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| ORIGINAL ARTICLE |
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| Year : 2020 | Volume
: 63
| Issue : 3 | Page : 113-121 |
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Muscle type from which satellite cells are derived plays a role in their damage response
Chuang-Yu Lin1, Chun-Yin Hou2, Chung-Min Tsai3, Hsi Chang4
1 Department of Clinical Application, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan
2 Department of Family Medicine, Taipei City Hospital, Zhongxiao Branch, Taipei, Taiwan
3 Graduate Institute of Medical Sciences, College of Medicine, Taipei Medical University, Taipei, Taiwan
4 Department of Pediatrics, School of Medicine, College of Medicine, Taipei Medical University; Department of Pediatrics, Taipei Medical University Hospital, Taipei, Taiwan
| Date of Submission | 24-Dec-2019 |
| Date of Decision | 15-Apr-2020 |
| Date of Acceptance | 07-May-2020 |
| Date of Web Publication | 23-Jun-2020 |
Correspondence Address:
Dr. Hsi Chang
Department of Pediatrics, Taipei Medical University Hospital, 252 Wu Xing Street, Taipei 11031
Taiwan
 Source of Support: None, Conflict of Interest: None  | 2 |
DOI: 10.4103/CJP.CJP_98_19
The aim of this study was to evaluate the response of satellite cells to muscular atrophies which possess different pathological characteristics and which were induced by distinct damages. Right lower limbs of rats were exposed to denervation or disuse and later its tibialis anterior (TA) or soleus (SOL) muscles were analyzed. After confirming their functional impairments indicated by common but distinct pathological and electrophysiological characteristics, the quantitative polymerase chain reaction analysis of Pax7 and Pax3 expressions and the number of Pax7+ve and Pax3+ve cells were analyzed sequentially at day 0, day 7, and day 14. TA muscles of both denervation- and disuse-induced atrophy models showed persisted low level of Pax7 expression from day 7 (0.91 ± 0.23 and 0.31 ± 0.07, P = 0.06, n = 6) through day 14 (1.09 ± 0.15 and 0.4 ± 0.09 [P < 0.05]). On the other hand, significant elevations were observed in Pax3 expression in both atrophy models (2.73 ± 0.46 and 2.75 ± 0.26 [P < 0.05]) at day 7. Similar to TA muscle, resembled pattern of Pax7 and Pax3 expression changes were observed between the SOL muscles of denervation- and disused-atrophy models. These trends were further confirmed by the changes in Pax7+ve and Pax3+ve cell numbers of TA and SOL muscles in both atrophy models. Despite the distinct pathological findings, similar patterns in the changes of Pax3 and Pax7 expressions and the changes of Pax7+ve and Pax3+ve cell numbers were observed between the denervation- and disuse-induced atrophy models and this commonality was admitted among the muscle type. Therefore, we claim that the muscle regeneration orchestrated by satellite cells was governed by the muscle type in which satellite cells reside.
Keywords: Denervation-induced atrophy model, disuse-induced atrophy model, muscle regeneration, muscle type, Pax3 and Pax7
How to cite this article:
Lin CY, Hou CY, Tsai CM, Chang H. Muscle type from which satellite cells are derived plays a role in their damage response. Chin J Physiol 2020;63:113-21 |
| Introduction | |  |
Postnatal skeletal muscle regeneration was mainly dependent on the activation of skeletal muscle stem cells, the satellite cells.[1] Muscle regeneration is immediately initiated whenever skeletal muscles are damaged by unaccustomed exercises[2] and a healthy skeletal muscle mass can be maintained only when the volumes of muscle degenerated and regenerated are balanced. Although physical exercise or training may damage skeletal muscles, avoiding physical activity is not conducive to skeletal muscle. In fact, the absence of physical activity leads to severe muscular deterioration.[3],[4],[5] Disuse-induced atrophy is a clinical situation associated with functional decline and disability which is often observed in bedridden patients while it seriously reduces their quality of life.[6] In addition to disuse, denervation is another frequently encountered clinical cause for muscular atrophy.[7],[8],[9],[10],[11] Its severity is various, depending on the regions of nerves that are damaged. For example, a spinal nerve injury is the most severe form, whereas facial palsy and carpal tunnel syndrome are considered milder forms.
As for disuse-induced atrophy, no effective treatment is available for denervation-induced atrophy at present and rehabilitation remains as the only evidence-based treatment.[12],[13],[14],[15],[16],[17],[18],[19] Although rehabilitation can delay the atrophy, it cannot completely arrest the progression, thereby avoiding the consequence of bedridden situation. To prevent such incidents, a combined strategy with rehabilitation such as diet regulation has been proposed.[20],[21],[22] As numerous studies have indicated an important role of satellite cell during muscle regeneration.[23],[24],[25],[26],[27],[28],[29] We believe that the goal of establishing a novel treatment could not be achieved without a thorough understanding of the responses of satellite cells during atrophy.
Clinically, atrophies induced by different causes show distinct pathological features. For example, the histological finding of a myopathic atrophy reveals rounded myofibers with marked variability in the fiber size. By contrast, muscles of neurologic atrophy exhibit “group atrophy,” the atrophy of an assembly of myofibers.[30],[31]
Since atrophies caused by different etiologies exhibit distinct histological characteristics, we raised the simple question regarding whether the satellite cells during the atrophies, which present distinct pathological characteristics, display differences in the responses. To answer this question, we used denervation- and disuse-induced atrophy rat models for our experiments. After confirmed their distinct pathological characteristics of two models, we examined the sequential changes in the expression of Pax7 and Pax3 and the cell number of Pax7+ve and Pax3+ve cells within tibia anterior (TA) and soleus (SOL) muscles of the two models. As we known, Pax genes play key roles in not only myogenesis during the embryonic stage but also postnatal muscle regeneration. Pax7 is a satellite cell marker that is specifically expressed in quiescent and newly activated satellite cells, and Pax3 is its paralog. Together Pax7 and Pax3 control the entry of myogenic program[24],[32] through the activation of myogenic regulatory factors (MRFs). The Myf5 and MyoD are also known as MRFs which profoundly involved with muscle regeneration.[33] However, since both MRFs locate at the downstream of Pax3 and Pax7; and the interaction between these Pax genes and MRFs are complicated. To address our observation specifically, the results of Pax3 and Pax7 were emphasized in this study. Although Pax7+ve and Pax3+ve cells were both known to participate in regeneration events of skeletal muscle[34] and possess similar properties, their different roles during adult muscle regeneration were confirmed.[32] In fact, Pax3 is expressed by a myogenic population distinct from the Pax7+ve satellite cell lineage which contributes to postnatal muscle regeneration;[35] therefore, we believe that the observation of changes in Pax7 and Pax3 expression and the Pax7+ve and Pax3+ve cell number in these two atrophy models which possess distinct pathological characteristics helps us to achieve a better understanding of how satellite cells response to the atrophy induced by distinct cause.
| Materials and Methods | | |
All animal handling procedures followed the Guide for the Care and Use of Laboratory Animals, published by the National Institutes of Health (NIH Publication No. 85-23, revised 1996), and the Guidelines of the Animal Research Committee of Taipei Medical University (approval number: LAC-2016-0458).
Denervation model
Male Sprague–Dawley rats (190–210 g) were used in this study. 1–3 rats were kept per cage (9 in ×12 in × 9 in). All animals were kept in a quiet animal facility with temperature of 21°C–23°C and a relative humidity of 30%–70% under a 12 h light/dark cycle. The procedure of denervation was as follows: in order to relieve the pain during the procedures, the rats were anesthetized using tiletamine and zolazepam (intraperitoneally, 25–30 mg/kg body weight) and 2% xylazine hydrochloride (intraperitoneally, 5–10 mg/kg body weight). The skin on the posterior surface of the right (Rt.) thigh was incised to expose the Rt. sciatic nerve. A 5-mm segment of the nerve was excised at the upper thigh level [Figure 1]a. The remaining ends of the nerve were allowed to retract spontaneously. The wound was closed in two layers using 6.0 nylon sutures. The rats were returned to the cages, where they had free access to food and water. There were six rats used as denervation model. | Figure 1: Both denervation and disuse models exhibited marked atrophy with functional impairment. The sciatic nerve of the Rt. lower limb was severed in rats in the denervation group (n = 6), and rats in the disuse group (n = 6) were covered with restriction suits at their Rt. lower limbs (a). Muscular atrophy was identified from the 1st week (b). The circumference of the Rt. tibialis anterior and soleus muscles in both groups decreased considerably in the 1st week (c). Functional impairment was also confirmed in both models (d). *P < 0.05.
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Disuse model
As in the denervation model, male Sprague–Dawley rats (190–210 g) were used for the experiment. The rats were maintained under the same condition as denervation group. Instead of nerve excision, the disuse rats were equipped with handmade restriction suits that were designed to inhibit the activity of their Rt. lower limbs [Figure 1]a during whole experimental period. All animals were confirmed to be accessible to food and water. There were six rats used as disuse model.
Control group
We used two different groups of rats as control for denervation and disuse model, respectively. Male Sprague–Dawley rats (190–210 g) with sham surgery were used for control group of denervation model. Sham surgery was performed as below under anesthesia (intraperitoneally, 25–30 mg/kg body weight); the Rt. thigh of rats was dissected with no sciatic nerve excision and the wound was closed in two layers using 6.0 nylon sutures. Later, the Rt. limb was used as control of denervation model. On the other hand, the Rt. limbs of male Sprague–Dawley rats (190–210 g) without any restriction were used as control of disused group. There were three rats used for each model and totally six rats were used as control group.
Footprint analysis
All the rats were subjected to footprint analysis. After dipping the hind feet in carbon ink, each rat was placed on a paper-lined walkway (8 cm × 42 cm) that led into a darkened cage.[36] The footprints of the Rt. limbs obtained on the paper as the rat walked down the walkway were measured and expressed as the mean ± standard deviation (SD) for comparison. For each rat, we measured three footprints out of the five steps taken by the rat to walk down the walkway. The footprints were postoperatively evaluated each week. The following three parameters of the footprints were measured: (i) print length (PL), the distance from the heel to the third toe; (ii) toe spread (TS), the distance from the first to the fifth toe, and (iii) intermediate TS, the distance from the second to the fourth toe. The data were used to compute the following parameters: (i) PL factor (PLF) = (experimental PL − normal PL [NPL])/NPL; (ii) TS factor (TSF) = (experimental TS − normal TS [NTS])/NTS, and (iii) intermediary TSF (ITSF) = (experimental intermediary TS − normal intermediary TS [NITS])/NITS, where E indicates the experimental side and N indicates the normal control. These factors were adapted to the Baine–Mackinnon–Hunter sciatic function index (SFI)[37] using the following formula:
SFI = −38.3 × PLF + 109.5 × TSF + 13.3 × ITSF − 8.8.
Electromyogram
The electromyogram (EMG) was performed as described previously.[38] Briefly, the EMG was performed on Rt. TA muscle of every rat in this study. The rats that received an EMG test were anesthetized using pentobarbital sodium (intraperitoneally, 25–30 mg/kg body weight) during the examination. The ADS3 constant current stimulator and CCU1 constant current unit (Upward Biosystems, Taipei, Taiwan) were used for performing the EMG test. The stimulating point and signal receiver were set at the upper quarter and lower quarter of the muscle, respectively. Electrical stimulation was provided in duration mode 4 at 200 mA.
Hematoxylin and eosin staining
TA and SOL muscles were isolated and frozen in liquid nitrogen-cooled isopentane. Serial transverse 5-μm-thick cryosections of muscle were cut and transferred onto slides. Later, the slides were incubated with Mayer's hematoxylin solution for 1 min and washed with running water for 5 min. After incubation in an eosin solution containing one drop of acetic acid, sections were rinsed quickly in water, hydrated using a gradient of ethanol solutions at 70%, 80%, 90%, 95%, and 100%, and mounted with xylene-based mounting media and covered with cover glass.
Quantitative polymerase chain reaction
Total RNA was isolated from skeletal muscles, namely the TA and SOL muscle, of the denervation and disuse model rats using the TRIzol reagent (Invitrogen). The following specific primers were used for quantitative polymerase chain reaction (qPCR): Pax3, sense, 5´-CAGCCCACGTCTATTCCACA-3´, and antisense, 5´-CACGAAGCTGTCGGTGTAGC-3´; Pax7, sense, 5´-AGCCGAGTGCTCAGAATCAA-3´, and antisense, 5´-TCCTCTCGAAAGCCTTCTCC-3´. Maxima® SYBR Gree/ROX qPCR Master Mix (Fermentas Life Sciences, OR, USA) was used for PCR amplification. The amplification program used was 40 cycles of 15 s at 95°C and 60 s at 60°C.
Immunocytochemical analysis
Immunocytochemical analysis was performed as described previously.[39] The antibodies used in this study were mouse anti-Pax7 and mouse anti-Pax3 (MAB1675, MAB2457; R&D Systems, Minneapolis, MN, USA) with a concentration of 10 μg/ml. Cy3-labeled antibodies against mouse or rabbit IgG and fluorescein isothiocyanate-labeled antibodies against mouse IgG (715-005-150, Jackson ImmunoResearch Laboratory, Bar Harbor, ME, USA) were applied as secondary antibodies. Hoechst 33324 (H3570; Molecular Probes) was used for nuclear staining. The samples were examined using a fluorescence microscope (Olympus, Tokyo, Japan) or an AS-MDW system (Leica Microsystems, Wetzlar, Germany). Micrographs were obtained using an AxioCam (Carl Zeiss Vision, Hallbergmoos, Germany) or a AS-MDW system (Leica Microsystems). In sections of muscles, the numbers of Pax7+ve and Pax3+ve cells were counted, per field, at a magnification of ×100. The average of ten fields in each tissue sample was calculated. The statistical analysis was performed for six rats for each group (n = 6).
Statistical analyses
The data were presented as mean ± SD. Statistical analyses were conducted using the unpaired Student's t-test, and significance was set at P < 0.05.
| Results | | |
Both denervation and disuse models exhibited marked atrophy with functional impairment
In this study, we intended to compare the responses of satellite cells between the denervation- and disuse-induced atrophy models, which showed distinct pathological characteristics. Prior to the comparison, we first confirmed whether atrophy with functional loss was evident in both models and further confirmed their distinct pathological characteristics. To prevent the bias caused by muscle type, we selected the TA and SOL muscles for analysis which are known to predominantly consist of fast and slow twitch myofibers, respectively. As expected, atrophy was clearly observed from the 1st week post of both denervation and disuse macroscopically [Figure 1]b. By measuring the circumference of the Rt. TA of denervated rats, the average circumference was 2.50 ± 0.21 cm (P < 0.05, n = 6), which showed significant atrophy relative to that of control (2.94 ± 0.23 cm) since 1st week. Resembled results were also observed at 2nd and 3rd weeks [Figure 1]c. The circumferences of Rt. SOL muscles were also significantly smaller than that of control (1.40 ± 0.18 cm, P < 0.05, n = 6). As denervation models, atrophies were also observed in Rt. TA and SOL muscles of disuse model from 1st week through 3rd week [TA: 2.65 ± 0.04 cm (P = 0.06), 2.3 ± 0.1 cm, and 1.79 ± 0.08 cm, SOL: 0.91 ± 0.18 cm, 0.8 ± 0.1 cm, and 0.53 ± 0.08 cm, P < 0.05, n = 6; [Figure 1]c. In addition to the morphological changes, we further evaluated the functional capability of both atrophic models using the SFI score. As expected, SFI score declined to 62.20 ± 10.08, which revealed a statistical significance (P < 0.05, n = 6) since 1st week in denervation model. The same as denervation model, the SFI score of disuse model was also decreased to 58.81 ± 3.27 [P < 0.05, n = 6; [Figure 1]d, Supplementary Video 1 ]. From these data, the remarkable atrophies accompanied with functional impairments were confirmed with both atrophy models.
[Additional file 1]
Histological and electrophysiological differences between denervation- and disuse-induced atrophy models
After morphological and functional assessment, the histological analysis was administered. The H and E staining of the Rt. TA muscles from the denervation model revealed “group atrophy” of myofibers (yellow arrowhead), which is a well-known pathological characteristic of neurogenic atrophy. By contrast, above-normal variability in fiber size due to the presence of both hypertrophic and atrophic fibers was observed in the Rt. TA muscles of the disuse model [Figure 2]a. In addition to the histological differences, discrepancies between the two models were also observed in the results obtained from the electrophysiological examination. Unlike the rapid amplitude depreciation observed in the EMGs of the Rt. TA muscle of denervation model, gradual amplitude depreciation was observed in the EMGs of the Rt. TA of disuse model [Figure 2]b. However, this difference in EMGs between denervation and disuse models become vague at 3rd week. | Figure 2: Histological and electrophysiological differences between the denervation- and disuse-induced atrophy models. Although both models exhibited muscular atrophy, the denervation model showed a form of group atrophy of myofibers (yellow arrowhead), whereas the disuse model exhibited a diffused and sporadic atrophic pattern (a). The denervation model exhibited an immediate deceleration of amplitude in the electromyogram, whereas the disuse model exhibited a gradual deceleration of amplitude in the electromyogram (b).
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Pax7 and Pax3 expression patterns were similar between muscle types but not between damage models
After confirming the pathological and electrophysiological differences, we investigated the sequential changes in Pax7 and Pax3 expression through qPCR at 1st and 2nd week post-denervation and disuse, respectively. The Pax7 expression in the Rt. TA muscles of the denervation model showed no significant elevation in both 1st and 2nd weeks [0.91 ± 0.23 and 1.09 ± 0.15, P = 0.06, n = 6; [Figure 3]a and e]. Similar to the results of denervation model, Pax7 expression in the Rt. TA muscles of the disuse model also remained low till 2nd week after disuse (0.31 ± 0.07 and 0.4 ± 0.09, P < 0.05, n = 6). By contrast, the Pax3 expression in the Rt. TA muscles in the denervation model increased rapidly from the 1st week but quickly declined in the 2nd week after damage (2.73 ± 0.46 and 1.79 ± 0.26, P < 0.05, n = 6). A similar trend was observed in Pax3 expression of Rt. TA muscles of disuse model [2.75 ± 0.26 and 1.25 ± 0.21, P < 0.05, n = 6; [Figure 3]b and e]. Like the changes in Rt. TA muscle, the expression patterns of Pax7 and Pax3 in Rt. SOL muscle for both models were also resembled. Both models exhibited a significant increase in the Pax7 expression in the 1st week (1.59 ± 0.29 and 3.84 ± 0.24, P < 0.05, n = 6), followed by a decrease in the 2nd week [0.76 ± 0.19 and 0.68 ± 0.28, P < 0.05, n = 6; [Figure 3]c and e], whereas the Pax3 expression in Rt. SOL muscle of both models showed a gradual increase [0.65 ± 0.31 to 1.45 ± 0.26 and 1.21 ± 0.44 to 1.82 ± 0.63, P < 0.05; [Figure 3]d and e]. These results indicated the similarities in Pax7 and Pax3 expression were observed between the muscle types but not between the damage models. | Figure 3: Pax7 and Pax3 expression in TA and SOL muscles of denervation and disuse models. The quantitative polymerase chain reaction expression of Pax7 in the tibialis anterior muscles for both groups remained constantly low (a). Pax3 expression pattern in the tibialis anterior muscles in both models rapidly increasing from the 1st week but declining from the 2nd week (b). The Pax7 expression in the soleus muscles of both models increased in the 1st week but decreased from 2nd week (c). Pax3 expression in the soleus muscles of both groups reached its peak in the 2nd week (d). The changes of Pax7 and Pax3 expression were summarized in a table (e).
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Changes in Pax7+ve and Pax3+ve cell numbers were also similar between muscle types
In addition to qPCR, an immunostained for Rt. TA and SOL muscles of two models with anti-Pax7 and anti-Pax3 antibodies [Figure 4]a was performed. The numbers of Pax7+ve and Pax3+ve cells were counted in the first and 2nd week postdamage, thereby confirming its consistency with the results of qPCR. Relative to the remarkable increase in Rt. SOL muscle in both denervation and disuse atrophy models in the 1st week [170.6 ± 8.1, 142.7 ± 6.1, P < 0.05, n = 6; [Figure 4]d and [Figure 4]f, mild increase of Pax7+ve cell was observed in the Rt. TA muscle of denervation and disuse induced atrophy models [46.5 ± 1.6 and 50.8 ± 4.8, P < 0.05, n = 6; [Figure 4]b and [Figure 4]f. On the other hand, a significant increase in Pax3+ve cells was observed in TA muscles at 1st week followed by a gradual decline at 2nd week in both models [from 139.4 ± 8.2 to 121.3 ± 8.4 and from 73.1 ± 4.1 to 38.5 ± 4.8, P < 0.05, n = 6; [Figure 4]c and [Figure 4]f. The elevation in Pax3+ve cell number were observed at 1st week for both models (101.7 ± 5.4 and 110.7 ± 11.9, P < 0.05) and followed by a decrease in the 2nd week after damage [56.9 ± 4.1 and 93.0 ± 2.1, P < 0.05; [Figure 4]e and [Figure 4]f. In line with the results of q-PCR, the similarity of changes in Pax7+ve and Pax3+ve cell number was observed between the muscle types but the atrophy models. | Figure 4: Changes in Pax7+ve and Pax3+ve cell numbers were also similar between muscle types. The numbers of Pax7+ve and Pax3+ve cells (Red) were counted sequentially (a). The number of Pax7+ve cell in the tibialis anterior muscles was markedly lower the number of Pax3+ve cell (b). The number of Pax3+ve cell in the tibialis anterior muscles increased from the 1st week and declined (c). In the soleus muscles, both Pax7+ve and Pax3+ve cell number increased from the 1st week, but declined from the 2nd week (d and e). The changes in the cell numbers were consistent with the results of quantitative polymerase chain reaction (f).
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| Discussion | | |
It is well known that muscular atrophies induced by different causes possess distinct pathological traits. Since satellite cells play a core role during muscle regeneration, to identify how the satellite cells responded while different types of damage induced atrophies showed distinct pathological characteristics is the purpose of this study.
In this study, the TA and SOL muscles of denervation- and disuse-induced atrophy models of rat were used and sequentially observed Pax3 and Pax7 expressions at 1st and 2nd week postdamage. Both models exhibited functional defects in common, but distinct results were confirmed in EMGs and histological examination. Despite the differences in EMGs and histology, similar patterns of expression for Pax7 and Pax3 were observed in the different damage models. By contrast, different expression patterns of Pax7 and Pax3 were observed between muscle types that consist of distinct proportions of fast and slow twitch muscle fibers. This phenomenon was observed not only in qPCR but also in the changes in Pax7+ve and Pax3+ve cell numbers. Therefore, we claim that the response of satellite cells that is induced by muscular damage is not altered by the mode of damage; however, the muscle type from which the satellite cells are derived or in which they reside plays a role to their response.
As we know, the skeletal muscle comprises fast and slow myofibers[40] and the proportion is unique according the muscle types. In this study, the commonality in the changes of expression of satellite cell markers and satellite cell numbers which responded to the damages was seen among the muscle type. Together, these data suggest a possible correlation of composition of myofibers (fast or slow) within the muscle types and the responses of satellite cells during muscular damage. In fact, our results are compatible to the theory previously reported which claimed the phenotype of myoblast reflects their fiber type of origin in a single-fiber culture.[41] Another study claimed the differential myogenicity among satellite cells from fast and slow muscles,[42] which is also consistent with our results.
Furthermore, a relatively limited plasticity of satellite cells under a normal physiological environment was speculated from the results of this study. Although further data are necessary to verify this hypothesis, previous reports of muscle flap engraftment technique support this idea. Muscle flap engraftment is a well-developed treatment in plastic surgery.[43],[44] The grafted muscle, along with the associated vessels and nerves, is transplanted onto the recipient region where the loss of skeletal muscle has occurred. In some cases, the transplantation has been performed between different muscle types of fast and slow. From those cases, the authors found that when the donor muscle tissue was successfully engrafted, the myofiber retained its original type and fused only with the muscle fibers of the donor site. Thus, when a slow muscle is transplanted onto a fast muscle, its property as a slow muscle is preserved even after a successful engraftment.[45],[46] However, the diversity of satellite cells was also been shown by other reports.[47],[48] As we mentioned, skeletal muscle comprises different ratio of fast and slow myofibers between muscle types. Therefore, we believe that further studies regarding the relationship between the satellite cells and myofiber type instead of muscle type are indispensable for solving this difficult question and consequently this will also be our next task.
Since the immobility is existed in both atrophic models, we could not completely exclude the effect of disuse in denervation model in this study. A new study model is needed to solve this problem and a regular passive muscle contraction after denervation may be a solution for it. Furthermore, there are deviations between the results of qPCR and immunohistochemistry. The reason for the difference may be due to the distinction of two methods. As the Western blot was used, the whole muscle as material but IHC analysis used single slice of muscle tissue for staining. Even though there are deviations between the results of qPCR and IHC analysis, the similar tendency can still be admitted among the same muscle type which received distinct types of muscle damage.
In this experiment, we observed that although differences in the mode of muscle damage caused differences in the electrophysiological and pathological characteristics of the damaged muscles, the responses of the satellite cells were similar in both cases. By contrast, different patterns of expression of Pax7 and Pax3 were observed between muscle types consisting of distinct proportions of fast and slow twitch muscle fibers after muscular damage. Therefore, we claim that the muscle regeneration orchestrated by satellite cells was governed by the muscle type from which the satellite cells were derived. Ascertaining whether separate satellite cells exist for fast twitch and slow twitch muscle fibers is difficult. From our experimental results, we believe that the plasticity of satellite cells is relatively limited. This is particularly relevant in considering satellite cells as a source of transplantation to muscle types other than their original type in the future.
Acknowledgments
We thank Dr. Geng-Chang Yeh and You-Shan Lin for their contribution to the concepts and statistical analysis contained in this manuscript.
Financial support and sponsorship
This study was supported by Grant-in-Aid for Scientific Research TMU100-AE1-B10, 101TMU-TMUH-11 from Taipei Medical University and Taipei Medical University Hospital and Grant-in-Aid for Scientific Research 101-2314-B-038 -017-MY3 from the Ministry of Science and Technology, R.O.C., Taiwan.
Conflicts of interest
There are no conflicts of interest.
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[Figure 1], [Figure 2], [Figure 3], [Figure 4]
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