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(American Journal of Pathology. 2005;167:1105-1117.)
© 2005 American Society for Investigative Pathology

NS-398, a Cyclooxygenase-2-Specific Inhibitor, Delays Skeletal Muscle Healing by Decreasing Regeneration and Promoting Fibrosis

Wei Shen*{dagger}, Yong Li{dagger}{ddagger}, Ying Tang{dagger}, James Cummins{dagger} and Johnny Huard*{dagger}{ddagger}

From the Departments of Bioengineering* and Orthopaedic Surgery,{ddagger} University of Pittsburgh, Pittsburgh; and the Growth and Development Laboratory,{dagger} Children’s Hospital of Pittsburgh, Pittsburgh, Pennsylvania


   Abstract
Top
 Abstract
Materials and Methods
 Results
 Discussion
 References
 
Nonsteroidal anti-inflammatory drugs are often prescribed after muscle injury. However, the effect of nonsteroidal anti-inflammatory drugs on muscle healing remains primarily controversial. To further examine the validity of using these drugs after muscle injury, we investigated the working mechanism of NS-398, a cyclooxygenase-2-specific inhibitor. In vitro experiments showed that NS-398 inhibited the proliferation and maturation of differentiated myogenic precursor cells, suggesting a detrimental effect on skeletal muscle healing. Using a mouse laceration model, we analyzed the in vivo effect of NS-398 on skeletal muscle healing at time points up to 4 weeks after injury. The in vivo results revealed delayed muscle regeneration at early time points after injury in the NS-398-treated mice. Compared to controls, lacerated muscles treated with NS-398 expressed higher levels of transforming growth factor-ß1, which corresponded with increased fibrosis. In addition, transforming growth factor-ß1 co-localized with myostatin, a negative regulator of skeletal muscle growth. We also found reduced neutrophil and macrophage infiltration in treated muscles, indicating that the delayed skeletal muscle healing observed after NS-398 treatment could be influenced by the anti-inflammatory effect of NS-398. Our findings suggest that the use of cyclooxygenase-2-specific inhibitors to treat skeletal muscle injuries warrants caution because they may interfere with muscle healing.

Muscle injuries due to trauma, sports, or military-related circumstances occur frequently. After injury, the healing process comprises usually sequential stages that include degeneration and inflammation, muscle regeneration, and fibrosis.1,2 The release of inflammatory metabolites, including prostaglandins, leukotriene, and thromboxane, produces pain, vasodilation, and a series of inflammation-related symptoms. To relieve pain, doctors for years have prescribed nonsteroid anti-inflammatory drugs (NSAIDs), a large and chemically diverse group of drugs that inhibit the cyclooxygenase (COX) pathways and thereby block the transition of arachidonic acid into prostaglandins and thromboxane. By so doing, these drugs relieve the pain associated with inflammation. Some NSAIDs also reportedly inhibit neutrophil aggregation,3,4 suppress the production of superoxide and nitric oxide by inflammatory cells,5,6 and disrupt the intracellular signaling in activated inflammatory cells, which contributes to the drugs’ anti-inflammatory effects.7 The use of traditional NSAIDs can result in numerous side effects, including gastrointestinal bleeding,8 liver damage,9 and heart failure.10 Identification of the constitutive (Cox-1) and inducible (Cox-2) isoforms of the Cox enzyme has resulted in widespread use of Cox-2-specific inhibitors (rather than traditional ones) to reduce the frequency and severity of these side effects.

Although some studies have shown that the administration of NSAIDs promotes muscle healing by reducing degeneration and inflammation,7,11 other research has demonstrated that NSAIDS are detrimental to the entire healing process.12-14 In response to increasing debate regarding both the beneficial and detrimental effects of the Cox-2-specific inhibitors, we performed this study to investigate the influence of these drugs on inflammatory responses, muscle regeneration, fibrosis, and, consequently, overall muscle healing after injury.

Here we examined the effects of NS-398, a Cox-2-specific inhibitor, both on myogenic precursor cells in vitro and in a mouse skeletal muscle laceration model in vivo. The administration of NS-398 inhibited the proliferation and maturation of differentiated myogenic precursor cells in vitro. Similarly, NS-398 delayed the maturation of regenerating myofibers in vivo at early time points (3 days, 5 days, and 7 days after injury) and led to fibrosis. However, at a later time point (28 days after injury) the control and NS-398-treated muscles appeared histologically similar. Further investigation indicated that the delayed healing in the treated muscles may have been due to reduced inflammatory responses and the up-regulation of transforming growth factor (TGF)-ß1 and myostatin after NS-398 administration—effects shown to be related to the dosage of NS-398 and the duration of its administration.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
Cell Isolation and Culturing

Myogenic precursor cells were isolated via a previously described preplate technique.15,16 Muscles were removed from 4-week-old C57BL/6J mice (Jackson Laboratories, Bar Harbor, ME). The muscles were minced with a surgical blade and were enzymatically digested by sequential exposure to collagenase type XI, dispase, and trypsin. The muscle cell extracts then were plated on collagen-coated flasks. Different populations were isolated by replating the extracts after different time intervals. The late preplate population contains cells with higher myogenic potential than cells in the earlier preplate populations.16 Late preplate cells were used for our in vitro experiments and were maintained in proliferation medium (Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum, 10% horse serum, and 0.5% chicken embryo extract).

Cell Proliferation and Differentiation

The myogenic precursor cells used for the cell proliferation experiments were plated at the same cell density per well in 12-well plates. On day 0, cells were grown overnight in serum-free medium to synchronize the cells by starvation. On day 1, the serum-free medium was replaced with proliferation medium. Different concentrations of NS-398 (0, 1, 10, or 100 µmol/L; Cayman Chemical, Ann Arbor, MI), selected on the basis of previous studies involving NS-398,17-19 were dissolved in culture media (with dimethyl sulfoxide, 10 mg/ml, as the stock solution) to examine the effect of NS-398 on cell proliferation. The proliferation medium and NS-398 were replenished on day 3. After cells were trypsinized and collected on days 1 to 4, a hemocytometer was used to count the cells manually. Cells in three wells/NS-398 concentration were counted each day.

The myogenic precursor cells used for the cell differentiation experiments were plated at the same cell density per well in six-well plates. On day 0, cells were grown overnight in serum-free medium to synchronize the cells by starvation. The following day, the serum-free medium was removed and replaced with differentiation medium (Dulbecco’s modified Eagle’s medium supplemented with 1% fetal bovine serum and 1% horse serum) supplemented with different concentrations of NS-398. As described above, the cells were permitted to grow for an additional 2 days. At that time, the cells and supernatants were collected for Western blot analysis and enzyme-linked immunosorbent assay (ELISA).

ELISA

Supernatants collected from the differentiation experiments were analyzed by ELISA. Analysis of PGE2 and PGF2{alpha} was performed as suggested in the instructions provided by the manufacturer (DE0100 PGE2 ELISA kit, DE1150 PGF2{alpha} ELISA kit; R&D Systems, Minneapolis, MN).

Animal Model

The gastrocnemius muscles (GMs) of 96 mice (C57BL/6J, female, 5 weeks of age; Jackson Laboratories) were lacerated in accordance with the injury model detailed below. The Animal Research and Care Committee at the authors’ institution approved all experimental protocols (protocol 5/01). The muscle injury model, developed in mice and used in previous studies,20,21 entailed laceration of the GMs in both legs. The mice were anesthetized by intramuscular injection of 0.03 ml of ketamine (100 mg/ml) and 0.02 ml of xylazine (20 mg/ml). A surgical blade (no. 11; SteriSharps, Mansfield, MA) was used to lacerate each GM at 60% of its length from its distal insertion through the lateral 50% of muscle width and 100% of muscle thickness. After laceration, the skin was closed with black silk 4-0 suture (Ethicon, Somerville, NJ).

NS-398 (Cayman Chemical) was dissolved in dimethyl sulfoxide and injected intraperitoneally into the mice, which received one of two doses (5 mg/kg of body weight or 10 mg/kg of body weight) for one of two durations (3 days or 5 days continuously) immediately after surgery. The same amount of only dimethyl sulfoxide (no NS-398) was injected intraperitoneally into other mice that served as the negative control. Thus, the mice used in the in vivo study received one of four treatment regimens: 5 mg NS-398 per day for 3 days (5 mg-3 day group), 5 mg NS-398 per day for 5 days (5 mg-5 day group), 10 mg NS-398 per day for 3 days (10 mg-3 day group), or only dimethyl sulfoxide (control group). The mice were sacrificed at different time points after injury (3, 5, 7, 10, 14, or 28 days), and the GMs were harvested from both legs, flash-frozen in 2-methylbutane precooled in liquid nitrogen, and stored at –80°C pending histological analysis.

Hematoxylin and Eosin (H&E) Staining

Cryosections were fixed in 1% glutaraldehyde for 1 minute, and then were dipped in hematoxylin for 30 seconds. After being washed with alcohol acid and ammonia water, they were immersed in eosin for 15 seconds. After each step, sections were rinsed with distilled water. The sections then were dehydrated by treatment with alcohols of increasing concentrations (70%, 80%, 95%, and 100%). Finally, the sections were treated with xylene and covered with glass slips.

Slides were analyzed manually via bright-field microscopy (Eclipse E800; Nikon, Tokyo, Japan) and by using Northern Eclipse software (Empix Imaging, Cheektowaga, NY). Sections containing the largest injury area were analyzed. The centronucleated regenerating myofibers in those sections were counted under x100 magnification (four animals per group), and results were recorded as the number of centronucleated myofibers/total number of myofibers in each section. In addition, an image of the central injury area was taken at x200 magnification, and the minor axis diameters (ie, the smallest diameter) of 200 centronucleated myofibers in each of the images were measured.

Immunohistochemistry

Standard techniques were used to prepare serial 8-µm cryostat sections and cell cultures. For immunohistochemistry, the following primary antibodies were used at the indicated dilutions: monoclonal mouse anti-myosin heavy chain-developmental (MHC-d) (1:100; Novocastra Laboratories, Ltd., Newcastle, UK), monoclonal mouse anti-MHC-mature (MHC-m) (1:150, M4276; Sigma, St. Louis, MO), rabbit anti-mouse collagen IV antibody (1:200; Biodesign, Saco, ME), rat anti-mouse TGF-ß1 (1:100; BD Biosciences Pharmingen, San Diego, CA), rabbit anti-mouse myostatin (1:200; Chemicon, Temecula, CA), polyclonal rabbit anti-desmin (D8281, 1:200; Sigma), monoclonal anti-mouse Pax7 (1:200; Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA), monoclonal rat anti-F4/80 (1:200; Abcam, Cambridge, MA), rat anti-CD-11b (1:150; Chemicon), and monoclonal anti-{alpha}-smooth muscle actin (fluorescein isothiocyanate-conjugated, 1:150; Sigma). The sections and cell cultures then were exposed to the following secondary antibodies for 50 minutes at room temperature: anti-mouse-conjugated Cy3 (1:250, Sigma) and anti-rabbit-conjugated fluorescein (1:100, Sigma). Collagen type IV was co-localized with MHC-d, MHC-m, and TGF-ß1, whereas myostatin was co-localized with TGF-ß1. Negative controls (stainings without the primary antibody) were performed concurrently with all immunohistochemical staining. The nuclei of the sections were revealed via 4,6-diamidino-2-phenylindole staining (Sigma). Fluorescent microscopy was used to visualize all immunofluorescence results (Nikon E800). Regenerating myofibers expressing MHC-d or MHC-m were counted at x200 magnification (slides from four animals per group). The area of TGF-ß1 expression (red after immunostaining) was measured with Northern Eclipse software (x200 magnification, slides from four animals per group; Empix Imaging).

Western Blot Analysis

Laemmli sample buffer (161-0737; Bio-Rad, Hercules, CA) was used to collect proteins from live cells. After being prepared via standard procedures, protein samples were separated on 10% sodium dodecyl sulfate-polyacrylamide electrophoresis gel and were transferred to nitrocellulose membranes that then were used to perform immunoblotting. Mouse anti-MyoD (554130, 1:250; Pharmingen), anti-myogenin (556358, 1:250; Pharmingen), and anti-MHC-d (1:500, Novocastra) were applied as primary antibodies, and mouse anti-ß-actin (1:8000, Sigma) was used for protein quantification. The horse-radish peroxidase-conjugated secondary antibodies (Pierce, Rockford, IL) were diluted to 1:5000 and applied. Blots were developed by using SuperSignal West Pico chemiluminescent substrate (Pierce), and positive bands were visualized on X-ray film. All results were analyzed with Northern Eclipse software (Empix Imaging).

Trichrome Staining

Trichrome staining was performed to determine the collagen content of the muscle tissue. Slides were processed as detailed in the manufacturer’s protocol (Masson Trichrome stain kit, K7228; IMEB, Inc., Chicago, IL). Northern Eclipse software (Empix Imaging) was used to measure the area of fibrous tissue (slides from three samples per group, x100 magnification).

Flow Cytometry Analysis

The GMs from nontreated (control) and NS-398-treated groups (10 mg-3 days group) were surgically removed before injury or 12 hours, 24 hours, 48 hours, 3 days, or 5 days after injury. Collagenase, dispase, and trypsin were used to digest the tissue matrix and isolate the cells. Debris was removed via filtration with 100-µm filters. Isolated cells first were treated with 10% mouse serum (Sigma) to block nonspecific binding sites. Primary rat anti-CD-11b (conjugated with fluorescein isothiocyanate, R&D Systems) and rat anti-F4/80 (conjugated with APC; Serotec, Raleigh, NC) antibodies were used in combination to distinguish the neutrophil and macrophage populations: F4/80 is specific to macrophages22 and CD-11b is expressed by macrophages and neutrophils.23,24 7-Amino-actinomycin D (7-AAD; Pharmingen) was added to all tubes to exclude nonviable cells from the analysis. Marked cell samples then were analyzed with a FACS Caliber flow cytometer (BD Biosciences) and CellQuest software (BD Biosciences).

Statistics

A Student’s t-test was used to compare the differences in MHC-d and MHC-m expression between the control and NS-398-treated groups. A {chi}2 test was used to analyze the percentage differences in the numbers of neutrophils and macrophages identified via flow cytometry. All other data were analyzed by one-way analysis of variance statistical analysis. Error bars on the figures represent the SD. P < 0.05 was considered statistically significant.


   Results
Top
 Abstract
 Materials and Methods
 Results
Discussion
 References
 
NS-398 Inhibited the Proliferation and Differentiation of Myogenic Cells in Vitro

Late preplate cells (preplate 5) isolated via the preplate technique15,16 were immunostained for two well-known markers of early myogenesis: desmin and Pax7.25-27 More than 90% of the cells were desmin-positive and 85% were Pax7-positive (data not shown). This finding confirms that these cells are myogenic precursor cells. NS-398 at a concentration of 100 µmol/L significantly inhibited the proliferation of late preplate cells at day 3 and day 4 (Figure 1A) . The addition of NS-398 (10 µmol/L or 100 µmol/L) to cells cultured in differentiation medium for 2 days significantly decreased their expression of the late myogenic differentiation markers myogenin and MHC-d but not of the early marker MyoD (P < 0.05; Figure 1, B and C ). The addition of NS-398 at any of the tested concentrations significantly inhibited the expression of PGE2 and PGF2{alpha} in culture supernatant (P < 0.05; Figure 1D ).



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  		Figure 1. A: NS-398 (100 µmol/L) significantly inhibited the proliferation of myogenic precursor cells at day 3 and day 4. B and C: Western blot results show that NS-398 (10 µmol/L and 100 µmol/L) decreased the expression of MHC-d and myogenin but not of MyoD. D: The addition of all tested concentrations of NS-398 in vitro severely reduced the expression of prostaglandins by myogenic precursor cells. The asterisks indicate a significant difference (P < 0.05) between the marked groups and the 0 µmol/L group.

 
Effect of NS-398 on Muscle Regeneration

H&E staining revealed the presence of centronucleated regenerating myofibers in both the NS-398-treated groups and the nontreated (control) group as early as 3 days after injury (results not shown). Seven days after injury, the injury site primarily contained regenerating myofibers. Seven and fourteen days after injury, significantly more regenerating myofibers were present in the nontreated control group than in each of the NS-398-treated groups (P < 0.05). Furthermore, the centronucleated myofibers in the nontreated control group were significantly larger than those in each of the NS-398-treated groups (P < 0.05), which indicates more advanced muscle regeneration in the control group. The dosage of NS-398 and the duration of NS-398 administration also contributed to these effects. Twenty-eight days after injury, the control group and the NS-398-treated groups exhibited similar degrees of muscle recovery. There were no significant differences between groups in terms of the number or size of regenerating myofibers at this time point (Figure 2) .



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  		Figure 2. H&E staining of muscle sections harvested 7 days (A and B), 14 days (C and D), and 28 days (E and F) after injury are shown. Seven and fourteen days after injury, muscle sections in the control group contained a significantly higher percentage of centronucleated myofibers than did muscle sections in the NS-398-treated groups. Furthermore, the minor axis diameters of the centronucleated myofibers in the control group were significantly larger than those of the centronucleated myofibers in the NS-398-treated groups. However, follow-up analysis of these parameters conducted 28 days after injury revealed no significant difference between the control and treated groups. The asterisks indicate a significant difference (P < 0.05) between the marked groups and the control group. Original magnifications: x200 (A–F); x100 (insets).

 
We stained muscle cryosections from all groups to assess MHC-d (immature MHC, Figure 3 ) and MHC-m (mature MHC, Figure 3 ) expression, which together serve as indicators of myofiber regeneration. We observed MHC-d and MHC-m expression within regenerating myofibers in injured skeletal muscle. Three days after injury, all groups contained numerous regenerating myofibers expressing MHC-d. At subsequent time points, the number of MHC-d-expressing myofibers began to decrease as we observed a transition from MHC-d (immature) to MHC-m (mature) expression. This transition was obvious and occurred rapidly in the control group (within 7 days after injury), whereas muscles harvested in the NS-398-treated groups continued to contain primarily MHC-d-expressing myofibers 7 days after injury and had just begun to exhibit this transition from immature to mature myofibers (Figure 3) . Our analysis to determine the different types of cells present within the injured area 5 days after injury revealed MHC-d- and MHC-m-expressing myofibers, immune cells (identified on the basis of F4/80 and CD-11b), and fibroblastic cells (identified by {alpha}-smooth muscle actin). Our comparison of the percentage of immune cells and fibroblastic cells found in the control group versus those found in the NS-398-treated group (10 mg-3 days) at this time point revealed no significant difference; however, as shown in Figure 3N , there was a significant difference in terms of the percentages of MHC-d- and MHC-m-expressing myofibers (P < 0.05).



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  		Figure 3. Immunostaining results of the injured area reveal the expression of MHC-d (red) and MHC-m (red) 3 days (A–D), 5 days (E–H), and 7 days (I–L) after injury. Collagen IV and cell nuclei are stained green and blue, respectively. During this period, the percentage of MHC-d-expressing myofibers in the muscle sections from the control group dropped from 1.42 to 1.11 to 0%, while the percentage of MHC-m-expressing myofibers increased from 0 to 5.34 to 6.65%. During the same period, the percentage of MHC-d-expressing myofibers in the muscle sections from the NS-398-treated group (10 mg-3 days) changed from 3.69 to 5.16 to 4.95%, while the percentage of MHC-m-expressing myofibers increased from 0 to 1.23 to 2.19%. Although we observed an isoform transition from MHC-d to MHC-m in both the NS-398-treated group and the control group during muscle regeneration, the transition was delayed in the NS-398-treated group. The asterisks in M and N indicate a significant difference (P < 0.05) between the control group and the NS-398-treated group (10 mg-3 days) in terms of the MHC-d-expressing myofibers and MHC-m-expressing myofibers. N depicts the densities of the main cell types (cells per unit area) found within the muscle 5 days after injury, including myogenic cells (MHC-d, MHC-m), immune cells (F4/80, CD-11b), and fibroblastic cells ({alpha}-smooth muscle actin). Original magnifications, x200.

 
Effect of NS-398 on Fibrosis

We used trichrome staining to observe fibrous tissue formation after injury in the different groups (Figure 4) . The results are comparable with those observed after H&E staining. At early time points (14 days after injury), the control group exhibited better recovery and contained less fibrous tissue than the NS-398-treated groups (P < 0.05). Muscles from the low-dose, short-duration group (5 mg-3 days) contained less fibrous scar tissue (P < 0.05) than muscles in the high-dose group (10 mg-3 days) or muscles in the longer NS-398 treatment group (5 mg-5 days). Although it did not reach statistical significance, the percentage of fibrous tissue in the NS-398-treated groups appeared to be higher than that recorded for the control group at the 28-day time point.



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  		Figure 4. Trichrome staining was used to observe fibrosis after injury. Fibrous tissue is stained blue. At one of the earlier time points (14 days after injury), the control group contained less fibrous tissue (less stained area) than did either of the NS-398-treated groups (P < 0.05). The low-dose, short-duration group (5 mg-3 days) contained less fibrous tissue than the high-dose (10 mg-3 days) and longer duration (5 mg-5 days) groups (P < 0.05). However, at the last time point (28 days after injury), no significant differences were observed. The asterisks indicate a significant difference (P < 0.05) between the marked groups and the control group. Original magnifications, x100.

 
Effect of NS-398 on the Expression of TGF-ß1 and Myostatin

We used double-labeled immunostaining to investigate the possible roles of TGF-ß1 and myostatin in muscle healing. In both the control group and the NS-398-treated group (10 mg-3 days), the first wave of TGF-ß1 expression was detectable on day 1 and day 3 after injury. We detected myostatin expression beginning on day 3 after injury, with most of the myostatin co-localizing with the TGF-ß1 expression (Figure 5, E, F, G, and H ; results from 5 mg-3 days group not shown). We found that 92% and 87% of the myostatin-expressing myofibers co-expressed TGF-ß1 in the control group and NS-398-treated group (10 mg-3 days), respectively. Our analysis of the percentage of myofibers expressing TGF-ß1 and myostatin 3 days after injury revealed a significantly higher percentage in the NS-398-treated muscles than in the controls (P < 0.05). A second wave of TGF-ß1 expression occurred in the extracellular matrix of all groups at later time points (7, 10, and 14 days after injury; Figure 6 ). The control muscles expressed significantly less TGF-ß1 than the NS-398-treated muscles (P < 0.05). This finding supports the possible involvement of TGF-ß1 in the promotion of fibrosis observed in the NS-398-treated muscles.



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  		Figure 5. Immunostaining results for TGF-ß1 (red) and myostatin expression (green). Cell nuclei are blue. There was a significantly higher percentage of TGF-ß1- or myostatin-expressing myofibers in the muscle sections from the NS-398-treated group than in the muscle sections from the control group 3 days after injury. The expression of myostatin was highly co-localized with the expression of TGF-ß1 in both the control (E and F) and the NS-398-treated groups (10 mg-3 days; G and H). The asterisks indicate a significant difference (P < 0.05) between the marked groups and the control group. Original magnifications, x200.

 


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  		Figure 6. Immunostaining results for TGF-ß1 expression. TGF-ß1, collagen IV, and cell nuclei are red (arrows), green, and blue, respectively. Both the control and the NS-398-treated groups (5 mg-3 days) exhibited relatively low TGF-ß1 expression 7 days after injury (A and D). However, 10 days and 14 days after injury the control group showed hardly any expression of TGF-ß1 (B and C), whereas, relative to the control group, the NS-398-treated groups exhibited significantly higher TGF-ß1 expression (E and F). Asterisks in the graph indicate that the TGF-ß1-expressing area was significantly less in the control group than in the various NS-398-treated groups (P < 0.05). Original magnifications: x200 (A–F); x100 (insets).

 
Effect of NS-398 on Immune Response

Using flow cytometry, we identified a population of CD-11b (fluorescein isothiocyanate)-positive cells and a population of CD-11b/F4/80 (fluorescein isothiocyanate/APC) double-positive cells in both the NS-398-treated groups and the nontreated (control) groups as early as 12 hours after injury; these two populations represent neutrophils and macrophages, respectively. The percentages of these cells (relative to the entire cell population) continued to increase until they peaked 48 hours after injury; these percentages dropped dramatically thereafter. At all time points, the percentages of neutrophils and macrophages in the NS-398-treated group were lower than those in the control group. However, statistically significant differences between the control group and the treated groups occurred only at 48 hours after injury for neutrophils and 24 hours after injury for macrophages (P < 0.05, Figure 7 ).



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  		Figure 7. Neutrophils (A) and macrophages (B) infiltrated the injury site as early as 12 hours after injury. The percentages of these cells (in terms of the entire muscle cell population) peaked 48 hours after injury. At all time points, the NS-398-treated groups contained lower percentages of inflammatory cells than did the control group, but we observed significant differences (P < 0.05) only at 48 hours after injury for neutrophils and at 24 hours after injury for macrophages.

 

   Discussion
Top
 Abstract
 Materials and Methods
 Results
 Discussion
References
 
This study investigated the effect of NS-398, a Cox-2-specific inhibitor, on skeletal muscle healing both in vitro and in vivo. We found that NS-398 1) inhibited the proliferation of myogenic precursor cells and their maturation when induced to differentiate, 2) decreased the regeneration of injured muscle by delaying the maturation of regenerating myofibers, and 3) promoted fibrosis by up-regulating TGF-ß1 expression. NS-398 administration also reduced the inflammatory response and up-regulated myostatin expression. These results suggest that NS-398 interferes with the healing of injured skeletal muscle.

Satellite cells are the cells primarily responsible for the regeneration of muscle tissue. When a muscle is injured, quiescent satellite cells begin to proliferate and generate myogenic cells, which are identifiable by their expression of the early myogenic markers MyoD and Myf5. Before they terminally differentiate to form new myofibers or fuse with previously existing myofibers, satellite cells begin to express myogenin and MRF4 (Figure 8) . The study reported here demonstrates that NS-398 decreased the expression of myogenin and MHC-d by myogenic precursor cells but did not affect MyoD expression. This finding suggests that NS-398 interferes with the maturation of myogenic cells (ie, with late-stage differentiation) but not with their early activation or differentiation.25,27-29 NS-398 administration also inhibited the proliferation of myogenic precursor cells, although a significant difference was only noted at the highest dose (100 µmol/L). Because the expression of prostaglandins (PGE2 and PGF2{alpha}) is severely impaired at low doses of NS-398 (1 µmol/L and 10 µmol/L), this proliferation inhibition may be at least partially due to a Cox-2-independent pathway. The results of trypan blue assay indicated good cell viability (data not shown), and cell numbers continued to increase in all NS-398-treated groups during the 4-day period of the experiment. Therefore the proliferation inhibition does not appear to be due to a toxic effect of NS-398. In a recently published study, Mendias and colleagues17 showed that NS-398 can inhibit the proliferation and differentiation of rat satellite cells, a finding that our results strongly support. In their study, however, NS-398 concentrations ranging from 0.01 µmol/L to 100 µmol/L showed similar inhibitory effects on the proliferation of the rat satellite cells. This discrepancy between their results and ours may be related to species-related differences.



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  		Figure 8. Proposed mechanism of myogenic differentiation. Different markers distinguish various stages of myogenic differentiation. MyoD and myogenin denote the early and late stages of muscle differentiation, respectively.27,29,62-64 The detrimental effects of NS-398 on myogenic cells likely occur via inhibition of both cell proliferation and maturation of differentiated myogenic cells.

 
We used the number of centronucleated regenerating myofibers and their minor axis diameters to quantify histological recovery of the injured muscles. Results from 7 days and 14 days after injury demonstrated better recovery in the control group than in the NS-398-treated groups. The low-dose group (5 mg-3 days) showed better recovery than the high-dose and the longer NS-398 treatment groups (5 mg-5 days and 10 mg-3 days, respectively). However, 28 days after injury, all groups exhibited similar final outcomes. We observed that NS-398 delayed muscle healing and that this effect was dependent on the dose of NS-398 and the duration of its administration. We hypothesize that, after cessation of NS-398 treatment and subsequent clearance of the compound from the body, the normal healing process resumes. Using a muscle freeze injury model, Bondesen and colleagues30 found that SC-236, another Cox-2-specific inhibitor, decreased the size of regenerating myofibers, even regenerating myofibers analyzed 5 weeks after injury. That study, however, involved the ongoing administration of SC-236 for the duration of the 5-week-long experiment, unlike our study that was administered for a maximum of 5 days. The prolonged inhibition of muscle healing observed in that study may be due to the extended administration of the Cox-2-specific inhibitor and gives more credence to the argument in favor of using these drugs only with caution—especially throughout the long term.

We also evaluated the regeneration of the injured muscle by detecting and measuring the expression of MHC-d and MHC-m. MHC is a major protein expressed in skeletal muscle. One of its isoforms, the developmental MHC (MHC-d), is expressed in embryonic and early regenerating myofibers.31,32 The appearance of MHC-d-expressing myofibers also correlates with the onset of muscle regeneration. During the maturation of regenerating myofibers, the isoform undergoes a transition from MHC-d (immature) to MHC-m (mature).33,34 In the control group and the NS-398-treated groups, the MHC-d-expressing myofibers appeared as early as 3 days after injury. However, the transition of these immature myofibers into mature myofibers was delayed in the NS-398-treated groups. This finding suggests that the administration of NS-398 in vivo can delay the maturation of regenerating myofibers and consequently slow the healing process. This phenomenon correlates with our in vitro findings, which show that NS-398 interfered with the maturation of differentiated myogenic precursor cells but not with their early activation or differentiation.

Fibrosis frequently occurs after injury in many organs and tissues. When fibrous tissue forms in an injured area, the recovery of normal tissue architecture and functionality is compromised. After muscle injuries, fibrosis causes decreased muscle contractility and range of movement.35 More importantly, the resultant fibrous tissue makes the repaired muscle more susceptible to reinjury. The results of trichrome staining in our study showed that fibrosis began as early as 10 days after injury and that fibrous tissue (rather than regenerating myofibers) filled the injury sites. The NS-398-treated muscles contained fewer regenerating myofibers and more fibrous tissue (in terms of area) than did control muscles. However, as demonstrated by the results of H&E staining of the muscles 28 days after injury, the final outcomes in all groups were similar.

To investigate how the administration of NS-398 promotes fibrosis, we used immunohistochemistry to examine the expression of TGF-ß1 in the area filled with fibrous tissue. TGF-ß1 plays a key role in initiating the fibrosis cascade.36,37 Its expression level correlates with the proliferation of fibroblasts and their production of collagen type I, the major component of fibrous tissue.38 The increased TGF-ß1 expression observed in the NS-398-treated groups relative to that observed in the control group suggests that NS-398 may slow muscle regeneration by increasing TGF-ß1 production, which in turn increases fibrosis.

Results from previous studies have suggested that the satellite cell population, as traditionally defined, probably contains committed myogenic progenitor cells and uncommitted stem cells.27 Muscle-derived stem cells are pluripotent cells that, when placed under proper conditions both in vivo and in vitro, can differentiate into hematopoietic,39 chondrogenic,40 osteoblastic,26,41 and myogenic26,27 lineages. After muscle injuries, growth factors such as TGF-ß1 stimulate muscle-derived stem cells to differentiate into fibroblast-like cells.42,43 The differentiation of such muscle-derived cells may explain the fibrosis that occurred in our experiments and occurs in clinical cases of muscle injury.

Our previous studies have shown that an early wave of TGF-ß1 expression occurs during the first 3 days after injury.43 To investigate the role of this early TGF-ß1 expression, we analyzed the expression of molecules related to TGF-ß1, including myostatin. Myostatin is a member of the TGF super family and plays a key role in regulating skeletal muscle growth.44 Mice lacking myostatin exhibit a dramatic and widespread increase in skeletal muscle mass due to myofiber hypertrophy and hyperplasia.44 The higher expression of myostatin observed in the NS-398-treated muscles compared with the control muscles indicates that NS-398 may negatively affect muscle regeneration by up-regulating myostatin. It is intriguing that we observed highly co-localized expression of TGF-ß1 and myostatin 3 days after injury. Furthermore, the chronological expression pattern that we observed suggests that the expression of TGF-ß1 preceded the expression of myostatin. Although characterization of the exact interrelationship of these two molecules will require additional investigation, some reports already have shown that TGF-ß1 regulates bone morphogenetic protein-1, a molecule necessary for myostatin activation.45,46

Inflammatory responses, including the infiltration and migration of neutrophils and macrophages, are integral to the healing process. Some researchers have hypothesized that neutrophils exacerbate muscle injuries by releasing reactive oxygen intermediates, including peroxides, hypochlorite, and superoxide.47-51 During muscle healing, different populations of macrophages perform different roles, including phagocytosis and the release of growth factors.52,53

CD-11b (also called Mac-1) is a neutrophil surface antigen.24 After activation on neutrophils, its expression is rapidly up-regulated.23 CD-11b also reportedly is present on other immune cells, including lymphocytes and monocytes.54,55 F4/80 antigen is a 160-kd glycoprotein that is expressed exclusively by most murine macrophages. Although this antigen is expressed on macrophages from different sites, it is not expressed on lymphocytes or polymorphonuclear cells.22,56 Our study involved the combined use of these two markers to identify and distinguish between the neutrophil and macrophage populations present within muscles after injury. The administration of NS-398 reduced the relative percentages of all inflammatory cell populations within the injured muscle in a time-dependent manner. This finding demonstrates the anti-inflammatory effects of NS-398. This attenuation of the inflammatory response may explain why some researchers have reported that NSAIDs appear to promote muscle healing at early time points after injury.13,14 On the other hand, reports have indicated that macrophages play an important role in the muscle healing process. By phagocytosing damaged tissue, macrophages help to remove the debris that can impede muscle regeneration. These cells also stimulate the proliferation of activated satellite cells by releasing growth factors and cytokines and by direct contact with them.57-60 Recent study results also suggest that macrophages may fuse with myofibers directly to promote regeneration.61 Our study showed that the administration of NS-398 reduced the infiltration of macrophages, especially at the early time point of 24 hours after injury. By reducing the number of infiltrating macrophages, the administration of NS-398 may have impeded the proliferation of satellite cells and reduced the secretion of some necessary growth factors and cytokines. We believe this inhibition of the immune response was at least partially responsible for the delays in MHC isoform transition and myofiber regeneration during the healing process.

Our study provides evidence that NS-398 has an effect on the proliferation and maturation of differentiated myogenic precursor cells in vitro. We also found that the in vivo administration of NS-398 delays muscle healing by interfering with the normal inflammatory response and the maturation of regenerating myofibers and by increasing fibrosis, possibly by up-regulating TGF-ß1 and myostatin. In light of the results from our study and other reports,17,30 the use of Cox-2-specific inhibitors to treat skeletal muscle injuries appears to warrant caution. When confronted with cases that require the clinical use of such inhibitors to treat skeletal muscle injuries, clinicians probably should consider a reduced dosage or a shorter duration of administration.


   Acknowledgements
 
We thank Alison Logan for her technical support with the flow cytometry experiments, Masamitsu Sakamoto and Hany Bedair for their assistance with animal surgeries, Jin Hong Zhu for her help with immunohistochemistry, Victor Prisk for his helpful input, and Ryan Sauder for his excellent editorial assistance during manuscript preparation.


   Footnotes
 
Address reprint requests to Johnny Huard, Ph.D., Growth and Development Laboratory, Children’s Hospital of Pittsburgh, 4100 Rangos Research Center, 3460 Fifth Ave., Pittsburgh, PA 15213-2583. E-mail: jhuard{at}pitt.edu

Supported by the National Institutes of Health (project grant 1 R01 AR 47973 to J.H. and research facility improvement program grant C06 RR-14489 from the National Center for Research Resources), the William F. and Jean W. Donaldson Chair at Children’s Hospital of Pittsburgh, and the Henry J. Mankin Endowed Chair for Orthopaedic Research at the University of Pittsburgh.

Accepted for publication June 15, 2005.


   References
Top
 Abstract
 Materials and Methods
 Results
 Discussion
 References
 

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