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MUSCLE CELL BIOLOGY AND CELL MOTILITY
Departments of 1Surgery, 2Epidemiology and Biostatistics, 3Pathology, 4Periodontics and 5Medicine, 6Sam and Ann Barshop Institute for Longevity and Aging Studies, University of Texas Health Science Center, San Antonio; 7Protein Design Labs, Fremont, California; and 8The South Texas Veterans Health Care System, San Antonio, Texas
Submitted 5 April 2006 ; accepted in final form 27 September 2006
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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adipocyte; macrophage; MyoD; myogenic progenitor cell; neutrophil; signal transducers of and activators of transcription-3
Of growing interest is the parallel and possibly direct contribution of this chemokine system to the progression of cellular events in skeletal muscle regeneration following ischemic injury. For instance, MCP-1 and CCR2 are expressed in injured skeletal muscle and the recovery of muscle strength is delayed in CCR2–/– mice following injury (64, 65). MCP-1 has also been immunolocalized to endothelial cells and macrophages in the ischemic muscles of C57Bl/6J mice (53). Furthermore, in unrelated in vitro studies, isolated macrophages enhanced proliferation, while delaying differentiation of myogenic progenitor cells (MPC) (38). MPC, also known as satellite cells, are normally quiescent, but upon injury become activated and proliferate (21). Activated satellite cells are referred to as myoblasts. Independently, we have observed impaired perfusion and altered muscle regeneration in MCP-1–/– mice following femoral artery excision (FAE; 52a). In combination, these findings suggest that the recruitment and activation of macrophages via MCP-1/CCR2 forms the foundation for normal muscle regeneration. We hypothesize that recruitment and activation of macrophages via MCP-1/CCR2 during early inflammatory events can impact muscle regeneration at the level of the MPC responsible for replacing damaged muscle fibers. Because MPC are activated immediately following muscle injury (21), the relationship between CCR2, macrophages, and regeneration is an intriguing one. However, whether CCR2-mediated signaling acts directly or indirectly to regulate MPC during regeneration remains speculative.
Given the diversity of cells that are involved in responses following muscle injury, a complex repertoire of signaling mechanisms is required to regulate dynamic reparative processes. For example, in regenerating muscle, the expression of the myogenic regulatory transcription factor, MyoD, promotes proliferation and myogenic differentiation of MPC (49). While in vitro MPC are multipotent and are capable of transdifferentiation into other cell types, including adipocytes (62), osteoblasts (57), and endothelial cells (55), the ability of MPC to transdifferentiate in vivo is not well established. These alternative cellular outcomes are the result of modified transcriptional steps critical to directing terminal differentiation. In addition to MyoD, other molecular signaling factors are involved in MPC regulation. One such factor, signal transducers of and activators of transcription-3 (STAT3) was associated with MPC proliferation in regenerating rat skeletal muscle (25, 26) and cultured C2C12 myoblasts (19). STAT3 is also involved in inflammatory cell activation and function (32). Thus complex signaling cascades are capable of modulating multiple types of cellular responses in the healing microenvironment following ischemic injury.
The present study examined the role of CCR2 during inflammation, restoration of perfusion, and skeletal muscle regeneration following FAE. Despite similar levels of ischemic muscle injury and restoration of perfusion compared with WT mice, CCR2–/– mice had increased neutrophil infiltration, decreased macrophage accumulation, impairments in muscle regeneration, and increased intermuscular adipocyte deposition in areas of regeneration. In parallel, MCP-1 levels were increased in the ischemic muscle of CCR2–/– mice and persisted for an extended time period. Though no significant differences in the MyoD and STAT3 activities were observed at any individual time point post-FAE between WT and CCR2–/– animals, overall transcription factor activity was increased in CCR2–/– mice. These findings suggest that alterations in inflammatory cell recruitment modulate signaling events via the MCP-1 and CCR2 axis, and may impinge on MPC fate and skeletal muscle regeneration.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Mouse FAE model and laser Doppler imaging. Mice were anesthetized and the femoral artery excised as previously described (54). Briefly, the right femoral artery was removed from the inguinal ligament to just proximal to the bifurcation of the popliteal and saphena arteries; the femoral vein and nerve were preserved. A sham surgery was performed on the left leg, where an incision was made and then closed. For vascular perfusion studies, laser Doppler imaging (LDI) (Moor Instruments, Wilmington, DE) was sequentially performed on each animal at all time points to determine perfusion to both hind limbs as previously described (54).
Tissue weights and lysate preparation. Mice were euthanized at various time points following FAE and sham surgery (days 1, 3, 7, and 14; n = 5/time point) and all of the muscles of the hindlimb were removed, weighed, and immediately utilized to prepare tissue lysates as described previously in detail (53). For Western blots, lysates were prepared as previously described (53) using a modified buffer [10 mM Tris, pH 7.4, containing 100 mM NaCl, 1% Triton X-100, 0.1% SDS, 0.5% sodium deoxycholate, 5.5 mM EDTA, 1 mM EGTA, 20 mM sodium pyrophosphate, 1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride, 65 µm bestatin hydrochloride, 7 µm trans-epoxysuccinyl-L-leucylamido-(4-guanidino) butane, 11 µm leupeptin, 0.15 µm aprotinin, and 1 mM PMSF]. "Calf" was designated as all muscles from the knee to the ankle and "thigh" was designated as all muscles above the knee to the inguinal ligament. Similar samples were collected from animals not subjected to surgery and served as negative controls (baseline).
Measurement of protein and MCP-1 levels and activity of myeloperoxidase and lactate dehydrogenase. Protein in tissue lysates was determined with bicinchoninic protein assay (Pierce Biotechnology, Rockford, IL) (53). Myeloperoxidase (MPO) and lactate dehydrogenase (LDH) activity in tissue lysates was measured and adjusted for the amount of protein in the tissue lysates (53). MCP-1 levels were assessed by an ELISA from Biosource (Camarillo, CA) with a slight modification of manufacturer's specifications as previously described (53). Data for weight, protein/weight, MPO, LDH, and MCP-1 for WT animals have been previously published (53) and are included here for comparison to the CCR2–/– mice.
Determination of MyoD and STAT3 activity and expression. MyoD and STAT3 activity assays were performed in tissue lysates using TransAM kits (10, 36) from Active Motif (Carlsbad, CA). This ELISA-like assay measures activated transcription factor binding to an oligonucleotide consensus binding sequence (5'-CACCTG-3' or 5'-TTCCCGGAA-3' for MyoD and STAT3, respectively) on a microtiter plate; the active form of the transcription factors specifically binds to the consensus sequence bound to the microtiter well is then identified via an enzyme-linked antibody detection system with colorimetric readout (45). For this assay, tissue lysates were thawed on ice and centrifuged at 16,100 g for 10 min at 4°C. All samples were initially diluted in tissue lysate buffer to a protein concentration of 2.0 mg/ml, followed by further serial dilutions. For MyoD determinations, 5 and 10 µg of protein from each lysate were assayed. For STAT3 assays, 5 µg of protein from each lysate were assayed. Nuclear extracts (Active Motif) of C2C12 or HepG2 cells were diluted in lysate buffer and used in every assay as a positive control for MyoD or STAT3 activity, respectively. Lysate buffer alone was used to determine background absorbance at 450 nm; the outer diameter values at 450 nm (OD450) of all unknown samples were corrected for background absorbance and values reported as OD450 after adjustment for mg of protein in the assay.
MyoD and STAT3 expression were evaluated in representative samples for each mouse strain and post-FAE time point by Western blot analyses. For Western blot analyses, PAGE-SDS was conducted as described by Laemmli and Favre (30) using 12.5% polyacrylamide/SDS resolving gels (Bio-Rad, Hercules, CA). Proteins were transferred to Immobilon (Millipore, Billerica, MA) polyvinylidene fluoride membrane. Immunoblot analyses were conducted using rabbit polyclonal STAT3 and rabbit monoclonal phospho-STAT3 (Tyr705) antibodies (Cell Signaling, Danvers, MA) and rabbit polyclonal MyoD (C-18) (Santa Cruz Biotechnology, Santa Cruz, CA). Immunoreactive proteins were detected with AlexaFluor 680 goat anti-rabbit secondary antibody (Invitrogen, Carlsbad, CA) using an infrared imaging system (Odyssey; LiCor, Lincoln, NE).
Measurement of tissue macrophages and neutrophils by fluorescence-activated cell sorting. Cells from FAE and sham surgery calf muscles were isolated as previously described by Rando and Blau (43). Briefly, all the muscles between the knee and ankle were removed from WT and CCR2–/– mice 3 days post-FAE. Fat was carefully removed from the tissues. Cells were enzymatically dissociated by incubation in a solution containing 1% collagenase type II and 2.4 U/ml dispase (both from Invitrogen) in phosphate-buffered saline (PBS) supplemented with CaCl to a final concentration of 2.5 mM at 37°C for 90 min. Muscle tissues were triturated with fire-polished Pasteur pipettes every 15 min and then passed through 40 µm cell strainer (BD Biosciences, San Jose, CA). The filtrate was centrifuged at 350 g to harvest the dissociated cells, which were further quantitatively analyzed for macrophages and neutrophils with a fluorescence-activated cell sorter (FACS).
Macrophages and neutrophils were analyzed by a modified procedure for muscle-associated cells as previously described by Lagasse and Weissman (31). Neutrophils were defined as CD11b+/Gr-1+, and macrophages were defined as CD11b+/Gr-1–. Single cell suspensions were treated with mAb 2.4G2 (1 µg/106 cells, BD Biosciences) for 20 min to block Fc
II/III receptors and incubated with conjugated antibodies at optimal concentration at 4°C for 25 min. The conjugated monoclonal antibodies were as follows (all from BD Biosciences): APC-anti-mouse Gr-1 (Clone RB6–8C5) at 0.2 µg/106 cells, PE-anti-mouse CD11b at 0.1 µg/106 cells (Clone, M1/70), as well as the corresponding isotype control antibodies. Cell analysis was performed on a FACSAria (BD Biosciences) with data analysis accomplished with FACSDiva software (BD Biosciences).
Histology, immunohistochemistry, oil red O, capillary counts, and histomorphometry. Mice were euthanized at 1, 3, 7, 14, 21, or 28 days (4–8 mice per time point) following FAE and sham surgery (right and left leg, respectively), and hindlimb tissues were collected for histology and immunohistochemistry with and without perfusion fixation as previously described (53). Routine, indirect immunohistochemical procedures were used for the localization of monocytes/macrophages (F4/80 and mac3) or neutrophils (Ly-6) in deparaffinized sections. Nonspecific binding of antibodies was blocked by treatment of sections (30 min) with 1% human serum albumin in PBS. Primary rat monoclonal antibodies were directed against mouse F4/80 (Serotec, Raleigh, NC), mouse mac3 (BD Biosciences), and mouse Ly-6 (BD Biosciences). Antibodies were diluted 1:500, 1:200, and 1:10, respectively, in 1% human serum albumin in PBS. A biotinylated secondary antibody (mouse absorbed rabbit anti-rat IgG used at a dilution of 1:200) and streptavidin-horseradish peroxidase were obtained from Vector Laboratories (Burlingame, CA). After enzymatic development in diaminobenzidine tetrahydrochloride and hydrogen peroxide, sections were counterstained with hematoxylin. MyoD and F4/80 immunolocalization on frozen sections was performed as previously described (53) using a primary antibody dilution of 1:80 for MyoD. Oil red O staining was performed on 4 µm cross-sections of frozen muscle using a kit from Poly Scientific (Bay Shore, NY) per manufacturer protocol. The biotinylated lectin Griffonia (Bandeiraea) Simplicifolia lectin I (Vector Laboratories) at 1:50 dilution was used to identify endothelial cells on deparaffinized cross-sections. Endogenous peroxidase activity was blocked by incubation in 3% H2O2, and nonspecific binding was blocked by treatment of sections with 1.67% horse serum in PBS. After incubation with lectin, sections were sequentially incubated with horseradish peroxidase-streptavidin, followed by diaminobenzidine tetrahydrochloride, and counterstained with hematoxylin and eosin (H&E).
Histology and immunohistochemistry images were captured with an Eclipse microscope (model E600; Nikon, Melville, NY) equipped with a high-resolution digital camera (Nikon DXM 1200) connected to a personal computer equipped with Act-1 (Nikon) software for image capture and archiving. Capillary counts and morphometric analyses were performed on images captured using a Nikon Eclipse TE2000-U microscope equipped with a high-resolution digital camera (Nikon DXM 1200F) interfaced with a personal computer equipped with Metamorph (Nikon) software.
For morphometric analyses, anterior compartment specimens were removed en bloc at 21 days post-FAE (n = 5 mice/strain), 2- to 3-µm-thick cross-sections were obtained through the midportion of the muscle and stained with H&E or Masson's trichrome. Additional sections were processed for capillary density. Within each section, eight nonoverlapping areas of the tibialis anterior (TA) muscle were randomly selected and digitally captured (x20 magnification); areas containing large blood vessels or fibrous tissue bands between muscle bundles were excluded. The percentage of the total cross-sectional area of the TA covered by the eight digital images was 35–50%. Preliminary studies demonstrated no significant difference in fiber cross-sectional area between H&E and trichrome-stained sections (data not shown). For fiber size and percent of intermuscular fat, two different microscopic sections separated by at least 100 µm were used for morphometric analyses for both the FAE and sham surgery muscle; data were similar at both levels. Results from replicate sections were averaged for a given animal.
In each of the digitally captured images, intermuscular adipocytes (i.e., between and among individual muscle fibers in a given muscle bundle) were manually outlined and used to calculate the total area of fat that was then divided by the total area of the image (0.278 µm2) to calculate the percent fat. The average percent fat area of eight images for each TA section was then determined. The average percentage of fat among replicate TA sections for each animal was calculated for subsequent use in comparisons of results for each group of animals.
The average cross-sectional area (µm2) of individual muscle fibers for a given animal was determined after manually outlining individual muscle fibers in each of the digitally captured images for a given TA; fibers that were only partially included within the images were excluded. In the TA of FAE limbs, only regenerated fibers [identified by the presence of a centrally located nucleus (6)] were measured, whereas only fibers with peripherally located nuclei (i.e., mature, nonregenerated fibers) were measured in the TA of the sham surgery limbs for both WT and CCR2–/– mice.
Capillary counts were performed on both the TA of FAE and sham surgery specimen using the eight fields described above. Images were digitally captured using phase contrast microscopy at x20 to enhance the identification of cell borders and lectin staining. The number of capillaries and muscle fibers in each image were counted using NIS-Elements Software (Nikon). Only capillaries associated with muscle fibers were counted and expressed as capillaries/muscle fiber. In addition, areas of fat were manually outlined and subtracted from the total area and capillary density was also expressed as capillaries/mm2.
Data analysis. SAS software (SAS, Cary, NC) was used for all statistical analyses. Results from corresponding time points of each group were averaged and used to calculate descriptive statistics (means ± SE).
Sequentially derived LDI ratios were analyzed by two different methods. First, to determine whether LDI ratios within a given group had returned to preoperative (baseline) levels, a Bonferroni-corrected multiple-comparisons procedure was used. Data collected preoperatively were subtracted from each of the other time points to produce a dataset of comparable LDI ratio differences. Second, to determine whether there were differences between CCR2–/– and WT mice at each time point, an analysis of variance incorporating repeated measures across time was used with a Bonferroni correction for multiple comparisons. Within animals, time correlations and heterogeneity of the variances across time were modeled using the ANTE (1) covariance structure. Animal variance was considered a random effect.
Protein/weight, weight, enzyme activity, transcription factors, and MCP-1 data were analyzed by a Dunnett-corrected multiple-comparisons procedure utilizing a two-way analysis of variance of least-square means to determine whether significant differences existed at different time points (1, 3, 7, and 14 days post-FAE) compared with baseline values. For transcription factors, MCP-1 and MPO, results were log transformed to adjust for unequal variances. For lysates samples with MCP-1 below the level of detection in the ELISA (<78 pg/ml), a value of 78/2 pg/ml was assigned to these samples (20) and this value was corrected for the protein in each specimen.
FACS data for neutrophil and macrophage quantitation were analyzed by paired (within strain) and unpaired (between strains) t-tests.
Histomorphometry data for percent fat and fiber cross-sectional area were analyzed as follows. First, a repeated measures linear model was used to determine that there were no significant interactions between the two different sections of the TA muscle for a given animal. Replicate data for all images and levels were then averaged to generate a single number for each limb for each mouse, i.e., fiber cross-sectional area was determined for both FAE and the contralateral limb whereas percent fat was only calculated for the FAE limb. Paired t-tests were used to assess the significance of mean differences in fiber cross-sectional area between FAE vs sham limbs within each strain. One-way ANOVA were used to assess the significance of mean differences between the limbs of CCR2–/– and WT mice for FAE (fiber cross-sectional area and percent fat) or sham (fiber cross-sectional area). Analysis of capillaries/fiber and capillaries/mm2 were performed in a similar manner, except only one level of the TA was used to generate a single number for each limb. All statistical testing was two sided with a significance level of 5%.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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50% of that in the normal, uninjured muscle (P 
0.001). LDH activity in postischemic calf muscle of WT mice remained significantly decreased at day 7 (P = 0.002) but WT LDH activity were similar to baseline by day 14 post-FAE. In contrast, LDH activity in CCR2–/– mice was significantly decreased at day 3 (P = 0.003) and remained significantly decreased compared with baseline at both days 7 and 14 (P 0.001). Furthermore, LDH activity was significantly decreased in CCR2–/– compared with WT mice at day 14 (P = 0.004). This enzymatic measure of tissue injury and repair paralleled the histological evaluation and extent of muscle regeneration described below.
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1 in both mouse strains (Fig. 2), i.e., perfusion was comparable in the right and left legs. Immediately following FAE, perfusion dropped to 15% of the perfusion levels observed in the contralateral, sham surgery leg. Thereafter, restoration of perfusion was similar in both mouse strains, increasing gradually throughout days 3-28 following FAE. In both WT and CCR2–/– mice, perfusion was not fully restored and remained at 75% of preoperative perfusion levels at day 28 (P 0.001 compared with baseline). There were no significant differences in the perfusion ratios between WT and CCR2–/– mice at any time point.
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The calf muscle of CCR2–/– mice had a similar histological appearance to WT mice at day 1 post-FAE, except neutrophils appeared to be more numerous. However, the injured muscle of CCR2–/– mice differed from that of WT mice at all subsequent times. CCR2–/– mice had a minimal mononuclear cell infiltrate at day 3 (Fig. 3B) and very few F4/80+ cells were identified in areas of necrotic muscle fibers (Fig. 3D), indicating that CCR2–/– mice have fewer intramuscular macrophages compared with WT mice. At day 7 post-FAE in CCR2–/– mice, an interstitial inflammatory infiltrate was prevalent in association with extensive residual necrotic myofibers (Fig. 3F). Necrotic myofibers persisted and were widespread at day 14 in CCR2–/– animals (Fig. 4, C and D), whereas muscle regeneration was essentially complete in WT mice at this time (Fig. 4A). Necrosis was still present in CCR2–/– mice at 21 days post-FAE (Fig. 4E). Thus CCR2–/– mice had decreased and delayed macrophage accumulation into the injured muscle in association with residual tissue necrosis. In addition, by day 14 post-FAE in CCR2–/– mice, there was a considerable accumulation of adipocytes between regenerated fibers (Fig. 4, C and D) while WT mice exhibited only small, scattered, foci of intermuscular adipocytes (Fig. 4B). The lipid content within the cytoplasmic vacuoles of these cells was confirmed by oil red O staining (Fig. 5, A and B). Interestingly, at 7 days post-FAE, large, vacuolated cells were prevalent among necrotic myofibers of CCR2–/– mice (Fig. 3G); these cells may represent the early precursor cells of adipocytes that prevailed at 14 and 21 days post-FAE (24). Note that the histological progression of muscle regeneration in both groups closely paralleled LDH measurements of tissue injury and regeneration (Fig. 1).
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Intermuscular fat was not identified in the sham surgery TA muscles of either mouse strain on post-operative day 21. However, in the TA of FAE limbs, intermuscular adipocytes were present in areas of muscle regeneration (Figs. 3H, 4B, and 4E). Compared with WT mice, CCR2–/– mice had significantly increased intermuscular fat within the TA (18.2 ± 3.6% vs. 3.3 ± 0.2% for CCR2–/– and WT mice, respectively, P = 0.003) (Fig. 6A).
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0.001) smaller than myofibers of the corresponding contralateral limb.
Finally, capillary density was measured in the FAE and sham 21-day TA muscles of CCR2–/– and WT mice (Fig. 6, C and D) and expressed as either capillaries/muscle fiber or capillaries/mm2. In the FAE TA muscles, only areas of regeneration, identified by centrally located nuclei in the muscle fibers, were used. For these measurements, only capillaries associated with muscle fibers were counted and areas of adipocyte accumulation or necrosis were excluded so that the significant increased adipocyte accumulation in the CCR2–/– regenerating muscle would not influence capillary density measurements. There were no significant differences in capillaries/fiber (Fig. 6C) or capillaries/mm2 (Fig. 6D) in the contralateral sham TA of CCR2–/– and WT mice. However, in the FAE TA, CCR2–/– mice exhibited significantly (P 0.001) decreased capillaries/fiber (0.97 ± 0.03) compared with WT mice (1.66 ± 0.04; Fig. 6C). Furthermore, compared with the sham TA, CCR2–/– mice had a significant decrease in capillaries/fiber in the FAE TA (P = 0.001), whereas the FAE TA of WT mice had similar capillaries/fiber.
Interestingly, the significant differences in capillary density between CCR2–/– and WT mice were not exhibited when capillary density was expressed in relation to the area of the regenerated muscle. Thus, between strains, similar numbers of capillaries/mm2 were present in regenerated and normal muscle (Fig. 6D). Therefore, the significant decrease in capillaries/fiber in the FAE TA of CCR2–/– animals reflected the decreased size of the regenerated muscle fibers (Fig. 6B) rather than differences in the number of capillaries at the 21-day time point.
FAE-induced changes in tissue weight, protein, MPO activity, and macrophage recruitment.
Tissue weight was measured at various time points following FAE as an indication of edema. To adjust for small differences in animal size, thigh and calf FAE specimen weights were normalized to the contralateral, sham surgery leg. For thigh muscles, there were no significant changes in the relationship between right and left weight ratios compared with baseline at any time point following FAE for either strain of mice (data not shown). In contrast, the relative weight of the ischemic calf of WT mice was significantly increased at day 3 (P 0.001) and returned to baseline at day 7 (Fig. 7A). In comparison, the calf weight ratio of CCR2–/– was significantly elevated at day 1 and remained elevated through day 7 before returning to baseline at day 14. CCR2–/– calf weight ratios were significantly elevated over WT at days 1 and 7 (P 0.01). The protein/weight ratios in calf tissue lysates had a similar but inverse pattern to tissue weight except for day 14 post-CT in the CCR2–/– mice, where the protein/weight ratio remained decreased from baseline secondary to decreased protein levels in the lysates (Fig. 7B). Thus the weight and protein/weight ratios during post-CT days 1–7 were consistent with edema in the CCR2–/– mice while the decreased protein in the 14-day post-CT lysates in the CCR2–/– mice suggested impaired regeneration; both of these observations were consistent with the histological features of the postischemic muscle tissues.
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0.002) post- FAE (Fig. 7C). In comparison, MPO activity in the calf muscle of CCR2–/– mice was elevated over baseline in an accelerated pattern, i.e., at day 1 post-FAE. At this time, MPO activity in the calf muscle of CCR2–/– mice was significantly elevated compared with WT mice (P 0.001), indicating an increased presence of neutrophils.
Further quantitation of tissue neutrophils and macrophages (cells x 106/g tissue) was accomplished by FACS analysis at day 3 post-FAE (Fig. 8). Neutrophils (P 
0.04) and macrophages (P 0.03) in both WT and CCR2–/– mice were elevated in the FAE vs the sham calf tissues. In the sham surgery calves, WT and CCR2–/– mice had similar numbers of neutrophils (0.15 ± 0.06 vs. 0.21 ± 0.19 cells x 106/g tissue, WT and CCR2–/–, respectively). In contrast, the CCR2–/– sham calves had significantly less macrophage recruitment than WT mice (0.07 ± 0.02 vs. 0.01 ± 0.00 cells x 106/g tissue for WT and CCR2–/– mice, respectively, P = 0.03). In the ischemic, FAE calves, neutrophil recruitment was similar at day 3 post-FAE (Fig. 8C) and this was consistent with the MPO data at day 3 post-FAE (Fig. 7C), described above. On the other hand, there was a severe impairment in macrophage recruitment in CCR2–/– compared with WT mice (Fig. 8C).
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0.002), which had returned to baseline levels at day 7.
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0.01) and significantly elevated at days 3 and 7 compared with WT mice (P 0.004; Fig. 9B).
To examine the molecular regulation of muscle regeneration and other cellular responses, MyoD and STAT3 transcription factor activities were measured in calf muscle tissue lysates following FAE. For both mouse strains, MyoD activity progressively increased after ischemic injury and decreased as the muscle regenerated (Fig. 10A). In WT mice, MyoD activity was significantly higher than baseline at days 3, 7, and 14 (P 0.02) with maximal expression at days 3 and 7. In CCR2–/– mice, MyoD activity was significantly elevated over baseline at all time points post-FAE (P 0.001). STAT3 activity in the post-ischemic calf muscle of WT mice was only increased above baseline at day 3 (P = 0.05; Fig. 10B), whereas in CCR2–/– mice, STAT3 activity was significantly elevated over baseline at days 7 and 14 post-FAE (P 0.05). Although there was a trend toward increased MyoD and STAT3 activities in CCR2–/– mice, there were no significant differences in the activities of these transcription factors between WT and CCR2–/– mice at any individual time point post-FAE. Nevertheless, when results from all time points post-FAE were taken altogether, there were significant differences between WT and CCR2–/– mice for both MyoD and STAT3 (P = 0.02 and P = 0.05, respectively). Western blot analyses for MyoD, STAT3, and phosphorylated STAT3 showed a pattern of expression consistent with that observed by transcription factor activity assay (Fig. 10C).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Recently, Reichel et al. (44) reported that mice deficient in certain CCRs, including CCR2, exhibited decreased neutrophil infiltration following ischemia. In contrast, we observed increased neutrophil infiltration into the ischemic calf muscle of CCR2–/– mice following FAE. The differences between these studies may arise from the assessment, theirs being made in an ischemia-reperfusion model immediately following reperfusion. In addition to using the FAE model, the current study used myeloperoxidase measurements to quantify neutrophils present in ischemic muscle. Other investigators have described increased neutrophil accumulation in CCR2–/– mice using various injury models (23, 29). Moreover, the antagonism of MCP-1 has resulted in delayed neutrophil clearance due to decreased numbers of macrophages and the reduced ability of macrophages to phagocytize apoptotic neutrophils (1, 35). Thus, inadequate macrophage recruitment and activation in the present study may underlie the increased accumulation of neutrophils.
Previously, Tang et al. (60) reported reduced macrophage recruitment following hindlimb ischemia in CCR2–/– mice. In that study, macrophages in ischemic calf muscle were decreased, but in the thigh, an area corresponding to the site of active arteriogenesis, equivalent amounts of macrophages and increased levels of MCP-1 mRNA were observed. Tang et al. (60) also described similar restoration of perfusion between CCR2–/– and WT mice following ischemia. In contrast, another study described decreased restoration of perfusion in CCR2–/– mice (17). The earlier findings by Tang et al. taken in conjunction with the findings described here pose a confounding contradiction: that despite an apparent normal restoration of perfusion, muscle was unable to regenerate properly in CCR2–/– mice. This indicates that events other than perfusion (i.e., inflammation, satellite cell activation, etc.) account for the delayed and impaired muscle regeneration observed in CCR2–/– mice. With regard to MCP-1-related arteriogenesis, the present study demonstrated that MCP-1 protein levels in thigh were negligible. Rather, elevated levels of MCP-1 protein observed in the calf suggest a more crucial function for MCP-1 and CCR2 at the site of ischemic tissue injury and regeneration.
Skeletal muscle regeneration is primarily dependent upon perfusion, inflammation, and satellite cells, the multipotent progenitor cells that reside in skeletal muscle (14). Although there appeared to be no major differences in restoration of perfusion in WT and CCR2–/– mice herein, it is still possible that the impairment in skeletal muscle regeneration was secondary to a more severe or prolonged ischemic insult. Alternatively, the impairment of skeletal muscle regeneration could be a primary defect, i.e., satellite cells may require CCR2 for efficient proliferation and differentiation. It is also possible that modifications in the nature or magnitude of the inflammatory infiltrate, especially macrophages, including the presence or absence of mediators derived following cell activation, lead to impairments in skeletal muscle regeneration.
Previous studies (33, 63) of ischemic muscle damage have demonstrated that inflammation is important for the removal of necrotic tissue and influences many aspects of regenerative processes. Consistent with these observations, decreased macrophage recruitment in CCR2–/– mice in the present study was associated with impaired muscle regeneration and increased intermuscular fat deposition. Similar impairments in muscle regeneration were provided in a recent study where macrophage influx was reduced by liposomal clodronate injections (56). It is conceivable that with reduced macrophage recruitment in both of these studies, inadequate macrophage-dependent growth factors in the regenerating environment lead to impaired and altered muscle regeneration. Indeed, macrophages secrete factors that modulate both physiological responses and differentiation of nearby cells (58). Thus, alterations in inflammation may influence not only the removal of damaged skeletal muscle, but also MPC proliferation and/or differentiation.
MCP-1 is just one of many factors secreted by macrophages (23). This chemokine is also generated by muscle (46). We observed significantly elevated levels of MCP-1 protein in CCR2–/– mice; however, the physiological consequence of increased MCP-1 remains unknown. At concentrations exceeding normal physiological levels, MCP-1 may interact with alternative chemokine receptors that are not typically favored in the presence of CCR2. In this regard, higher concentrations of MCP-1 are required to activate CCR3 while subnanomolar concentrations are sufficient for CCR2 activation (41). Conversely, CCR2 can bind other chemokines, including MCP-2–5 (27, 47) and eotaxin-1 (41). Normal cellular responses to these other chemokines could also be inhibited in the absence of CCR2. Likewise, potential interactions between excessive amounts of MCP-1 and other chemokine receptors may have functions redundant to the normal MCP-1/CCR2 axis but may also direct an array of unforeseen events.
The mechanism of MCP-1 signal transduction via the CCR2 receptor is not well established. It has been suggested that MCP-1 may trigger activation of the Janus kinase 2 (JAK2)/STAT3 pathway and tyrosine phosphorylation of the CCR2 receptor in macrophages (4). However, based on the current study, it appears that MCP-1 does not directly activate JAK2 via the CCR2 receptor. Further studies are needed to clarify the role of the MCP-1/CCR2 axis in STAT3 signaling.
The effect of STAT3 signaling during regeneration, particularly in multipotent MPC, is unknown. However, STAT3 activation normally regulates several biological functions in myoblasts including proliferation, differentiation, and survival (25, 26). While STAT3 activation in CCR2–/– mice was not significantly different than WT at specific time points, STAT3 activity in CCR2–/– mice tended to be elevated overall compared with WT. Interestingly, in human adipogenesis, STATs, including STAT3, are upregulated (15). Further evaluation of STAT3 activation in CCR2–/– mice will be important in determining the contribution of this factor to the increased adipocyte accumulation. In addition, the kinetics of STAT3 activation paralleled macrophage recruitment in both WT and CCR2–/– mice. This is interesting because macrophage activation is also associated with the activation of STAT3 (13, 32). Additional studies are required to clarify the role of CCR2 and STAT3 signaling in regenerating muscle following ischemic injury.
In addition to the above, STAT3 induces the expression of the proangiogenic cytokine, VEGF (19) and macrophages can also produce VEGF (58). Since capillary density was similar in CCR2–/– and WT mice in both uninjured and ischemic muscle at day 21 post-FAE and STAT3 activities leading to this point were similar between mice, it seems unlikely that STAT3-dependent angiogenic events were modified in CCR2–/– mice. While capillaries/muscle fiber was decreased in regenerated muscle of CCR2–/– mice, this reflected the smaller size of regenerated fibers. Thus, although it remains to be determined whether earlier angiogenic events are also intact in CCR2–/– mice, the current results do not support the idea that revascularization underlies the observed alterations in skeletal muscle regeneration. Rather, our findings support the hypothesis that the primary defect in skeletal muscle regeneration in CCR2–/– mice involves impairment of myoblast regeneration and/or differentiation.
The results of the present study demonstrate that myoblast regeneration, as indicated by MyoD activity, is independent of CCR2-mediated signaling as the amount and duration of MyoD activity increased similar to WT in the absence of CCR2. Earlier studies reported no difference in MyoD mRNA levels following macrophage depletion (56) or between WT and CCR2–/– mice following freeze injury (64). Current findings of MyoD activity and protein expression are consistent with this previous study. However, we observed an overall trend toward elevated MyoD activity in CCR2–/– mice. Clearly, further studies are required to establish the relative activities of transcription factors that are known to regulate muscle regeneration in CCR2–/– mice compared with WT and thus define the basis for CCR2-dependent impairments in muscle regeneration.
An important question that was not addressed in the current study is what cell types are expressing MyoD and STAT3. Previous studies have documented that MyoD localization is limited to satellite cells, myoblasts, and some muscle fibers (9) in regenerating skeletal muscle; the result herein are consistent with this observation. In contrast to the cell-specific localization of MyoD to muscle, STAT3 is expressed in a variety of cells, including muscle (25), inflammatory cells (34), and vascular endothelial cells (18). A detailed study of STAT3 expression in regenerating rat muscle demonstrated that phosphorylated STAT3 was first detected in the nuclei of activated satellite cells and remained activated in proliferating, but not differentiating, myoblasts. STAT3 nuclear staining was also identified in cells outside of the myofiber and presumably represented inflammatory cells (25). Additional studies are required to identify the cellular sources of STAT3 signaling in regenerating muscle.
The most significant observation in this study was adipocyte formation in regenerating skeletal muscle of CCR2–/– mice. This suggests that locally impaired CCR2-mediated signaling, in association with reduced myogenic and/or regenerative capacity of skeletal muscle progenitor cells following injury, might favor differentiation of mesenchymal progenitor cells toward an adipocytic phenotype (28). This possibility is supported by results from in vitro studies documenting that satellite cells can be redirected, by altered molecular signaling, to accumulate intracellular fat and express adipocyte markers (61, 62). Alternatively, adipocytes from the surrounding areas may migrate into the injured muscle (2) or the adipocytes may be the result of differentiation/transdifferentiation of other progenitor populations, such as bone marrow-derived progenitor cells (22, 52). Regardless of the origin of the adipocytes, our results are consistent with those of a recent report demonstrating adipocyte formation in regenerating muscles of CCR2–/– mice following freeze-induced skeletal muscle injury (64). Together, these findings suggest that the propensity to develop adipocytes in mice with impairments in CCR2 is a generalized reaction to muscle injury, regardless of the basis for muscle tissue damage. Additional evidence for a relationship between altered macrophage recruitment/function and increased intermuscular fat has been observed when macrophages are partially depleted in WT mice by clodronate-containing liposomes (56).
Interestingly, in humans, skeletal muscle regeneration is also impaired with aging (8, 14) and there is an increase in fat within and between the muscle fibers (40, 48). Furthermore, aged mice (12, 39, 59), similar to elderly humans (3, 37), have alterations in macrophage function. Thus CCR2–/– mice may represent a unique animal model for studies designed to elucidate the inflammation-based mechanisms involved in impaired skeletal muscle regeneration that may occur in normal human aging. This is particularly important because in addition to the physiological consequences of impaired muscle regeneration, such as reductions in force-producing muscle fibers, impairments in contraction, and metabolic dysfunction (28), adipose tissue is recognized as having a paracrine/endocrine function with secretion of many factors that influence diverse cell types as well as attract macrophages (5). Therefore, the accumulation of adipocytes could serve as a compensatory mechanism to recruit macrophages for the purpose of enhancing impaired healing and regeneration in this model. Interestingly, the majority of adipose tissue-derived cytokines may emanate from tissue-resident macrophages (5). The possible paracrine effects of intermuscular adipocytes on the surrounding muscle tissue are an active area of ongoing investigation in our laboratory.
In conclusion, the current study supports a role for CCR2-mediated events in skeletal muscle regeneration but not restoration of perfusion. Increases in MCP-1 indicate potentially profound changes in cellular signaling events in CCR2–/– mice in association with enhanced neutrophil and impaired macrophage recruitment. In these animals, there was defective muscle regeneration and increased fat accumulation, despite apparently similar activities of MyoD and STAT3. These events suggest that the functional outcome of the reparative process, including signaling events, is impaired in the absence of CCR2. Importantly, these alterations in neutrophil and macrophage recruitment, and the reparative processes, direct regenerating muscle toward an adipogenic lineage by mechanisms which remain to be established.
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ACKNOWLEDGMENTS |
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Present affiliation for V. Contreras-Shannon: Department of Cellular and Structural Biology, University of Texas Health Science Center, San Antonio, TX.
Present affiliation for S. M. Reyes-Reyna: Department of Medicine, University of Texas Health Science Center, San Antonio, TX.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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