analyzed uPA and tPA activities in regenerating
muscle tissues at different times AI. Tissue extracts were prepared
from noninjured muscle (0 days AI); from 2-, 4-, and 7-day-injured
muscle (2, 4, and 7 days AI); and analyzed by zymography
(Figure 3). As mentioned above, this
assay allows clear distinction between uPA and tPA, which migrate at 45 and 72 kd, respectively. No uPA activity was detectable in tissue
extracts from noninjured muscle. However, it was induced in the
regenerating muscle samples, reaching a peak at 4 days AI, 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 muscle extracts analyzed, while
purified human uPA (55 kd), utilized as a control for activity (C), was detected in an adjacent lane. These results suggested that uPA, rather
than tPA, might be involved in Plg activation during skeletal muscle
regeneration in vivo.
Expression of myogenin is reduced in regenerating muscle tissue of Plg-deficient mice To evaluate the consequences of loss of plasmin in the skeletal muscle regeneration process, we induced muscle injury in WT and Plg/ mice by glycerol injection, and the expression
myogenin was analyzed because the expression of this transcription
factor is known to be induced in satellite cells during muscle
regeneration in vivo. RNA and protein lysates were prepared from
muscles on the fourth day after injury (W, wound) as well as from
noninjured muscles (C, control). As shown by Northern blotting,
myogenin RNA levels were very low in quiescent muscle from WT mice and
were dramatically induced in injured muscle (Figure
4A, compare C and W in WT), indicating
that the regeneration process was proceeding normally. However,
myogenin transcript levels were induced to a lesser extent in
Plg-deficient mice than in WT mice after injury (Figure 4A, compare W
in WT and Plg/), while the 18S RNA level was comparable
in all lanes. Similarly, following muscle injury, myogenin protein
levels were induced to a greater extent in WT mice than in
Plg-deficient mice (Figure 4B, compare W in WT and
Plg/). These results indicated that Plg is
required for efficient expression of muscle regeneration specific gene
products. As a control for the the Plg/ genotype, PCR
genotyping of WT, Plg/, and Plg+/ mice
was performed using specific primers. As shown in Figure 4C, PCR
products of 450 and 480 bp, corresponding to WT Plg allele and targeted
allele, respectively, were obtained.
Plg-deficient mice show a severe regeneration defect with enhanced fibrosis and myotube degeneration To analyze the functional significance of the increased plasmin activity in skeletal muscle after damage, we performed glycerol-induced muscle injury in WT mice and in Plg/
mice, and we analyzed comparatively the histopathological changes induced by the lesion in both types of mice. Skeletal muscle of Plg-deficient mice showed a dramatic regeneration defect after glycerol-induced injury, whereas regeneration in WT mice proceeded normally (Figure 5A). This regeneration
defect in Plg/ mice was apparent by 4 days AI but was
most striking 10 to 30 days after injury. Analysis of H/E-stained
cross-sections of WT and Plg-deficient mice 2 days after injury showed
that muscles of all mice were edematous, presenting fibrotic
infiltrates within the enlarged intercellular space separating the
necrotic myofibers. Cross-sections of muscles 4 days after injury
already showed features of ongoing regeneration in WT mice, with many
new myofibers characterized by small size and single nuclei, and with a
reduction in fibrotic infiltrates (Figure 5A). In contrast, in
Plg-deficient mice, at 4 days AI the muscle appeared edematous and no
new uninucleated, small myofibers were detected yet. In WT mice, at 7 days AI most injured fibers regenerated into groups of centrally
nucleated myotubes, an indication of advanced regeneration, and a very
small number of necrotic fibers could be observed; in addition, the size of newly formed myofibers had increased in the WT animals. In
contrast, in Plg-deficient mice, the muscle appeared necrotic, showing
still extensive fibrosis 7 days after damage. Ten days after injury,
the muscle of WT mice was completely regenerated; no sign of previous
damage was detectable in WT mice, except for the presence of centrally
located nuclei inside the regenerated fibers (Figure 5A). In
Plg-deficient mice, however, a high number of degenerated myotubes were
still visible. Thirty days after injury, the lesion was no longer
noticeable in WT mice, except for the central myonuclei. In
Plg/ mice, however, the muscle presented extensive
fibrosis with high numbers of degenerated myotubes. Notably, extensive
muscle degeneration was still observed in Plg-deficient mice 50 days
after injury, suggesting that regeneration is not only retarded, but
dramatically impaired, in mice deficient in Plg (see Figure
5A).
To confirm further the muscular regeneration defect in
Plg Plg deficiency alters the inflammatory response after muscle injury Circulating inflammatory cells are recruited after injury to skeletal muscle during the inflammatory phase.3 While neutrophils have been detected in the early hours following muscle injury, maximal levels of macrophages and T lymphocytes appear later during the muscle injury response. It is widely held that Plg system plays a role in inflammation through plasmin-mediated directional cell migration. To directly characterize plasmin's involvement in the inflammatory response following muscle injury, the presence of neutrophils, macrophages, and T lymphocytes in normal and in regenerating muscle of WT and Plg-deficient mice was analyzed by immunohistochemistry using antibodies against Gr-1, Mac-1, and T11, well-characterized neutrophil, macrophage, and T-lymphocyte markers, respectively. No Gr-1+ cells were detected in noninjured muscle sections of WT or Plg-deficient mice (Figure 6A, 0 days AI). Twelve hours after injury, there was a similar increase in Gr-1-expressing cells at the sites of injury in both types of animals (Figure 6A, 0.5 days AI, P < .05). Only a few resident Mac-1+ cells were detected in the intact muscle sections of WT and Plg/ mice (Figure 6B, 0 days AI). Two days AI, the
number of Mac-1-expressing cells augmented at the injury site of both
types of animals; however, the number of Mac-1+ cells in
Plg-deficient mice was reduced to almost 50% with respect to WT mice
(Figure 6B, 2 days AI, P < .05). Similarly, 2-days AI the
number of T11-expressing cells increased with respect to resting muscle
in both WT and Plg-deficient mice; however, the number of
T11-expressing cells at the injury site in muscles of Plg-deficient
mice was 35% lower than in WT mice (Figure 6C, 2 days AI,
P < .05). Quantification of these data are represented in
Figure 6D. These results suggested that the recruitment of both
macrophages and T lymphocytes to the injured muscle was reduced in the
absence of plasmin.
Systemic fibrinogen depletion restores muscle regeneration in Plg-deficient mice The classical role of plasmin is the degradation of fibrin deposits or fibrinolysis (reviewed by Lijnen and Collen23). In a previous study, we demonstrated an accumulation of fibrin following muscle injury in Plg-deficient mice, but not in WT mice, correlating with the persistence of muscular degeneration features in Plg/ mice after muscular
injury.15 Depletion of circulating fibrinogen can be
achieved by administration of the viper venom ancrod.15,24 Ancrod- or saline-delivering osmotic pumps (3 U/d) were implanted in
Plg/ mice during 3 days before muscle injury and then
throughout the experimental period (up to 10 days AI). Western blot
analysis using an antifibrinogen antibody demonstrated the reduction of plasma fibrinogen upon ancrod administration, without any effect on
survival (Figure 7A, compare ancrod and
saline). Furthermore, signs of improved muscular regeneration were
detectable in ancrod-treated Plg-deficient mice 10 days after injury,
with centrally located nuclei inside the regenerated fibers. In
contrast, histologic features of muscle degeneration persisted in
saline-treated Plg-deficient mice at the same stage AI, including high
numbers of degenerated myotubes and fibrosis in the intercellular space
throughout the damaged muscle (Figure 7B). The extent of muscle
regeneration of ancrod-treated Plg-deficient mice 10 days AI resembled
the regeneration status of a WT mouse at the same time AI. This
observation supports the idea of a potential pathogenic role of
fibrin(ogen) accumulation during muscle regeneration in
Plg-deficient mice.
The aim of our work was to investigate the involvement
of the broad-spectrum serine protease plasmin in cellular and tissular remodeling occurring during skeletal myogenesis in vitro and skeletal muscle regeneration in vivo. Our results clearly show that inhibition of plasmin activity by Extravascular fibrin deposition is a key feature in pathologies characterized by inflammation and tissue repair, including impaired skin wound healing25 and glomerulonephritis.26 In these 2 situations, failure to remove plasmin was attributable to reduced uPA- or tPA-mediated fibrinolysis, respectively. In experimentally induced muscle degeneration, we also found that uPA deficiency led to increased levels of muscular fibrin, indicating the importance of uPA in extravascular fibrin clearance.15 In the present study, using Plg-deficient mice we have demonstrated the importance of plasmin for fibrin clearance following muscle injury. Fibrin accumulation in the extracellular basal membrane may have deleterious effects, such as the impediment of normal nutrition to the muscle tissue. The hypothesis that increased fibrin levels may contribute to impaired muscular regeneration was further explored by systemic fibrinogen depletion of Plg-deficient mice. Administration of ancrod reduced plasma fibrinogen levels in Plg-deficient mice, resulting in an almost complete restoration of the normal muscle regeneration process. These results clearly show that fibrin(ogen) accumulation has a pathogenic role in sustaining muscle degeneration. Our finding that loss of plasmin activity impedes muscle regeneration
raises the question of how this proteolytic activity 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, neutrophils, macrophages, and T lymphocytes accumulated near
the injury site in the first days AI. Similar results in macrophage
recruitment were reported in a study of regeneration following
bupivacaine-induced muscle injury in the rat27 and
following crush injury in mouse.3 We have observed that
mice with a specific deficit in Plg show a reduced staining for
Mac-1+ and T11+ cells 48 hours AI, indicating
that the number of macrophages and T lymphocytes reaching the injury
site is reduced in the absence of Plg. Recently, deficient macrophage
recruitment was also reported by our group in regenerating muscle of
uPA-deficient mice.15 This suggests that uPA/plasmin
activity may have a profound effect on inflammation and
inflammation-related muscular disease. Previous studies with
uPA-deficient mice 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.28 Similarly, monocyte and lymphocyte
recruitment was significantly diminished in Plg-deficient mice after
thioglycollate-induced acute peritoneal inflammation.29 Additionally, components of the Plg system may regulate the expression or/and activity of cytokines involved in inflammatory processes. For
example, endogenously produced uPA can amplify tumor necrosis factor- The hypothesis that, in general, Plg activation facilitates cellular
penetration of fibrin-containing matrices fits 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 serineproteases, metalloproteases,
and other classes of proteases, which together serve in matrix
remodeling and cellular reorganization of wound fields. Our data
suggest that plasmin is particularly useful in fibrin solubilization
and, without it, cellular reorganization of fibrin reach matrices is
severely impeded. We conclude that the uPA/Plg system is required for
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 Plg-deficient animals may be the
diminished ability of responding macrophages to proteolytically dissect
their way through the extracellular matrix to 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 Plg is fibrin. However, based on the existing data, we
cannot exclude that the plasmin 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. Our results have shown that plasmin is required for myogenesis in vitro, because inhibition of plasmin activity with In conclusion, our results demonstrate that plasmin 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 dystrophic muscular therapies. Future experiments will be directed toward further definition of the benefit of defibrinogenating, anticoagulant, or fibrinolytic agents in dystrophinopathies. This report constitutes the first demonstration for a role of plasmin in skeletal myogenesis in vitro and muscle regeneration in vivo. Rescue experiments will show whether exogenously supplied plasmin can compensate for the mutation. If this is the case, PAIs might prove to be useful for the treatment of muscle injuries or muscle dystrophies, particularly if uPA/plasmin evokes the response preferentially in muscle.
We are very grateful to Dr Keld Dano for the antifibrinogen
antibody and Dr R. Lijnen for
Submitted June 26, 2001; accepted December 11, 2001.
Supported by grants from the European Union 1999/C361/06 (E.S., G.A., M.S.), DGES PM97-008, Fundació La Marató-TV3 (F.L., M.P.), and FIS 01/1474. M.S. was also supported by the Catalan government (Reincorporació de Doctors).
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 latent basic fibroblast growth factor, and these
activities have been proposed to be crucial for cell migration and
tissue remodeling in vivo. Thus, the plasmin system has been shown to play a role in a wide variety of physiopathological processes involving
extracellular matrix degradation, tissue remodeling, and cell
migration, including ovulation, trophoblast invasion, postlactational
mammary involution, neurite outgrowth, wound healing, inflammation,
angiogenesis, and tumor cell invasion (reviewed by Irigoyen et
al
) or presence of 
), SB203580 (
), or BSA (
). Fusion is represented as
percentage of nuclei in myotubes. Values are mean of 3 independent
experiments. (B) Lower-power views of representative fields.
Preconfluent cells in DMEM/10% FBS (i), confluent cells in
DMEM/insulin (ii), confluent cells in DMEM/insulin plus
