Abstract
Skeletal muscle injury is one of the most common injuries in sports medicine. Our previous study found that macrophage depletion impairs muscle regeneration and that mechano growth factor (MGF) may play an important role in this process. However, whether injection of MGF protects against impaired muscle regeneration after macrophage depletion has not been explored. Therefore, we generated a muscle contusion and macrophage depletion mouse model and injected MGF into the damaged muscle. Comprehensive morphological and genetic analyses were performed on the injured skeletal muscle after macrophage depletion and MGF injection. The results showed that injection of MGF did not exert a protective effect on muscle fiber regeneration; however, it did decrease fibrosis in the contused skeletal muscle after macrophage depletion. Moreover, MGF injection decreased the expression of muscle inflammatory cytokines (TNF-α, IFN-γ, IL-1β, and TGF-β), chemokines (CCL2, CCL5, and CXCR4), oxidative stress factors (gp91phox) and matrix metalloproteinases (MMP-1, MMP-2, MMP-9, MMP-10, and MMP-14). These results suggest that the impairment of skeletal muscle regeneration induced by macrophage depletion could be partly ameliorated by MGF injection and that inflammatory cytokines, oxidative stress factors, chemokines, and MMP may be involved in this process.
Introduction
Skeletal muscle accounts for almost half of the body weight in humans. As a power-generating tissue, it is one of the most important organs in mammals. Skeletal muscle injuries occur very often in humans and are inevitable. Skeletal muscle regeneration following an injury is a complicated process that involves many cells and cytokines ().
Inflammation, especially macrophage inflammation, plays a crucial role in skeletal muscle regeneration (). Experiments have revealed that skeletal muscle injury induces extensive macrophage infiltration at the injury site (). There are two different phenotypes of macrophages, M1 and M2. Classically activated macrophages, typically designated M1 macrophages, are characterized by the expression of proinflammatory cytokines (IL-1β and TNF-α), high expression of the CD68 surface marker and absence of the CD163 marker (). These M1 macrophages infiltrate injury sites following muscle damage and are involved in the phagocytosis of muscle debris (). Then, the M1 macrophages are replaced by “alternatively activated” macrophages (M2 phenotype), which are characterized by high expression of CD163 and anti-inflammatory cytokines (such as IL-10) and contribute to muscle regeneration (; ).
Studies have shown that macrophages not only act as regulators of inflammation but also as secretory cells that produce many cytokines, such as IL-1β, IL-10, IGF-1, HGF, uPA and mechano growth factor (MGF) (; ; ; ; ). Our previous study indicated that MGF expression in the macrophages of overload training rats increased significantly compared to expression levels in the control animals (). A recent study found that macrophages are the main contributors to the upregulation of MGF in injured skeletal muscle (). Furthermore, our previous study found that macrophage depletion significantly decreased the expression of MGF and the regeneration of impaired skeletal muscle (). However, whether injection of MGF protects against impaired muscle regeneration after macrophage depletion has not been explored. We hypothesize that the injection of MGF at least partially protects against impaired skeletal muscle regeneration after macrophage depletion.
To explore this concept, we generated a muscle contusion and macrophage depletion mouse model and injected MGF into the damaged muscle of the mice. The histomorphology, fibrosis and expression of inflammatory cytokines, chemokines, oxidative stress factors, and matrix metalloproteinases (MMPs) were tested in the injured skeletal muscle (Figure 1).
FIGURE 1
Materials and Methods
Animals
One hundred fifty-four C57BL/6J mice [7 weeks old, Shanghai Laboratory Animal Center (SLAC), Co., Ltd.] were used in this study. After acclimatization to the local environment for 1 week, the mice were divided into three groups: skeletal muscle contusion mice (S group), skeletal muscle contusion and macrophage depletion mice (TS group), muscle contusion, macrophage depletion and MGF-treated mice (TSM group). The animals were housed at a constant temperature of 25°C with free access to pellet food and water. The study and all protocols were approved by the Ethics Review Committee for Animal Experimentation of Shanghai University of Sport.
Skeletal Muscle Contusion Model
The mouse gastrocnemius muscle (GM) contusion model was generated according to the methods used in our previous study (; ). Briefly, animals were anesthetized via intraperitoneal injection of 35 mg/kg pentobarbital sodium. The knee was extended, and the ankle was plantar-flexed at 90° after the mice were anesthetized. A 16.8 g (diameter: 15.9 mm) stainless steel ball was dropped from a height of 125 cm through a plastic tube with an interior diameter of 16 mm onto an impactor.
Macrophage Depletion
To generate a macrophage depletion model, clodronate-containing liposomes1 were used as previously described (; ; ). Briefly, 2 mg of clodronate-containing liposomes (CL) or control liposomes was introduced by intrapulmonary injection 3 days before the contusion injury. A total of 0.5 mg of CL or control liposomes was then injected at 0, 3, 6, 9, and 12 days after the muscle was contused. This macrophage depletion protocol was effective and has been used by many researchers (; ; ; ; ).
MGF Treatment
A total of 5 μg/20 μl MGF peptide (GL Biochem, Ltd., Shanghai; Lot No. P140627-25-LR066649) was injected into each GM immediately and at 1 and 2 days after muscle injury. The control mice were injected with PBS-BSA buffer (; ).
Hematoxylin and Eosin (H&E) Staining
After the mice were sacrificed, the GMs were collected and placed in 4% paraformaldehyde before routine paraffin embedding. For the morphometric analysis, 5-μm cross sections of GMs were generated with a microtome (Leica-EG 1160, Germany) and were deparaffinized and stained with H&E. Using the 40× lens objective of a light microscope (Labphot-2, Nikon), image was captured for each muscle section.
Masson’s Trichrome Staining
Masson’s trichrome staining was used to measure the fibrotic tissue area in the injured GMs. After the staining, ImageJ software (ImageJ 1.44, Bethesda, MD, United States) was used to estimate the fibrotic area of the injured skeletal muscle. The sum of the fibrotic area was quantified as previously described ().
Immunofluorescence Staining
Identification of gp91phox in injured muscle was performed by immunofluorescence staining of the GMs. Slides of the GM sections were incubated with a bovine serum albumin (BSA) blocking buffer for 1.5 h and with the gp91phox primary antibody (Abcam, 1:1000) overnight at 4°C. The slides were washed three times in PBS, for 5 min per wash. The sections were incubated with a goat anti-rabbit IgG (Abcam, 1:800) secondary antibody for 60 min, washed three times in PBS, and incubated with 4′,6-diamidino-2-phenylindole (Beyotime Biotech, Co., Ltd.) for 5 min. Using a 20× lens objective (LSM700, Zeiss), an image was captured for each muscle section. Immunofluorescence staining intensity was semiquantitatively analyzed by ImageJ 1.44 image analysis software (National Institutes of Health, Bethesda, MD, United States).
RNA Extraction, cDNA Synthesis, and Real-Time Polymerase Chain Reaction (PCR)
Total RNA was isolated from the GMs using the guanidinium isothiocyanate-CsCl method as previously described (). Total RNA (2 μg) was reverse-transcribed into cDNA (RevertaidTM First Strand cDNA Synthesis Kit, Thermo Scientific). Q-PCR was performed on a StepOnePlus PCR-Cycler (Life Technologies) using the SYBR Green/ROX qPCR Master Mix (Vazyme Biotech, Co., Ltd.) with the following components: 1 μl of cDNA, 7.8 μl of nuclease-free water and 300 nM of each primer. The activation step was performed at 95°C for 10 min, which was followed by 40 cycles of denaturation at 95°C for 15 s and annealing/extension at 60°C for 1 min. The relative mRNA expression was determined using the 2ΔΔCT method (). The primer sequences are listed in Table 1.
Table 1
| Target gene | Forward primer sequences | Reverse primer sequences |
|---|---|---|
| TNF-α | 5′-CTTCTGTCTACTGAACTTCGGG-3′ | 5′-CACTTGGTGGTTTGCTACGAC-3′ |
| INF-γ | 5′-GCTTTGCAGCTCTTCCTCAT-3′ | 5′-GTC ACC ATCCTTTTGCCAGT-3′ |
| IL-1β | 5′-TGACGTTCCCATTAGACAACTG-3′ | 5′-CCGTCTTTCATTACACAGGACA-3′ |
| TGF-β | 5′-TGCGCTTGCAGAGATTAAAA-3′ | 5′-CGTCAAAAGACAGCCACTCA-3′ |
| Col1a1 | 5′-GAGCGGAGAGTACTGGATCG-3′ | 5′-GCTTCTTTTCCTTGGGGTTC-3′ |
| Col3a1 | 5′-GTCCACGAGGTGACAAAGGT-3′ | 5′-GATGCCCACTTGTTCCATCT-3′ |
| MMP-1 | 5′-AGTTGACAGGCTCCGAGAAA-3′ | 5′-CACATCAGGCACTCCACATC-3′ |
| MMP-2 | 5′-ACCCTGGGAGAAGGACAAGT-3′ | 5′-ATCACTGCGACCAGTGTCTG-3′ |
| MMP-9 | 5′-CGTCGTGATCCCCACTTACT-3′ | 5′-AACACACAGGGTTTGCCTTC-3′ |
| MMP-10 | 5′-GAGTGTGGATTCTGCCATTGA-3′ | 5′-TCTCCGTGTTCTCCAACTGC-3′ |
| MMP-14 | 5′-CCTGGCTCATGCCTACTTCC-3′ | 5′-GCACAGCCACCAAGAAGATG-3′ |
| CCL2 | 5′-GCTCAGCCAGATGCAGTTAAC-3′ | 5′-CTCTCTCTTGAGCTTGGTGAC-3′ |
| CCR2 | 5′-GAAAAGCCAACTCCTTCATCAG-3′ | 5′-TCTAAGCACACCACTTCCTCTG-3′ |
| CCL5 | 5′-CATATGGCTCGGACACCA-3′ | 5′-ACACACTTGGCGGTTCCT-3′ |
| MyoD | 5′-GAGCGCATCTCCACAGACAG-3′ | 5′-AAATCGCATTGGGGTTTGAG-3′ |
| myogenin | 5′-CCAGTACATTGAGCGCCTAC-3′ | 5′-ACCGAACTCCAGTGCATTGC-3′ |
| F4/80 | 5′-AACATGCAACCTGCCACAAAC-3′ | 5′-ACAGGATTCGTCCAGGC-3′ |
| CXCR4 | 5′-CAAGGCCCTCAAGACGACAG-3′ | 5′-CCCCCAAAAGGATGAAGGAG-3′ |
| gp91phox | 5′-CCAGTGAAGATGTGTTCAGCT-3′ | 5′-GCACAGCCAGTAGAAGTAGAT-3′ |
| GAPDH | 5′-ACTCCACTCACGGCAAATTC-3′ | 5′-TCTCCATGGTGGTGAAGACA-3′ |
Primers used for qRT-PCR.
Statistical Analysis
The data were analyzed using SPSS 20.0 software for Windows (IBM, United States). The number, diameter and area of regenerating myofibers or fibrotic regions were compared using independent samples t-tests. A factorial analysis of the multi-way ANOVA with the one-way ANOVA was used to analyze the main effects of the S, TS, and TSM groups. Post hoc Scheffe tests were performed when significance was detected by the one-way and multi-way ANOVA tests. All values are expressed as the mean ± SEM, and statistical significance was set at p < 0.05.
Results
Effect of Clodronate-Containing Liposomes on Macrophage Marker in Injured Skeletal Muscle
RT-PCR results indicated that clodronate-containing liposomes significantly decreased the expression of the F4/80 macrophage marker at 1, 3, and 7 days (p < 0.01) post-skeletal muscle injury. In addition, the injection of MGF did not influence expression of the macrophage marker after treatment of injured skeletal muscle with clodronate-containing liposomes (p > 0.05) (Figure 2).
FIGURE 2
MGF Injection Did Not Exert a Protective Effect on Muscle Fiber Regeneration After Macrophage Depletion
H&E staining showed that macrophage depletion significantly decreased the sums of the diameters (7927 ± 991.20 vs. 3318.76 ± 503.19 μm, p < 0.01), numbers (706 ± 63.64 vs. 348 ± 37.15, p < 0.01) and areas (103,461.79 ± 17,145.27 vs. 36,055.28 ± 71,57.94 μm2, p < 0.01) of the regenerating myofibers at 14 days post-injury. However, unexpectedly, MGF injection had no effect on the sums of the diameters, numbers and areas of the regenerating myofibers (p > 0.05) (Figure 3d–f).
FIGURE 3
MGF Injection Decreased the Fibrosis of Contused Skeletal Muscle After Macrophage Depletion
Masson’s trichrome staining was performed to evaluate whether MGF injection improved GM fibrosis after macrophage depletion in injured skeletal muscle. Masson’s trichrome staining showed that fibrosis increased significantly in the GMs in the TS group compared with the fibrosis in the S group at 14 days post-injury (496.34 ± 162.06 vs. 3047.86 ± 857.20 μm2, p < 0.01). However, interestingly, the fibrotic area decreased significantly after MGF injection compared with the fibrotic area of the TS group at 14 days post-injury (3047.86 ± 857.20 vs. 809.44 ± 254.23 μm2, p < 0.05) (Figure 4a–d).
FIGURE 4
In addition, we tested the mRNA expression of collagen I and III in injured skeletal muscle. The results revealed that MGF injection significantly decreased the expression of collagen I mRNA levels at 1 day (p < 0.01) and 3 days (p < 0.01) after muscle injury compared with the TS group (Figure 5a). The expression of collagen III mRNA decreased in the MGF-treated mice at 1 (p < 0.01) and 3 days (p < 0.01) post-injury compared with the collagen III mRNA levels of the TS group (Figure 5b).
FIGURE 5
MGF Injection Did Not Influence the Functional Status of Satellite Cells in Injured Skeletal Muscle After Macrophage Depletion
Myogenic differentiation antigen (MyoD) is an important factor for muscle regeneration and is considered a marker of proliferation in satellite cells, and myogenin is a marker of differentiation in satellite cells (). In this study, we found that macrophage depletion did not influence the expression of MyoD (Figure 6a) but did significantly decrease the expression of myogenin at 3 days (p < 0.01) post-muscle injury (Figure 6b). However, injection of MGF did not influence the expression of MyoD and myogenin in the injured skeletal muscle after macrophage depletion (Figure 6a,b).
FIGURE 6
MGF Injection Decreased Expression of Inflammatory Cytokines in Injured Skeletal Muscle After Macrophage Depletion
RT-PCR demonstrated that levels of pro-inflammatory cytokines (TNF-α, IL-1β, and TGF-β) were significantly higher in the TS group than in the S group 3 days after the skeletal muscle injury. In contrast, the injection of MGF resulted in significantly decreased TNF-α, IFN-γ, IL-1β, and TGF-β levels in the TSM group post-injury compared with the TS group (Figure 7a–c).
FIGURE 7
MGF Injection Regulated Chemokine Expression in Injured Skeletal Muscle After Macrophage Depletion
Chemokines are important cytokines in skeletal muscle regeneration and fibrosis (; ). Compared with the S group, the group of mice with macrophage depleted muscle showed higher levels of CCL2, CCL5, and CXCR4 at 3 and 7 days post-injury (Figure 8a,b,d). Compared with the TS group, the MGF-treated group showed significantly decreased expression levels of CCL2, CCL5, and CXCR4 at 3 and 7 days post-injury. However, there was no significant change in the expression of CCR2 mRNA in the TSM group compared with the TS group post-injury (p > 0.05) (Figure 8c).
FIGURE 8
MGF Injection Decreased the Expression of gp91phox in Injured Skeletal Muscle After Macrophage Depletion
Oxidative stress contributes to fibrotic scar formation after skeletal muscle injury (). In this study, we tested whether MGF injection improves oxidative stress in injured skeletal muscle after macrophage depletion. The RT-PCR results showed that MGF injection significantly decreased the expression of gp91phox, a key subunit of NADPH oxidases (), in the TSM group compared with the TS group at 3 and 14 days post-injury (p < 0.05) (Figure 9). In addition, the immunofluorescence staining showed similar results (Figure 10).
FIGURE 9
FIGURE 10
MGF Injection Decreased the Expression of Matrix Metalloproteinases in Injured Skeletal Muscle After Macrophage Depletion
Macrophage depletion caused significant increases in MMP-1, MMP-2, MMP-9, MMP-10, and MMP-14 levels in injured skeletal muscle compared with the MMPs levels in the S group after injury. Compared with the TS group, the group that received the injection of MGF exhibited significantly increased levels of skeletal muscle MMP-1, MMP-2, MMP-9, MMP-10, and MMP-14 (p < 0.01) at 3 days after muscle injury (Figure 11a–e).
FIGURE 11
Discussion
To investigate the protective effect of MGF in impaired skeletal muscle regeneration after macrophage depletion and the underlying mechanisms involved, we generated a muscle contusion and macrophage depletion mouse model. In our previous study, we found that injection of clodronate-containing liposomes was an effective method to deplete the macrophage in injured skeletal muscle (). Consistent with that study, we found that injection of clodronate-containing liposomes significantly decreases the expression of a macrophage marker (F4/80) (Figure 2). These results suggested that macrophages were effectively depleted. In addition, we found that MGF injection has no influence on the expression of the macrophage marker (F4/80) after clodronate-containing liposomes were injected into injured skeletal muscle.
The H&E results indicated that macrophage depletion impairs muscle regeneration and that MGF injection exerts no protective effect, as evidenced by the sum of the diameters, numbers, and areas of the regenerating muscle fibers (Figure 3). These results are consistent with those of , who found that MGF overexpression did not notably affect muscle regeneration outcomes.
Furthermore, we tested the effects of MGF treatment on the fibrosis of injured muscle after macrophage depletion. Masson’s trichrome staining results showed that MGF injection significantly decreased the fibrotic area in the GMs in the TSM group compared with the fibrotic area in the TS group at 14 days post-injury (Figure 4). The fibrotic areas of the injured skeletal muscles exhibited markedly increased levels of collagen types I and III in the ECM of the muscle (; ). indicated that exogenous MGF-E peptide reduced collagen type I/III synthesis in cultured anterior cruciate ligament fibroblasts. Consistent with the aforementioned result, the levels of collagens I and III decreased significantly in the TSM group compared with the levels in the TS group at 1 and 3 days post-injury (Figure 5a,b). This result suggests that MGF injection can decrease the fibrosis of injured muscle after macrophage depletion.
To study the mechanism mediating in these results, we examined the expression of MyoD (a marker of proliferation of satellite cells) and myogenin (a marker of differentiation of satellite cells) (). Our results showed that injection of MGF did not influence the expression of MyoD and myogenin in the injured skeletal muscle after macrophage depletion (Figure 6a,b). These results suggest that MGF injection improves skeletal muscle fibrosis after macrophage depletion but not by regulating the functional status of satellite cells.
Furthermore, we examined the expression of inflammatory cytokines after muscle injury. Previous studies have shown that TGF-β plays a central role in skeletal muscle fibrosis (; ). Our data showed that MGF treatment significantly decreased the expression of TGF-β mRNA in the TSM group compared with the expression in the TS group at 3 days post-injury (p < 0.01, Figure 7d). This result may explain why MGF injection significantly decreases muscle fibrosis after macrophage depletion. Furthermore, we analyzed the expression of IL-1β, TNF-α, and IFN-γ mRNA after muscle injury. Some inflammatory cytokines (such as IL-1β, TNF-α, and TGF-β) are significantly higher after macrophage depletion in injured skeletal muscle, which may come from other inflammatory cells (). Furthermore, our results showed that MGF treatment significantly decreased the levels of IL-1β, TNF-α, and IFN-γ (Figure 7a–c) in the TSM group compared with the levels in the TS group. These results were similar to the findings of another study that showed that exogenous MGF-E peptide downregulates the protein levels of IL-1β and TNF-α in fibroblast-like synoviocytes in osteoarthritis (). The present results suggest that MGF injection improves skeletal muscle fibrosis after macrophage depletion by decreasing the expression of pro-inflammatory cytokines (TNF-α, IL-1β, and TGF-β).
Chemokines are not only involved in the migration and activation of monocytes, neutrophils, macrophages and lymphocytes but also play a key role in muscle regeneration (). In this study, we found that macrophage depletion significantly increased the expression of chemokines (CCL2, CCL5 and CXCR4) (Figure 8a,b,d). However, MGF injection significantly decreased the fibrosis of injured muscle tissue after macrophage depletion and decreased the expression of the three chemokines (CCL2, CCL5, and CXCR4). In other disease models, chemokines are involved in the fibrosis process of injured tissues, much like CCL5 contributes to liver fibrosis in non-alcoholic fatty liver disease progression () and CXCR4 contributes to cardiac fibrosis in diabetes (). This result suggests that the decreased levels of chemokines may be conducive to improving muscle fibrosis after MGF injection.
Because NADPH oxidase-induced reactive oxygen species (ROS) are involved in the fibrosis of injured skeletal muscle (; ), we analyzed the expression of gp91phox, a key subunit and common marker of NADPH oxidase (). The results showed that the injection of MGF significantly decreased the expression of gp91phox in the TSM group compared with the expression in the TS group at 3 days post-injury (p < 0.01) (Figure 9). Several studies using gene knockout mice or pharmacological agents to deplete gp91phox resulted in significantly reduced fibrosis in the kidney, liver, and heart (). This result suggests that gp91phox may be involved in the process underlying the improved muscle fibrosis after macrophage depletion as a result of MGF treatment.
In addition, we investigated the expression of MMPs, which play important roles in skeletal muscle regeneration and fibrosis. It has been acknowledged that MMPs can electively digest the extracellular matrix (ECM) in physiological and pathological states (). Inappropriate ECM digestion contributes to muscle fibrosis. The mRNA levels of MMPs (MMP-1, MMP-2, MMP-9, and MMP-10) (Figure 11a–d) were significantly downregulated in the MGF-treated group after macrophage depletion in the injured muscle compared to the MMP mRNA levels in the TS group. In other muscle injury disease models (such as dystrophic muscle in mdx mice), MMPs are significantly upregulated, whereas inhibitors of MMPs alleviate the related pathology and improve skeletal muscle function (). This observation may be help explain why the injection of MGF improves muscle fibrosis.
Conclusion
The results suggest that the impairment of skeletal muscle regeneration induced by macrophage depletion could be partly ameliorated by MGF injection and that inflammatory cytokines, oxidative stress factors, chemokines and matrix metalloproteinases may be involved in the process.
Statements
Ethics statement
The study was approved by the Ethics Review Committee for Animal Experimentation of Shanghai University of Sport, Shanghai, China (Reference No. 2014025).
Author contributions
WX and PC designed this study and helped to draft the manuscript. XL carried out data analysis and drafted the manuscript. XL, ZZ, and LZ performed the histological staining and carried out the real time PCR. All authors have read and approved the final version of the manuscript, and agreed with the order of presentation of the authors.
Funding
The study was supported by grants from the National Natural Science Foundation of China (Grant No. 31300975), the Natural Science Foundation of Shanghai (Grant No. 18ZR1437100), and the Key Lab of Development and Protection of Human Sports Ability in Shanghai (Grant No. 11DZ2261100).
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Footnotes
References
1
BrzoskaE.KowalewskaM.Markowska-ZagrajekA.KowalskiK.ArchackaK.ZimowskaM.et al (2012). Sdf-1 (CXCL12) improves skeletal muscle regeneration via the mobilisation of Cxcr4 and CD34 expressing cells.Biol. Cell104722–737. 10.1111/boc.201200022
2
Cabello-VerrugioC.AcunaM. J.MoralesM. G.BecerraA.SimonF.BrandanE. (2011). Fibrotic response induced by angiotensin-II requires NAD(P)H oxidase-induced reactive oxygen species (ROS) in skeletal muscle cells.Biochem. Biophys. Res. Commun.410665–670. 10.1016/j.bbrc.2011.06.051
3
ChanE. C.JiangF.PeshavariyaH. M.DustingG. J. (2009). Regulation of cell proliferation by NADPH oxidase-mediated signaling: potential roles in tissue repair, regenerative medicine and tissue engineering.Pharmacol. Ther.12297–108. 10.1016/j.pharmthera.2009.02.005
4
ChenX.LiY. (2009). Role of matrix metalloproteinases in skeletal muscle: migration, differentiation, regeneration and fibrosis.Cell Adh. Migr.3337–341.
5
ChuP. Y.WalderK.HorlockD.WilliamsD.NelsonE.ByrneM.et al (2015). CXCR4 antagonism attenuates the development of diabetic cardiac fibrosis.PLoS One10:e0133616. 10.1371/journal.pone.0133616
6
CornelisonD. D.WoldB. J. (1997). Single-cell analysis of regulatory gene expression in quiescent and activated mouse skeletal muscle satellite cells.Dev. Biol.191270–283.
7
GhalyA.MarshD. R. (2010). Ischaemia-reperfusion modulates inflammation and fibrosis of skeletal muscle after contusion injury.Int. J. Exp. Pathol.91244–255. 10.1111/j.1365-2613.2010.00708.x
8
GrefteS.Kuijpers-JagtmanA. M.TorensmaR.Von den HoffJ. W. (2010). Skeletal muscle fibrosis: the effect of stromal-derived factor-1α-loaded collagen scaffolds.Regen. Med.5737–747. 10.2217/rme.10.69
9
HuebnerK. D.JassalD. S.HalevyO.PinesM.AndersonJ. E. (2008). Functional resolution of fibrosis in mdx mouse dystrophic heart and skeletal muscle by halofuginone.Am. J. Physiol. Heart Circ. Physiol.294H1550–H1561. 10.1152/ajpheart.01253.2007
10
KharrazY.GuerraJ.MannC. J.SerranoA. L.Munoz-CanovesP. (2013). Macrophage plasticity and the role of inflammation in skeletal muscle repair.Mediat. Inflamm.2013:491497. 10.1155/2013/491497
11
KozakowskaM.Pietraszek-GremplewiczK.JozkowiczA.DulakJ. (2015). The role of oxidative stress in skeletal muscle injury and regeneration: focus on antioxidant enzymes.J. Muscle Res. Cell Motil.36377–393. 10.1007/s10974-015-9438-9
12
KumarA.BhatnagarS.KumarA. (2010). Matrix metalloproteinase inhibitor batimastat alleviates pathology and improves skeletal muscle function in dystrophin-deficient mdx mice.Am. J. Pathol.177248–260. 10.2353/ajpath.2010.091176
13
LiB. H.HeF. P.YangX.ChenY. W.FanJ. G. (2017). Steatosis induced CCL5 contributes to early-stage liver fibrosis in nonalcoholic fatty liver disease progress.Transl. Res.180:e4. 10.1016/j.trsl.2016.08.006
14
LiH.LeiM.YuC.LvY.SongY.YangL. (2015). Mechano growth factor-E regulates apoptosis and inflammatory responses in fibroblast-like synoviocytes of knee osteoarthritis.Int. Orthop.392503–2509. 10.1007/s00264-015-2974-5
15
LieberR. L.WardS. R. (2013). Cellular mechanisms of tissue fibrosis. 4. Structural and functional consequences of skeletal muscle fibrosis.Am. J. Physiol. Cell Physiol.305C241–C252. 10.1152/ajpcell.00173.2013
16
LiuX.LiuY.ZhaoL.ZengZ.XiaoW.ChenP. (2016). Macrophage depletion impairs skeletal muscle regeneration: the roles of regulatory factors for muscle regeneration.Cell Biol. Int.41228–238. 10.1002/cbin.10705
17
LivakK. J.SchmittgenT. D. (2001). Analysis of relative gene expression data using real-time quantitative PCR and the 2(T)(-Delta Delta C) method.Methods25402–408.
18
MillsP.DominiqueJ. C.LafreniereJ. F.BouchentoufM.TremblayJ. P. (2007). A synthetic mechano growth factor E Peptide enhances myogenic precursor cell transplantation success.Am. J. Transplant.72247–2259.
19
NovakM. L.BryerS. C.ChengM.NguyenM. H.ConleyK. L.CunninghamA. K.et al (2011). Macrophage-specific expression of urokinase-type plasminogen activator promotes skeletal muscle regeneration.J. Immunol.1871448–1457. 10.4049/jimmunol.1004091
20
NovakM. L.KohT. J. (2013). Macrophage phenotypes during tissue repair.J. Leukoc. Biol.93875–881. 10.1189/jlb.1012512
21
NovakM. L.Weinheimer-HausE. M.KohT. J. (2014). Macrophage activation and skeletal muscle healing following traumatic injury.J. Pathol.232344–355. 10.1002/path.4301
22
PanduroM.BenoistC.MathisD. (2018). Treg cells limit IFN-gamma production to control macrophage accrual and phenotype during skeletal muscle regeneration.Proc. Natl. Acad. Sci. U.S.A.115E2585–E2593. 10.1073/pnas.1800618115
23
PhilippouA.StavropoulouA.SourlaA.PissimissisN.HalapasA.MaridakiM.et al (2008). Characterization of a rabbit antihuman mechano growth factor (MGF) polyclonal antibody against the last 24 amino acids of the E domain.In Vivo2227–35.
24
PillonN. J.BilanP. J.FinkL. N.KlipA. (2013). Cross-talk between skeletal muscle and immune cells: muscle-derived mediators and metabolic implications.Am. J. Physiol. Endocrinol. Metab.304E453–E465. 10.1152/ajpendo.00553.2012
25
PopovichP. G.GuanZ.WeiP.HuitingaI.van RooijenN.StokesB. T. (1999). Depletion of hematogenous macrophages promotes partial hindlimb recovery and neuroanatomical repair after experimental spinal cord injury.Exp. Neurol.158351–365.
26
SaclierM.Yacoub-YoussefH.MackeyA. L.ArnoldL.ArdjouneH.MagnanM.et al (2013). Differentially activated macrophages orchestrate myogenic precursor cell fate during human skeletal muscle regeneration.Stem Cells31384–396. 10.1002/stem.1288
27
SawanoS.SuzukiT.DoM. K.OhtsuboH.MizunoyaW.IkeuchiY.et al (2014). Supplementary immunocytochemistry of hepatocyte growth factor production in activated macrophages early in muscle regeneration.Anim. Sci. J.85994–1000. 10.1111/asj.12264
28
ShaY.AfandiR.ZhangB.YangL.LvY. (2017). MGF E peptide pretreatment improves collagen synthesis and cell proliferation of injured human ACL fibroblasts via MEK-ERK1/2 signaling pathway.Growth Factors3529–38. 10.1080/08977194.2017.1327856
29
ShonoJ.SakaguchiS.SuzukiT.DoM. K.MizunoyaW.NakamuraM.et al (2013). Preliminary time-course study of antiinflammatory macrophage infiltration in crush-injured skeletal muscle.Anim. Sci. J.84744–750. 10.1111/asj.12105
30
SummanM.WarrenG. L.MercerR. R.ChapmanR.HuldermanT.Van RooijenN.et al (2006). Macrophages and skeletal muscle regeneration: a clodronate-containing liposome depletion study.Am. J. Physiol. Regul. Integr. Comp. Physiol.290R1488–R1495.
31
SunK.-T.CheungK.-K.AuS. W. N.YeungS. S.YeungE. W. (2018). Overexpression of mechano-growth factor modulates inflammatory cytokine expression and macrophage resolution in skeletal muscle injury.Front. Physiol.9:999. 10.3389/fphys.2018.00999
32
TidballJ. G. (2011). Mechanisms of muscle injury, repair, and regeneration.Compr. Physiol.12029–2062. 10.1002/cphy.c100092
33
TynerJ. W.UchidaO.KajiwaraN.KimE. Y.PatelA. C.OSullivanM. P.et al (2005). CCL5-CCR5 interaction provides antiapoptotic signals for macrophage survival during viral infection.Nat. Med.111180–1187.
34
WarrenG. L.OFarrellL.SummanM.HuldermanT.MishraD.LusterM. I.et al (2004). Role of CC chemokines in skeletal muscle functional restoration after injury.Am. J. Physiol. Cell Physiol.286C1031–C1036.
35
XiaoW.ChenP.WangR.DongJ. (2013). Overload training inhibits phagocytosis and ROS generation of peritoneal macrophages: role of IGF-1 and MGF.Eur. J. Appl. Physiol.113117–125. 10.1007/s00421-012-2418-5
36
XiaoW.LiuY.ChenP. (2016a). Macrophage depletion impairs skeletal muscle regeneration: the roles of pro-fibrotic factors, inflammation, and oxidative stress.Inflammation392016–2028.
37
XiaoW.LiuY.LuoB.ZhaoL.LiuX.ZengZ.et al (2016b). Time-dependent gene expression analysis after mouse skeletal muscle contusion.J. Sport Health Sci.5101–108. 10.1016/j.jshs.2016.01.017
38
ZhuJ.LiY.ShenW.QiaoC.AmbrosioF.LavasaniM.et al (2007). Relationships between transforming growth factor-beta1, myostatin, and decorin: implications for skeletal muscle fibrosis.J. Biol. Chem.28225852–25863.
Summary
Keywords
MGF, muscle regeneration, inflammatory cytokines, oxidative stress factors, chemokines, matrix metalloproteinases
Citation
Liu X, Zeng Z, Zhao L, Chen P and Xiao W (2019) Impaired Skeletal Muscle Regeneration Induced by Macrophage Depletion Could Be Partly Ameliorated by MGF Injection. Front. Physiol. 10:601. doi: 10.3389/fphys.2019.00601
Received
28 January 2019
Accepted
26 April 2019
Published
17 May 2019
Volume
10 - 2019
Edited by
Stefano Biressi, University of Trento, Italy
Reviewed by
Marina Bouche, Sapienza University of Rome, Italy; Jacob Sorensen, University of Minnesota Twin Cities, United States
Updates
Copyright
© 2019 Liu, Zeng, Zhao, Chen and Xiao.
This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Peijie Chen, chenpeijie@sus.edu.cn Weihua Xiao, xiao_weihua@163.com
†These authors have contributed equally to this work
This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology
Disclaimer
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.