Oxidative Medicine and Cellular Longevity

Oxidative Medicine and Cellular Longevity / 2018 / Article

Research Article | Open Access

Volume 2018 |Article ID 2393570 | 9 pages | https://doi.org/10.1155/2018/2393570

Mild Hyperthermia-Induced Myogenic Differentiation in Skeletal Muscle Cells: Implications for Local Hyperthermic Therapy for Skeletal Muscle Injury

Academic Editor: Steven McAnulty
Received08 Feb 2018
Accepted05 Jun 2018
Published27 Jun 2018

Abstract

The percutaneous application of controlled temperature on damaged muscle is regarded as a prevalent remedy. However, specific mechanisms are not completely understood. Therefore, cellular behaviors of myoblasts were investigated under a physiological hyperthermic temperature. The myoblasts were cultured under no treatment (NT, 37°C, 24 h/day), intermittent heat treatment (IHT, 39°C, 2 h/day), and continuous heat treatment (CHT, 39°C, 24 h/day) during proliferation, migration, or myogenic differentiation. Although the effects of mild heat on migration were not observed, the proliferation was promoted by both IHT and CHT. The myogenic differentiation was also enhanced in a treatment time-dependent manner, as evidenced by an increase in myotube size and fusion index. The gene expressions of mitochondrial biogenesis (Pgc-1α, Nrf1, and Tfam), a subset of mitochondrial dynamics (Mfn1 and Drp1), and a myogenic regulatory factor (myogenin) were increased in a heat treatment time-dependent manner. Interestingly, the mild heat-induced myogenic differentiation and myogenin expression were retarded significantly in PGC-1α-targeted siRNA-transfected cells, suggesting that mild hyperthermia promotes myogenic differentiation via the modulation of PGC-1α. This study provides cellular evidence supporting that local hyperthermic treatment at 39°C is regarded as an effective therapeutic strategy to promote satellite cell activities in regenerating myofibers.

1. Introduction

Recovery from skeletal muscle injury is orchestrated by a series of events consisting of degeneration, inflammation, and regeneration [1]. In the early phase of muscle recovery, significant destruction of myofibers called necrosis occurs, which is immediately followed by the removal of damaged cellular debris [2]. It is reported that, based on the level of injury state, the degeneration and inflammatory phase last up to 1 to 7 days [3, 4]. During the inflammatory stage, macrophages matured from monocytes not only remove damaged cells by phagocytosis but produce chemotactic signaling molecules leading to the activation of satellite cells for the proper regeneration of injured myofibers [3].

In the field of rehabilitation and sports medicine, there has been a great deal of effort in promoting skeletal muscle recovery. The most prevalent noninvasive strategy is the local application of controlled temperature on the damaged skeletal muscle. In the past, the local hypothemic therapy was regarded to be an effective strategy in the management of injury-induced pain, swelling, and inflammatory reactions. [58]. However, a study suggested that skeletal muscle regeneration is attenuated by hypothemic therapy due to the excessive suppression of inflammatory reactions, suggesting that a proper inflammatory process in the initial phase of recovery is critical for the facilitated regeneration of myofibers [9]. In line with this, heat treatment has been demonstrated to increase macrophage activation, leading to an increase in the number of regenerating muscle fibers in the damaged extensor digitorum longus of experimental rats [4]. In addition, several lines of studies proved that the application of mild heat treatment at 39°C is conducive to myogenesis while temperature at 41°C is detrimental to satellite cell activities [10, 11], providing a piece of evidence that the intrinsic biological capacity of satellite cell is tightly modulated in a temperature-sensitive manner.

Satellite cells are multipotent myogenic stem cells and exist between the sarcolemma and the basement membrane of myofibers [12]. Satellite cell-mediated muscle regeneration involves the orchestration of a series of cellular processes. Specifically, once activated, quiescent satellite cells enter proliferation phase and migrate to the site of injury, followed by myogenic differentiation to form new fibers or to incorporate into existing fibers [13, 14], all which steps are critical for the proper remodeling of the skeletal muscle. However, specific mechanisms of heat-mediated myogenic differentiation are not fully understood.

To elucidate cellular behaviors of myoblasts, cells were cultured under three different conditions in the study. The value of chronic heat treatment (CHT) was set at 39°C since this temperature can be easily reached in deep skeletal muscles under physiological systems [15]. In comparison, the myoblasts (IHT) were subjected to a cycle of 2 hour 39°C and 22 hour 37°C until being assayed. The heat-treated cells were compared with the cells cultured at 37°C.

2. Materials and Methods

2.1. Cell Culture

C2C12 myoblasts were seeded onto collagen-coated 6-well plates and were maintained in Dulbecco’s Modified Eagle’s Medium (Welgen, Kyungsan, Korea) containing 10% fetal bovine serum (FBS), 100 units/mL penicillin, and 100 mg/mL streptomycin (Welgene, Kyungsan, Korea) in a humidified atmosphere of 5% CO2 at 37°C or 39°C. When the myoblasts were confluent (95%), growth medium (10% FBS) was changed to differentiation medium (DM) supplemented with 2% horse serum, 100 units/mL penicillin, and 100 mg/mL streptomycin (Welgene, Kyungsan, Korea). Cells were differentiated for 3 to 5 days at 37°C or 39°C. For chronic heat treatment, proliferating or differentiating myoblasts were incubated at 39°C continuously for the indicated time. For intermittent heat treatment (IHT), proliferating or differentiating myoblasts were incubated for 2 hours at 39°C, followed by 22 hours at 37°C a day. This cycle was repeated until the end of each experimental protocol. The specific incubation time for each experiment is described in each figure legend. Cell culture was performed in a minimum of four independent times. siRNA transfection was performed as previously described [16]. Briefly, Pgc-1α siRNA or scrambled siRNA (Santa Cruz Biotechnology, CA, USA) was prediluted in OPTI-MEM medium (Gibco, MD, USA) containing Lipofectamine 2000 (Gibco, MD, USA). Lipofectamine-siRNA complexes were added into each well immediately after cells were plated in DMEM + 5% FBS without 100 units/mL penicillin, and 100 mg/mL streptomycin (final concentration of each siRNA was 10 nM). After 6 h of incubation, the media was exchanged with DMEM supplemented with 10% FBS for 18 h.

2.2. Cell Counting Assay

Cell counting assay was conducted using the trypan blue exclusion assay with minor modification [17]. C2C12 myoblasts were plated onto 60 mm culture dishes at 30% confluence and grown in growth medium at the indicated temperatures. Medium was changed every 24 h for 48 hours. The cells were trypsinized and stained with trypan blue. The nonstained viable cells were counted using a hemacytometer. The number of cells was expressed as myoblast number/mL.

2.3. Myoblast Migration Assay

An acellular area was generated in 95% confluent monolayers of myoblasts by scraping with a 200 μL pipette tip. Myoblast migration test was performed in a culture medium (DMEM) supplemented with 0.1% FBS and 100 units/mL penicillin and 100 mg/mL streptomycin for 24 hours in a humidified atmosphere of 5% CO2 at the indicated temperature conditions. The myoblasts that migrated into the acellular area were measured. The images were acquired using a phase contrast microscope (Carl Zeiss, Jena, Germany) 24 hours after the generation of the cellular area.

2.4. Immunocytochemistry for Myosin Heavy Chain and Myonuclei

Differentiated myotubes were fixed and permeabilized by incubating in ice-cold methanol (Sigma-Aldrich, Saint Louis, USA) for 10 minutes. After rehydration in Dulbecco's phosphate-buffered saline (DPBS) three times, the myotubes were incubated in blocking solution including 2% bovine serum albumin (BSA) for 30 minutes at room temperature, followed by incubation in 2% BSA solution containing anti-sarcomeric myosin antibody MF-20 conjugated with Alexa Fluor 488 (eBioscience, San Diego, USA). After myonuclei were stained with 4,6-diamidino-2-phenylindole (DAPI) (Molecular Probes, Eugene, OR, USA), the micrographs were acquired under a fluorescence microscope (Carl Zeiss, Jena, Germany).

2.5. Fusion Index

Fusion index is a myological indicator showing the degree of myogenic differentiation to myotubes. The index is calculated as the number of nuclei in myotubes divided by the total number of nuclei observed in the field, which is expressed as %. The cells with a minimum number of three myonuclei were counted as differentiated myotubes.

2.6. qRT-PCR

Differentiated myotubes were washed with cold DPBS and lysed with TRIzol reagent. Chloroform was added for the separation of RNA from DNA and protein fractions. The RNA fraction was precipitated by the addition of isopropanol, followed by centrifugation at 12,000 for 8 minutes at 4°C. The RNA pellet was washed with 75% ethanol and centrifuged at 12,000 for 5 minutes at 4°C. After removal of the ethanol, the RNA pellets were air-dried before resuspending with RNAse free water. The RNA concentration was quantified using a NanoDrop spectrophotometer (NanoDrop Technologies, DE, USA). Reverse transcription was performed with a cDNA synthesis kit according to manufacturer’s instruction (Bioneer, Daejeon, Korea). Following complementary DNA synthesis, qRT-PCR was performed in a Real-Time PCR system (Applied Biosystems, CA, US) using a SYBR Green Master Mix (Bioline, Hanam, Korea), and the primer pair sets were described in Table 1. Cycle threshold (Ct) values were normalized to the housekeeping gene (HPRT1-F: 5- GACTTGCTCGAGATGTCATG -3, HPRT1-R: 5- TACAGTCATAGGAATGGACC -3).


GenePrimer sequences

Pgc-1αForwardTGATGTGAATGACTTGGATACAGACA
ReverseGCTCATTGTTGTACTGGTTGGATATG
Ncor1ForwardGACCCGAGGGAAGACTACCATT
ReverseATCCTTGTCCGAGGCAATTTG
Nrf1ForwardGAACGCCACCGATTTCACTGTC
ReverseCCCTACCACCCACGAATCTGG
TfamForwardCTGATGGGTATGGAGAAGGAGG
ReverseCCAACTTCAGCCATCTGCTCTTC
Mfn1ForwardTTGATCGAATAGCATCCGAGGA
ReverseCACAGCATTGCATTGATGACAG
Opa1ForwardGGTCACCACGAGAAATCTCAG
ReverseTCTTCCATTCCGTCTCTAGGTT
Drp1ForwardCGGGACAAGTTAATTCAGGACA
ReverseGTTCTCGGGCAGACAGTTTTC
Fis1ForwardGCTCTAAAGTATGTGCGAGGG
ReverseTGCCTACCAGTCCATCTTTCTT
MyogeninForwardCCTTGCTCAGCTCCCTCA
ForwardTGGGAGTTGCATTCACTGG
HPRTForwardGACTTGCTCGAGATGTCATG
ReverseTACAGTCATAGGAATGGACC

2.7. Statistical Analysis

The results were presented as mean ± SEM for a minimum of four independent experiments. One-way ANOVA was followed by Tukey’s post hoc test to present statistical difference among groups. The statistical significance was set at .

3. Results

3.1. Effects of IHT and CHT on Migration and Proliferation of Myoblasts

The migrating myoblasts (%) were not changed statistically in the IHT and CHT groups compared to NT (Figures 1(a) and 1(b)). The myoblast proliferation was increased by the IHT and CHT in a treatment time-dependent manner (, Figure 1(c))

3.2. Effects of IHT and CHT on Myogenic Differentiation

On day 3, the number of myotubes formed during differentiation was higher in the CHT group compared to the IHT and NT groups (, Figures 2(a) and 2(b)). On day 4, the number of myotubes was higher in the IHT group compared to NT group while the number was the highest in the CHT group (Figures 2(a) and 2(b)). To explore myotube morphology in depth, the myosin heavy chain and myonuclei in myotubes differentiated for 4 days were stained with MF20 and DAPI (4,6-diamidino-2-phenylindole) (Figure 2(b)). The total myonuclei number was lower in the CHT group compared to the IHT and NT groups (Figure 2(d)) while the myotube fusion index (calculated as the number of nuclei in myotubes divided by the total number of nuclei counted) was greatest in the CHT group (, Figure 2(e)), indicating that more myoblasts were involved in myogenic differentiation. The width and length of myotubes were the greatest in the CHT, followed by the IHT and NT (, Figures 2(f) and 2(g)).

3.3. Effects of IHT and CHT on the Gene Expressions of Mitochondrial Remodeling

The multiple gene expressions of mitochondrial biogenesis (Pgc1-α, Nrf1, and Tfam) were increased in a treatment time-dependent manner (, Figure 3(a)). The expression of Ncor1, a negative regulator of mitochondrial biogenesis, was not changed by both treatments (Figure 3(a)). The gene expressions of all the mitochondrial dynamics markers (Mfn1, Opa1, Drp1, and Fis1) were increased by both treatment methods. Specifically, the expressions of Mfn1 and Drp1 were the greatest in the CHT group, followed by the IHT and NT group (, Figure 3(b)).

3.4. Effects of IHT and CHT on the Gene Expressions of Myosin Heavy Chain Isoforms and Myogenin

The gene expressions of myosin heavy chain isoforms and myogenic markers were measured. The expression of MyHCI was increased in both treatment groups in a treatment time-dependent manner. The expression of MyHCIIa was increased only in the CHT group compared to NT group (, Figure 4(a)). The expression of MyHCIIx and IIb were not altered by both treatments. The expression of myogenin was increased in a treatment time-dependent manner (, Figure 4(b)).

3.5. PGC-1α-Dependent Mild Heat-Induced Myogenic Differentiation

The mild heat-induced width and length of myotubes and myogenin expression were decreased to the level of nonheat-treated cells in the PGC-1α-targeted siRNA-transfected cells (, Figure 5).

4. Discussion

The application of controlled temperature on injury site has been regarded as a prevalent noninvasive method for myofiber regeneration [8, 18]. However, cellular mechanisms are not completely understood. Here, we demonstrated that mild hyperthermic treatment promotes myoblast proliferation and myogenic differentiation in a treatment time-dependent manner, but not migration. Our results also suggested that mild heat-induced myogenic differentiation is controlled in PGC-1α-dependent manner.

Satellite cells exist in a sublaminar niche in a mitotically quiescent state under basal conditions [19]. Once activated by injury-induced extracellular stimuli such as hepatocyte growth factor and nitric oxide, quiescent satellite cells enter the phase of self-renewing process to transform to proliferating myogenic cells called myoblasts [20]. In the current study, the mild hyperthermic treatments stimulated the rate of myoblasts proliferation. This result is in accordance with a previous study reporting that nuclear DNA contents, a surrogate marker of proliferation, were increased in a temperature-sensitive manner [21], suggesting a possibility that mild heat treatment at 39°C promotes an initial step of satellite self-renewing process. This idea is further supported by a rodent study demonstrating that the percutaneous application of a 20 minute heat treatment, immediately after injury, to the extensor digitorum longus (EDL) increased the number of regenerating fibers [4].

Satellite cells move along myofibers in an external chemotactic molecule-dependent manner [22]. Since satellite cells are sparsely scattered along the myofiber, an efficient motility of myoblasts is warranted for rapid myofiber recovery [23]. In the present study, the hyperthermic treatment did not change the rate of myoblast migration (Figures 1(a) and 1(b)). This suggests that hyperthermia alone is not sufficient to accelerate the motility of satellite cells. Currently, the effect of mild heat treatment on myoblast migration has not been reported elsewhere. As it has been known that the rate of myoblast migration is significantly affected by external molecular signals such as HGF, FGF, and SDF-1 [22], future studies are warranted to test if heat treatment would have synergistic effects on satellite cell migration under the treatment of known external chemoattractants.

Myoblast fusion is initiated by myogenin-induced myogenic differentiation to incorporate into myofibers, which is considered a critical and irreversible step in satellite cell-mediated muscle regeneration [19]. In the present study, a greater number of myotubes was observed in both CHT and IHT cells compared to NT at the fourth day of differentiation. The increased myogenic differentiation is further supported by the enhanced fusion index and myotube sizes in a heat treatment time-dependent fashion (Figures 2(a)2(g)). In previous in vivo studies using rodents, the effects of hyperthermia have been demonstrated by immersing animals with injured legs into warm water for only 20 to 30 minutes a day to avoid the excessive increase of the body temperature [4, 24, 25]. Therefore, it still remains elusive if a longer treatment would further enhance the rate of skeletal muscle regeneration. Regarding this notion, our results imply a piece of cellular possibility that, at a phase of myogenic differentiation, the duration of local hyperthermic therapy to injured skeletal muscle could be extended to facilitate the rate of satellite cell differentiation in regenerating myofibers.

Mitochondria are dynamic subcellular organelles regulating their size and mass as necessary, and their biogenesis and remodeling through fusion and fission are essential to maintain metabolic capacity in myofibers [2628]. In this study, the hyperthermic treatments increased the expressions of mitochondrial biogenic (Pgc-1α, Nrf1, and Tfam) and remodeling (Mfn1, Opa1, Drp1, and Fis1) markers (Figures 3(a) and 3(b)). In line with this, multiple studies proved that mitochondrial biogenesis and remodeling of myoblasts are essential steps in myogenic differentiation [29, 30]. This suggests that metabolic shift to mitochondrial oxidative phosphorylation take a central role in mild heat-mediated myogenic differentiation. This notion is further supported by the enhanced expressions of mitochondria-rich oxidative fibers (MyHCI and MyHCIIa) and myogenic regulatory factor (myogenin) in the heat-treated cells (Figure 4). Of the myogenic regulatory factors, myogenin has been known not only to initiate the processes of myogenesis but also to act as a significant determinant to myofiber characteristics. Specifically, it has been documented that myogenin is predominately expressed in slow-twitch muscles [31, 32], indicating a distinctive role of myogenin in transforming muscle fiber to more oxidative fiber type. As the myogenin was expressed in a treatment time-dependent manner, it appears that myogenin plays a significant role in increasing the expressions of more oxidative fibers (MyHCI and MyHCIIa) during heat-mediated myogenesis.

It has been widely accepted that PGC-1α, a master regulator of mitochondrial biogenesis, plays a central role in a regulatory network controlling numerous mitochondria-related transcription factors during myogenic differentiation [33, 34]. As it has been proven that depletion of PGC-1α attenuates myogenin expression [35], myoblasts were transfected with PGC-1α siRNA and differentiated under hyperthermic condition to investigate a possible mechanism of heat-mediated myogenic differentiation. Interestingly, the mild heat-induced differentiation potentials were significantly attenuated in the PGC-1α-targeted siRNA-transfected cells (Figures 5(a)5(c)). This is consistent with the reduced expressions of myogenin under heat treatment in the transfected cells (Figure 5(d)), indicating that mild heat-induced myogenic differentiation is controlled via the modulation of PGC-1α and myogenin axis.

In summary, this study demonstrated that mild heat treatment promotes the rate of proliferation and myogenic differentiation in a treatment time-dependent manner, which is regarded to provide an important piece of cellular evidence that clinical local hyperthermic therapy could be considered as an effective strategy to enhance the rate of regenerating myofibers (Figure 6). However, on the interpretation of our study, it should be noted that the temperature was set at 39°C as a mild hyperthermic condition. Given that the temperature in skeletal muscles could be elevated up to 44°C during hyperthermia and myogenic differentiation potential is significantly dysregulated at 41°C [36], a careful consideration should be taken into account to develop an effective hyperthermic therapy.

Data Availability

The data used to support the findings of this study are available from the corresponding author upon request.

Conflicts of Interest

The authors declare that there is no conflict of interest regarding the publication of this paper.

Authors’ Contributions

All authors participated in the design, interpretation of the studies, and review of the manuscript.

Acknowledgments

This work was supported by National Research Foundation of Korea (Grant NRF-2016R1C1B2015125) and Kyungsung University Research Grants in 2018.

References

  1. A. J. Quintero, V. J. Wright, F. H. Fu, and J. Huard, “Stem cells for the treatment of skeletal muscle injury,” Clinics in Sports Medicine, vol. 28, no. 1, pp. 1–11, 2009. View at: Publisher Site | Google Scholar
  2. T. A. H. Järvinen, T. L. N. Järvinen, M. Kääriäinen, H. Kalimo, and M. Järvinen, “Muscle injuries: biology and treatment,” The American Journal of Sports Medicine, vol. 33, no. 5, pp. 745–764, 2005. View at: Publisher Site | Google Scholar
  3. J. G. Tidball, “Inflammatory processes in muscle injury and repair,” American Journal of Physiology. Regulatory, Integrative and Comparative Physiology, vol. 288, no. 2, pp. R345–R353, 2005. View at: Publisher Site | Google Scholar
  4. K. Takeuchi, T. Hatade, S. Wakamiya, N. Fujita, T. Arakawa, and A. Miki, “Heat stress promotes skeletal muscle regeneration after crush injury in rats,” Acta Histochemica, vol. 116, no. 2, pp. 327–334, 2014. View at: Publisher Site | Google Scholar
  5. S. Michlovitz, W. Smith, and M. Watkins, “Ice and high voltage pulsed stimulation in treatment of acute lateral ankle sprains,” The Journal of Orthopaedic and Sports Physical Therapy, vol. 9, no. 9, pp. 301–304, 1988. View at: Publisher Site | Google Scholar
  6. T. L. Smith, W. W. Curl, B. P. Smith et al., “New skeletal muscle model for the longitudinal study of alterations in microcirculation following contusion and cryotherapy,” Microsurgery, vol. 14, no. 8, pp. 487–493, 1993. View at: Publisher Site | Google Scholar
  7. D. N. Deal, J. Tipton, E. Rosencrance, W. W. Curl, and T. L. Smith, “Ice reduces edema. a study of microvascular permeability in rats,” The Journal of Bone and Joint Surgery-American volume, vol. 84, no. 9, pp. 1573–1578, 2002. View at: Publisher Site | Google Scholar
  8. C. Bleakley, S. McDonough, and D. MacAuley, “The use of ice in the treatment of acute soft-tissue injury: a systematic review of randomized controlled trials,” The American Journal of Sports Medicine, vol. 32, no. 1, pp. 251–261, 2004. View at: Publisher Site | Google Scholar
  9. R. Takagi, N. Fujita, T. Arakawa, S. Kawada, N. Ishii, and A. Miki, “Influence of icing on muscle regeneration after crush injury to skeletal muscles in rats,” Journal of Applied Physiology, vol. 110, no. 2, pp. 382–388, 2011. View at: Publisher Site | Google Scholar
  10. Q. Guo, D. Miller, H. An et al., “Controlled heat stress promotes myofibrillogenesis during myogenesis,” PLoS One, vol. 11, no. 11, article e0166294, 2016. View at: Publisher Site | Google Scholar
  11. K. Ikeda, A. Ito, M. Sato, S. Kanno, Y. Kawabe, and M. Kamihira, “Effects of heat stimulation and l-ascorbic acid 2-phosphate supplementation on myogenic differentiation of artificial skeletal muscle tissue constructs,” Journal of Tissue Engineering and Regenerative Medicine, vol. 11, no. 5, pp. 1322–1331, 2017. View at: Publisher Site | Google Scholar
  12. P. S. Zammit, T. A. Partridge, and Z. Yablonka-Reuveni, “The skeletal muscle satellite cell: the stem cell that came in from the cold,” The Journal of Histochemistry and Cytochemistry, vol. 54, no. 11, pp. 1177–1191, 2006. View at: Publisher Site | Google Scholar
  13. F. Relaix and P. S. Zammit, “Satellite cells are essential for skeletal muscle regeneration: the cell on the edge returns centre stage,” Development, vol. 139, no. 16, pp. 2845–2856, 2012. View at: Publisher Site | Google Scholar
  14. F. Kawano, Y. Takeno, N. Nakai et al., “Essential role of satellite cells in the growth of rat soleus muscle fibers,” American Journal of Physiology. Cell Physiology, vol. 295, no. 2, pp. C458–C467, 2008. View at: Publisher Site | Google Scholar
  15. V. J. Wirth, B. van Lunen, D. Mistry, E. Saliba, and F. McCue, “Temperature changes in deep muscles of humans during upper and lower extremity exercise,” Journal of Athletic Training, vol. 33, no. 3, pp. 211–215, 1998. View at: Google Scholar
  16. R. Srikuea, X. Zhang, O. K. Park-Sarge, and K. A. Esser, “VDR and CYP27B1 are expressed in C2C12 cells and regenerating skeletal muscle: potential role in suppression of myoblast proliferation,” American Journal of Physiology. Cell Physiology, vol. 303, no. 4, pp. C396–C405, 2012. View at: Publisher Site | Google Scholar
  17. A. Duttaroy, J. F. Qian, J. S. Smith, and E. Wang, “Up-regulated P21CIP1 expression is part of the regulation quantitatively controlling serum deprivation-induced apoptosis,” Journal of Cellular Biochemistry, vol. 64, no. 3, pp. 434–446, 1997. View at: Publisher Site | Google Scholar
  18. B. F. Taylor, C. A. Waring, and T. A. Brashear, “The effects of therapeutic application of heat or cold followed by static stretch on hamstring muscle length,” The Journal of Orthopaedic and Sports Physical Therapy, vol. 21, no. 5, pp. 283–286, 1995. View at: Publisher Site | Google Scholar
  19. F. Le Grand and M. A. Rudnicki, “Skeletal muscle satellite cells and adult myogenesis,” Current Opinion in Cell Biology, vol. 19, no. 6, pp. 628–633, 2007. View at: Publisher Site | Google Scholar
  20. X. Shi and D. J. Garry, “Muscle stem cells in development, regeneration, and disease,” Genes & Development, vol. 20, no. 13, pp. 1692–1708, 2006. View at: Publisher Site | Google Scholar
  21. R. L. Harding, O. Halevy, S. Yahav, and S. G. Velleman, “The effect of temperature on proliferation and differentiation of chicken skeletal muscle satellite cells isolated from different muscle types,” Physiological Reports, vol. 4, no. 8, article e12770, 2016. View at: Publisher Site | Google Scholar
  22. A. L. Siegel, K. Atchison, K. E. Fisher, G. E. Davis, and D. D. W. Cornelison, “3D timelapse analysis of muscle satellite cell motility,” Stem Cells, vol. 27, no. 10, pp. 2527–2538, 2009. View at: Publisher Site | Google Scholar
  23. R. Bischoff, “Chemotaxis of skeletal muscle satellite cells,” Developmental Dynamics, vol. 208, no. 4, pp. 505–515, 1997. View at: Publisher Site | Google Scholar
  24. T. Shibaguchi, T. Sugiura, T. Fujitsu et al., “Effects of icing or heat stress on the induction of fibrosis and/or regeneration of injured rat soleus muscle,” The Journal of Physiological Sciences, vol. 66, no. 4, pp. 345–357, 2016. View at: Publisher Site | Google Scholar
  25. T. Yoshihara, H. Naito, R. Kakigi et al., “Heat stress activates the Akt/mTOR signalling pathway in rat skeletal muscle,” Acta Physiologica, vol. 207, no. 2, pp. 416–426, 2013. View at: Publisher Site | Google Scholar
  26. A. Garnier, D. Fortin, J. Zoll et al., “Coordinated changes in mitochondrial function and biogenesis in healthy and diseased human skeletal muscle,” The FASEB Journal, vol. 19, no. 1, pp. 43–52, 2005. View at: Publisher Site | Google Scholar
  27. H. Ding, N. Jiang, H. Liu et al., “Response of mitochondrial fusion and fission protein gene expression to exercise in rat skeletal muscle,” Biochimica et Biophysica Acta (BBA) - General Subjects, vol. 1800, no. 3, pp. 250–256, 2010. View at: Publisher Site | Google Scholar
  28. Z. Yan, V. A. Lira, and N. P. Greene, “Exercise training-induced regulation of mitochondrial quality,” Exercise and Sport Sciences Reviews, vol. 40, no. 3, pp. 159–164, 2012. View at: Publisher Site | Google Scholar
  29. A. H. V. Remels, R. C. J. Langen, P. Schrauwen, G. Schaart, A. M. W. J. Schols, and H. R. Gosker, “Regulation of mitochondrial biogenesis during myogenesis,” Molecular and Cellular Endocrinology, vol. 315, no. 1-2, pp. 113–120, 2010. View at: Publisher Site | Google Scholar
  30. J. Sin, A. M. Andres, D. J. R. Taylor et al., “Mitophagy is required for mitochondrial biogenesis and myogenic differentiation of C2C12 myoblasts,” Autophagy, vol. 12, no. 2, pp. 369–380, 2015. View at: Publisher Site | Google Scholar
  31. L. N. Zhu, Y. Ren, J. Q. Chen, and Y. Z. Wang, “Effects of myogenin on muscle fiber types and key metabolic enzymes in gene transfer mice and C2C12 myoblasts,” Gene, vol. 532, no. 2, pp. 246–252, 2013. View at: Publisher Site | Google Scholar
  32. S. M. Hughes, K. Koishi, M. Rudnicki, and A. M. Maggs, “MyoD protein is differentially accumulated in fast and slow skeletal muscle fibres and required for normal fibre type balance in rodents,” Mechanisms of Development, vol. 61, no. 1-2, pp. 151–163, 1997. View at: Publisher Site | Google Scholar
  33. M. T. Ryan and N. J. Hoogenraad, “Mitochondrial-nuclear communications,” Annual Review of Biochemistry, vol. 76, no. 1, pp. 701–722, 2007. View at: Publisher Site | Google Scholar
  34. R. C. Scarpulla, “Transcriptional paradigms in mammalian mitochondrial biogenesis and function,” Physiological Reviews, vol. 88, no. 2, pp. 611–638, 2008. View at: Publisher Site | Google Scholar
  35. S. Baldelli, K. Aquilano, and M. R. Ciriolo, “PGC-1α buffers ROS-mediated removal of mitochondria during myogenesis,” Cell Death & Disease, vol. 5, no. 11, article e1515, 2014. View at: Publisher Site | Google Scholar
  36. R. M. Borrell, R. Parker, E. J. Henley, D. Masley, and M. Repinecz, “Comparison of in vivo temperatures produced by hydrotherapy, paraffin wax treatment, and fluidotherapy,” Physical Therapy, vol. 60, no. 10, pp. 1273–1276, 1980. View at: Publisher Site | Google Scholar

Copyright © 2018 Hojun Lee and Seung-Jun Choi. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


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