Loss of skeletal muscle mass is generally accompanied by a decline in muscle strength, a reduction in functional capacity, and an increased risk of developing chronic metabolic diseases. Skeletal muscle mass maintenance is regulated by an intricate balance between muscle protein synthesis and muscle protein breakdown. Physical activity is one of the main factors responsible for regulating skeletal muscle protein turnover (18). A reduction in physical activity level can cause a substantial disruption in muscle protein balance, resulting in a decline in skeletal muscle mass. Accordingly, the decline in or complete loss of physical activity is considered the main cause responsible for the loss of skeletal muscle mass in various models of muscle atrophy (35). In this context, sarcopenia and spinal cord injury (SCI) have been studied intensively.
The loss of skeletal muscle mass with aging (sarcopenia) has been, at least partly, attributed to a more sedentary lifestyle in the elderly population (10). Sarcopenia has been characterized by Type II muscle fiber–specific atrophy (20,22). Interestingly, with the discovery of satellite cells as stem cells responsible for myofiber maintenance and growth (14), we recently observed that Type II muscle fiber atrophy with aging is accompanied by a Type II muscle fiber–specific decline in satellite cell content (37). This finding was later confirmed by other groups (39) and implies that the age-related loss of Type II muscle fiber satellite cell content plays a key role in muscle fiber atrophy and, as such, in the etiology of sarcopenia (38).
Interestingly, much of the decline in Type II muscle fiber size in the elderly can be reversed by prolonged resistance type exercise training (19,33). We recently reported that the increase in muscle mass and strength after 3 months of supervised resistance type exercise training in healthy elderly men is accompanied by a specific increase in Type II muscle fiber size and muscle fiber satellite cell content (36). Clearly, the level of physical (in)activity is instrumental in the regulation of skeletal muscle maintenance with aging.
SCI represents another model in which substantial atrophy is observed in the affected muscle tissue. The total absence of voluntary muscle contractile activity and/or loss of neurological input are followed by a rapid loss of skeletal muscle mass (6). Previously, it has been reported that the loss of muscle mass after SCI is characterized by a shift toward more Type II muscle fibers and severe atrophy of both the Types I and II muscle fibers (3,6,17,27). Given the regulatory role of satellite cells in myofiber maintenance, it could be speculated that the muscle fiber atrophy in SCI is also accompanied by a decline in the number of satellite cells (3). Furthermore, from a clinical perspective, satellite cell content in SCI subjects may represent a key factor determining the skeletal muscle adaptive capacity to respond to anabolic stimuli, such as neuromuscular electrical stimulation protocols. However, data on satellite cell content in a human SCI model are currently lacking. Therefore, we investigated whether our previous observations on muscle fiber type–specific atrophy and an accompanying decline in satellite cell content in a human aging model would also be observed in an SCI model of muscle atrophy. We hypothesized that muscle fiber type–specific atrophy in SCI is different from muscle atrophy with aging. In addition, we hypothesized that muscle fiber type–specific atrophy in both aging and SCI are accompanied by a fiber type–specific decline in satellite cell content.
In the present study, we collected muscle biopsies from eight young males with SCI, eight age-matched, able-bodied controls, and eight healthy elderly men and assessed muscle fiber type composition, muscle fiber size, and muscle fiber myonuclear and satellite cell content to investigate the changes in fiber type–specific muscle characteristics with aging and SCI.
A total of 24 male subjects volunteered to participate in this study. Only men were recruited to reduce within-group variability in muscle fiber characteristics. Paraplegic, spinal cord–injured, wheelchair-dependent subjects were recruited through patient organizations. The mean number of years postinjury for the SCI subjects was 9 ± 2 yr. The level of injury was between C5–C6 and T9-T10 for all SCI subjects and was incomplete for three subjects; however, no voluntary contractile activity of the vastus lateralis was reported. Three of the eight subjects reported regular spasms, which included spasms in the vastus lateralis muscle. Able-bodied, age-matched men were selected to participate as controls. In addition, eight elderly subjects were recruited through advertisements in local newspapers. All subjects were screened with an oral glucose tolerance test (OGTT) to exclude type 2 diabetes according to the World Health Organization guidelines, and medical history was evaluated. All subjects were healthy, independently living volunteers without any major comorbidity (apart from SCI). All elderly and able-bodied young subjects were recreationally active (i.e., walking/cycling/running two to three times a week) but had not participated in any structured (leg) endurance and/or resistance type exercise training program designed to improve performance for the past 3 yr. All subjects were informed of the nature and possible risks of the experimental procedures before their written informed consent was obtained. All procedures were performed in compliance with the Declaration of Helsinki, and the study was approved by the Medical Ethics Committee of the Maastricht University Medical Centre+.
All subjects received the same standardized meal (approximately 3.0 MJ: 57 energy% (En%) carbohydrate, 30 En% fat, and 13 En% protein) the evening before the test day and refrained from strenuous physical activity for 3 d before testing. On the test day, subjects arrived at the laboratory by car or public transportation after an overnight fast. After 30 min of supine rest, a single-slice axial image of the upper leg muscles was obtained 15 cm above the patella by either computed tomography or magnetic resonance imaging scanning to determine quadriceps muscle cross-sectional area (CSA) as described previously (36). Then, a percutaneous needle biopsy (20–80 mg) was taken from the vastus lateralis muscle, approximately 15 cm above the patella, after local anesthesia. Muscle biopsies were collected from the self-reported dominant leg in all subjects. Any visible nonmuscle tissue was removed from the biopsy samples, which were then embedded in Tissue-Tek (Sakura Finetek, Zoeterwoude, The Netherlands), immediately frozen in liquid nitrogen–cooled isopentane, and stored at −80°C until further analyses.
Blood samples from the OGTT were collected in ethylenediaminetetraacetic acid containing tubes and centrifuged at 1000g and 4°C for 10 min. Aliquots of plasma were frozen in liquid nitrogen and stored at −80°C. Plasma samples were analyzed for glucose (COBAS FARA, Uni Kit III; Roche, Basel, Switzerland) and insulin concentrations (Insulin RIA Kit; LINCO Research, Inc., St. Charles, MO). Plasma glucose and insulin concentrations from the OGTT were used to estimate whole-body insulin sensitivity using the oral glucose insulin sensitivity index.
From all muscle biopsies, 5-μm-thick cryosections were cut at −20°C. Biopsy samples from one subject in each group were mounted together on uncoated glass slides. Care was taken to properly align the samples for cross-sectional fiber analyses. Serial cross-sections were stained for muscle fiber typing and myocellular satellite cell content as described previously (36). Because it was recently reported that the number of Type IIx fibers in a muscle biopsy sample is generally insufficient to allow a reliable assessment of Type IIx muscle fiber size and/or satellite cell content (24,37), a differentiation was only made between Types I and II muscle fibers. In short, staining procedures were as follows: After fixation (5-min acetone), slides were air dried and incubated for 60 min at room temperature with primary antibodies directed against laminin and either myosin heavy chain (MHC)-I or CD56 to stain Type I muscle fibers and satellite cells, respectively. Slides were then washed (3× 5-min phosphate-buffered saline). Appropriate secondary antibodies were applied, which were diluted together with 4’,6-diamidino-2-phenylindole (DAPI) to stain myonuclei. After a final washing step, all slides were mounted with cover glasses. After staining, digital images were captured using fluorescence microscopy (36). All image recordings and analyses were performed by an investigator blinded to subject coding.
For the fiber typing slides, the number of muscle fibers, the mean fiber CSA, the number of myonuclei, and the myonuclear domain (i.e., fiber CSA/number of myonuclei) were measured for the Types I and Type II muscle fibers separately (Fig. 1). For further insight into the distribution and variability of muscle fiber size, the frequency distribution was calculated. Intervals of 1000 μm2 were defined and the percentage of muscle fibers in each interval was determined for the Types I and II muscle fibers separately.
For the satellite cell slides, fiber type was determined on the basis of the serial fiber typing slides. Satellite cells were determined at the periphery of the muscle fiber and stained positive for both CD56 and DAPI (Fig. 2). The number of satellite cells per muscle fiber, the number of satellite cells per square millimeter of muscle fiber, and the percentage of satellite cells (number of satellite cells / [number of myonuclei + number of satellite cells]× (mult) 100) were determined for the Types I and II muscle fibers separately. A mean total of 308 ± 40 muscle fibers were analyzed for each subject, with a minimum of 75 fibers to determine satellite cell content in Types I and II muscle fibers separately (24). Because of the low number of Type I muscle fibers in SCI biopsies, reliable Type I muscle fiber satellite cell counts could only be made for three of the eight subjects.
Some of the muscle samples from SCI subjects showed sarcoplasmic staining of CD56 in small muscle fibers (Fig. 2A, B). Additional analyses were performed to determine whether the latter was related to satellite cells per se or to degenerating or regenerating fibers (5,15). A pax7 antibody was used as described previously (37) to determine whether CD56-positive fibers might represent very small fibers containing a satellite cell (Fig. 2E, F). Furthermore, staining for embryonic MHC was performed as a marker for regenerating and/or newly formed fibers (16,23) (Fig. 2G, H). Finally, a cleaved caspase-3 antibody was used to determine whether apoptotic nuclei were present in muscle fibers (7) (Fig. 2I, J). Staining procedures were similar as described for CD56 (i.e., including laminin and DAPI).
Antibodies for immunohistochemical analyses were purchased from the Developmental Studies Hybridoma Bank (Iowa City, IA: A4.951 for MHC-I, F1.652 for embryonic MHC, and pax7), Sigma (Zwijndrecht, The Netherlands: laminin), BD Biosciences (San Jose, CA: CD56), Molecular Probes (Invitrogen, Breda, The Netherlands: DAPI and secondary antibodies), and Cell Signaling Technology (Bioke, Leiden, The Netherlands: caspase-3). Examples of all stainings are shown in Figures 1 and 2.
All data are expressed as means ± SEM. Normality of the data was confirmed with Shapiro–Wilk tests. Differences between groups were analyzed by one-way ANOVA. In case of significant differences, Tukey post hoc tests were performed to locate between-group differences. In addition, muscle fiber type–specific variables (i.e., muscle fiber CSA, myonuclear content, and satellite cell content) were analyzed by paired samples t-tests in the young control and elderly groups separately. The latter was not possible in the SCI group because only few Type I muscle fibers were present in these biopsies. All analyses were performed using SPSS version 18.0 (Chicago, IL). An α level of 0.05 was used to determine statistical significance.
Subjects’ characteristics are provided in Table 1. Mean age was similar for the young controls (31 ± 3 yr) and SCI subjects (32 ± 4 yr) but higher in the elderly (75 ± 2 yr). Body mass and body mass index were similar between groups. In contrast, quadriceps muscle CSA was significantly smaller in the order SCI < elderly < young controls (P < 0.001). Although fasting blood glucose concentrations were significantly higher in the elderly when compared with the young controls, all subjects had normal glucose tolerance.
Muscle tissue characteristics.
Muscle biopsies from both young and elderly subjects showed normal staining patterns and morphological appearance (Fig. 1A, B). Biopsy material from SCI subjects showed severe signs of degeneration and atrophy (Fig. 1C, D). Tearing and folding of the basal lamina were often observed, along with large heterogeneity in muscle fiber CSA (ranging from 50 to >10,000 μm2). These characteristics did not differ between SCI subjects with or without spasms. Furthermore, generalized CD56-positive staining of very small fibers was often observed in muscle tissue from SCI subjects (Fig. 2A, B), whereas this was not observed in the other groups. Therefore, additional pax7 staining was performed to confirm the presence of satellite cells in these very small CD56-positive fibers (37). In addition, staining with embryonic MHC and cleaved caspase-3 was performed to further characterize muscle tissue in SCI subjects (Fig. 2).
Muscle fiber type size, composition, and frequency distribution.
The percentage of Type I muscle fibers and the area percentage occupied by Type I muscle fibers were significantly lower in the SCI subjects when compared with both the able-bodied young and elderly subjects (P < 0.001, Table 2). In four of the eight SCI subjects, no Type I muscle fibers were present in the muscle tissue samples. No differences were observed in muscle fiber type percentage and/or muscle fiber area percentage between the able-bodied young and elderly subjects.
Mixed muscle fiber CSA was significantly different between groups and was smaller in the order SCI < elderly < young controls. For the Type I muscle fibers, CSA was significantly smaller in the subjects with SCI when compared with the able-bodied young (P < 0.001) and elderly subjects (P = 0.003). No differences were observed between the elderly and young controls (Fig. 3). In contrast, Type II muscle fiber CSA was much smaller in both the SCI (P < 0.001) and elderly (P = 0.001) subjects when compared with the young controls, with no differences between SCI and elderly (Fig. 3). Within the elderly subgroup, muscle fiber CSA was significantly smaller in the Type II versus Type I muscle fibers (P = 0.002). No differences were observed between Types I and II muscle fiber CSA within the SCI and young control groups. In addition, no differences were evident between SCI subjects with or without spasms.
Figure 4 shows the frequency distribution for Types I and II muscle fibers separately. For both the Types I and II muscle fibers, an obvious shift was observed toward the smaller muscle fibers for the SCI subjects when compared with the young controls. Furthermore, the large variability in muscle fiber size in the SCI subjects is clearly illustrated in Figure 4, with 4%–5% of all fibers being larger than 10,000 μm2. In the elderly, a shift toward a larger percentage of small muscle fibers was observed only for the Type II muscle fibers.
Whereas the number of nuclei per Type I muscle fiber did not differ between groups, the number of nuclei per Type II muscle fiber was significantly lower in both the SCI (P = 0.005) and elderly (P = 0.003) subjects when compared with the young controls (Table 2). In addition, within the elderly subgroup, the number of nuclei was lower in the Type II versus Type I muscle fibers (P = 0.020). Type I muscle fiber myonuclear domain was significantly larger in the elderly when compared with the SCI subjects (P = 0.012). No group differences were observed for Type II muscle fiber myonuclear domain. Notably, central myonuclei were observed in 1%–10% of the muscle fibers of the SCI subjects (Fig. 1C). In contrast, central myonuclei were hardly observed in the able-bodied young and elderly subjects.
The number of satellite cells per muscle fiber was lower in the SCI subjects when compared with the young controls for both the Type I (0.049 ± 0.019 vs 0.104 ± 0.011, respectively; P = 0.017) and the Type II muscle fibers (0.050 ± 0.005 vs 0.117 ± 0.009, respectively; P < 0.001) (Fig. 3). When expressed per square millimeter of muscle fiber and relative to the total number of myonuclei, differences in satellite cell content between SCI subjects and young controls were less evident (Table 2). Interestingly, many of the small fibers in SCI subjects were CD56 positive (Fig. 2A, B). However, these CD56-positive fibers did not represent satellite cells as evidenced by the absence of pax7 staining.
The number of satellite cells per muscle fiber was lower in the Type II muscle fibers in the elderly when compared with the young controls (P < 0.001, Fig. 3). Type II muscle fiber satellite cell content in the elderly remained significantly lower compared with the young controls when expressed per square millimeter of muscle fiber (P = 0.011) and relative to the total number of myonuclei (P = 0.006, Table 2). Within the elderly subgroup, the number of satellite cells per muscle fiber (P < 0.001), the number of satellite cells per square millimeter of muscle fiber (P = 0.012), and the percentage of satellite cells relative to the total number of myonuclei (P = 0.010) were significantly lower in the Type II versus Type I muscle fibers (Fig. 3, Table 2). No differences were observed in satellite cell content between the SCI and elderly subjects, although the number of satellite cells per square millimeter of Type II muscle fiber tended to be lower in the elderly (P = 0.105).
Embryonic MHC and caspase-3.
Figure 2G–J shows examples of the staining for embryonic MHC and caspase-3. Muscle fibers positive for embryonic MHC were observed in biopsy samples from two control subjects, three elderly subjects, and six SCI subjects. However, the number of fibers that stained positive for embryonic MHC was less than 1%, including the very small fibers in SCI subjects. Myonuclei staining positive for caspase-3 were only rarely observed (<0.5% of all myonuclei), with no differences between groups.
In the present study, we show that skeletal muscle atrophy as observed in SCI and aging is associated with fiber type–specific alterations at the myocellular level. Muscle atrophy in SCI is attributed to both Types I and II muscle fiber atrophy, with a shift toward more Type II muscle fibers. Muscle atrophy in senescent muscle is attributed to specific Type II muscle fiber atrophy, with no apparent shift in muscle fiber type composition. Muscle fiber atrophy as observed with SCI (Types I and II fibers) and aging (Type II fibers) is accompanied by a muscle fiber type–specific reduction in satellite cell content.
SCI and sarcopenia are both characterized by the loss of skeletal muscle mass and function. Despite obvious similarities, the extent of muscle atrophy as well as its underlying causes is quite different between these models. Such differences most likely reside at the myocellular level. In the present study, we assessed fiber type–specific differences in muscle fiber size and satellite cell enumeration in tissue collected from young males with SCI and elderly men. We observed specific Type II muscle fiber atrophy in muscle biopsies collected from the vastus lateralis muscle in elderly men (Fig. 3), which is in line with previous data from our laboratory (36,37) as well as others (20–22) showing a specific reduction in Type II muscle fiber size at a more advanced age. In line with and extending on these findings, we now show a shift in the frequency distribution of Type II muscle fibers with aging (Fig. 4). In the elderly, more than 50% of all Type II muscle fibers were smaller than 4000 μm2, compared with less than 15% in the young controls.
In the subjects with SCI, muscle fiber size was much smaller for both the Types I and II muscle fibers when compared with the young controls. Whereas the elderly showed an approximately 40% reduction in Type II muscle fiber size, the young males with SCI showed massive Types I and II muscle fiber atrophy of more than 50% (Fig. 3). Furthermore, muscle fiber type composition had shifted toward approximately 90% Type II muscle fibers in the subjects with SCI, whereas no such shift was observed in the elderly. The reduction in the number of slow-twitch Type I muscle fibers with SCI is in accordance with earlier reports (4,27,30) and has been suggested to be caused by changes in the expression of myogenic regulatory factors (i.e., MyoD and myogenin) and nuclear factor of activated T-cells, in response to reduced neuromuscular activity (34). Also, the histological findings in muscle tissue from subjects with SCI—with folding of the basal lamina, large variability in muscle fiber size, and the presence of central nuclei—are in agreement with the myopathic alterations and ultrastructural disorganization typically observed in previous studies examining denervation in humans (2,3,17,30). We now extend on these findings by quantifying the frequency distribution of muscle fibers on the basis of different fiber sizes (Fig. 4). As expected, the majority of muscle fibers in SCI subjects was very small, with approximately 60% of fibers being smaller than 3000 μm2 compared with only approximately 30% in the elderly. Strikingly however, we observed that approximately 5% of all muscle fibers of SCI subjects were larger than 10,000 μm2 (Fig. 4). Although the existence of such large muscle fibers many years post-SCI is interesting, it remains to be determined whether these fibers still possess a physiological function, i.e., more so than those fibers with a substantially smaller size. Taken together, our data provide further evidence that muscle atrophy at the myocellular level differs substantially between the human SCI and aging model, at least partly explaining the difference in quadriceps CSA between SCI subjects and the elderly. These findings are likely associated with the nature, duration, and extent of the reduction in physical and/or neuromuscular activity (34). In accordance, it could be speculated that an affected muscle that remains partially active (either through spasms or an incomplete lesion) might to a certain extent be protected against the consequences of SCI. Although we did not observe any differences in muscle fiber characteristics between SCI subjects with or without spasms, caution should be taken with interpreting these findings because only three spasmodic subjects were included. As such, it remains to be elucidated to what extent the presence of muscle spasms might affect the myocellular changes after SCI.
Over the last few years, the role of satellite cells in adult myogenesis has gained much interest. Satellite cells represent the only known source for the formation of new myonuclei in vivo (26). Satellite cells play an essential role in muscle fiber growth and repair and, as such, have been implicated as a key regulator of muscle mass maintenance in humans (14). In accordance, changes in satellite cell content and/or function have been associated with both skeletal muscle atrophy and hypertrophy (14). We were the first to report that Type II muscle fiber atrophy with aging is accompanied by a muscle fiber type–specific decline in satellite cell content (37,39). In the present study, we confirm (Fig. 3) and extend these findings by the observation that satellite cell content also declines in subjects with SCI, a human model of severe physical inactivity leading to profound muscle atrophy. Until now, there has been no consensus on the effect of SCI on satellite cell content. Whereas in animal models some have reported a reduction in satellite cell content after SCI and/or denervation (9,31), others have failed to confirm these findings (12). So far, data on satellite cell enumeration after SCI in a human model have been lacking. The present study is the first to show that muscle fiber satellite cell content is substantially reduced in subjects with SCI when compared with young controls (Fig. 3). Furthermore, the decline in satellite cell content was evident in both the Types I and II muscle fibers. Despite the reduced Type I muscle fiber size in SCI subjects compared with the elderly, no differences in Type I muscle fiber satellite cell content were observed. This is likely explained by the limited power to detect a seemingly large difference (Fig. 3). In fact, care should be taken when interpreting these Type I muscle fiber data because we were only able to reliably assess Type I muscle fiber characteristics in three of eight SCI subjects. In summary, the fiber type–specific atrophy in SCI (Types I and II muscle fibers) and aging (Type II fibers) is accompanied by a fiber type–specific decline in satellite cell content. These findings further support the idea that satellite cells play a key role in regulating muscle fiber size and, as such, muscle maintenance (38). However, it remains to be established whether the decline in satellite cell content is indeed a cause or rather a consequence of the muscle fiber atrophy observed in SCI and aging models of muscle atrophy.
Despite substantial muscle fiber atrophy, myonuclear domain size and the number of satellite cells expressed per area of muscle fiber had hardly been affected after more than 9 yr of SCI. This observation may likely explain why affected muscle tissue after SCI seems to retain a remarkable degree of plasticity in response to electrical stimulation (11). Regained muscle function and muscle (fiber) hypertrophy has previously been reported after 1 up to 9 yr of functional electrical stimulation regimens (1,3,17,32). In contrast with SCI subjects, Type II muscle fiber satellite cell content in the elderly remained low when corrected for muscle fiber area. Whether this phenomenon results in an attenuated response to intervention strategies aimed to increase muscle mass and function in the elderly is a matter of debate. Previous studies indicate that the extent of skeletal muscle hypertrophy in response to exercise is affected by both satellite cell content and by the ability to increase satellite cell pool size, providing additional myonuclei to fuse with existing myofibers (28,29,36). As such, the hypertrophic response after neuromuscular electrical stimulation in SCI subjects could be driven by the incorporation of new myonuclei provided through proliferation and differentiation of a dormant but viable pool of satellite cells. Future work will need to establish the capacity of the resident satellite cell pool to activate, proliferate, and differentiate, both with SCI and aging. Such insight may lead to the development of tailored interventional strategies to augment muscle mass and function in various models of muscle atrophy.
In the present study, we used the CD56 antibody as a marker for satellite cells located at the periphery of muscle fibers. In line with previous findings in paralyzed muscle from both human and animal studies (5,8,15), we observed a severely altered histological pattern and a substantial number of very small muscle fibers with CD56 expression scattered throughout the sarcoplasm in the subjects with SCI (Figs. 1 and 2). To further characterize this muscle tissue, some additional stainings were performed. The absence of pax7 immunostaining in small muscle fibers showed that they did not represent satellite cells. Alternatively, the CD56-positive cells may represent either denervated or regenerating muscle fibers (5,15). Based on the presence of some embryonic MHC-positive muscle fibers, regenerative processes seem to continue in skeletal muscle tissue that has been paralyzed for many years (17,40). However, the fibers expressing embryonic MHC were very small both in number and in size. As such, those fibers that continue to express CD56 in the sarcoplasm clearly fail to fully regenerate and presumably represent a population of severely atrophied and denervated muscle fibers (13). Finally, the absence of caspase-3 immunoreactivity showed that, although some of the histological findings may be the result of cell death, no current signs of apoptosis were evident.
In conclusion, SCI is associated with severe atrophy of both the Types I and II muscle fibers, with a shift toward approximately 90% Type II fibers. In contrast, atrophy in senescent muscle is characterized by specific Type II muscle fiber atrophy, without a shift in muscle fiber type composition. In both muscle atrophy models, a decline in muscle fiber size is accompanied by a muscle fiber type–specific reduction in satellite cell content.
DHJT is financially supported by the Netherlands Heart Foundations (E Dekker stipend, 2009T064).
The authors thank all the participants who volunteered for this study. None of the authors declared any conflict of interest.
The results of the present study do not constitute endorsement by the American College of Sports Medicine.
1. Andersen JL, Mohr T, Biering-Sorensen F, Galbo H, Kjaer M. Myosin heavy chain isoform transformation in single fibres from m. vastus lateralis in spinal cord injured individuals: effects of long-term functional electrical stimulation (FES). Pflugers Arch
. 1996; 431: 513–8.
2. Biral D, Kern H, Adami N, Boncompagni S, Protasi F, Carraro U. Atrophy-resistant fibers in permanent peripheral denervation of human skeletal muscle. Neurol Res
. 2008; 30: 137–44.
3. Boncompagni S, Kern H, Rossini K, et al.. Structural differentiation of skeletal muscle fibers in the absence of innervation in humans. Proc Natl Acad Sci U S A
. 2007; 104: 19339–44.
4. Burnham R, Martin T, Stein R, Bell G, MacLean I, Steadward R. Skeletal muscle fibre type transformation following spinal cord injury. Spinal Cord
. 1997; 35: 86–91.
5. Cashman NR, Covault J, Wollman RL, Sanes JR. Neural cell adhesion molecule in normal, denervated, and myopathic human muscle. Ann Neurol
. 1987; 21: 481–9.
6. Castro MJ, Apple DF Jr, Staron RS, Campos GE, Dudley GA. Influence of complete spinal cord injury on skeletal muscle within 6 mo of injury. J Appl Physiol
. 1999; 86: 350–8.
7. Conraads VM, Hoymans VY, Vermeulen T, et al.. Exercise capacity in chronic heart failure patients is related to active gene transcription in skeletal muscle and not apoptosis. Eur J Cardiovasc Prev Rehabil
. 2009; 16: 325–32.
8. Covault J, Sanes JR. Neural cell adhesion molecule (N-CAM) accumulates in denervated and paralyzed skeletal muscles. Proc Natl Acad Sci U S A
. 1985; 82: 4544–8.
9. Dedkov EI, Kostrominova TY, Borisov AB, Carlson BM. Reparative myogenesis in long-term denervated skeletal muscles of adult rats results in a reduction of the satellite cell population. Anat Rec
. 2001; 263: 139–54.
10. Doherty TJ. Invited review: aging and sarcopenia
. J Appl Physiol
. 2003; 95: 1717–27.
11. Dudley-Javoroski S, Shields RK. Muscle and bone plasticity after spinal cord injury: review of adaptations to disuse and to electrical muscle stimulation. J Rehabil Res Dev
. 2008; 45: 283–96.
12. Dupont-Versteegden EE, Murphy RJ, Houle JD, Gurley CM, Peterson CA. Activated satellite cells fail to restore myonuclear number in spinal cord transected and exercised rats. Am J Physiol
. 1999; 277: C589–97.
13. Gosztonyi G, Naschold U, Grozdanovic Z, Stoltenburg-Didinger G, Gossrau R. Expression of Leu-19 (CD56, N-CAM) and nitric oxide synthase (NOS) I in denervated and reinnervated human skeletal muscle. Microsc Res Tech
. 2001; 55: 187–97.
14. Hawke TJ, Garry DJ. Myogenic satellite cells: physiology to molecular biology. J Appl Physiol
. 2001; 91: 534–51.
15. Illa I, Leon-Monzon M, Dalakas MC. Regenerating and denervated human muscle fibers and satellite cells express neural cell adhesion molecule recognized by monoclonal antibodies to natural killer cells. Ann Neurol
. 1992; 31: 46–52.
16. Kadi F, Eriksson A, Holmner S, Butler-Browne GS, Thornell LE. Cellular adaptation of the trapezius muscle in strength-trained athletes. Histochem Cell Biol
. 1999; 111: 189–95.
17. Kern H, Boncompagni S, Rossini K, et al.. Long-term denervation in humans causes degeneration of both contractile and excitation-contraction coupling apparatus, which is reversible by functional electrical stimulation (FES): a role for myofiber regeneration? J Neuropathol Exp Neurol
. 2004; 63: 919–31.
18. Koopman R, van Loon LJ. Aging, exercise, and muscle protein metabolism. J Appl Physiol.
2009; 106: 2040–8.
19. Kosek DJ, Kim JS, Petrella JK, Cross JM, Bamman MM. Efficacy of 3 days/wk resistance training on myofiber hypertrophy and myogenic mechanisms in young vs. older adults. J Appl Physiol
. 2006; 101: 531–44.
20. Larsson L, Sjodin B, Karlsson J. Histochemical and biochemical changes in human skeletal muscle with age in sedentary males, age 22–65 years. Acta Physiol Scand
. 1978; 103: 31–9.
21. Lexell J. Human aging, muscle mass, and fiber type composition. J Gerontol A Biol Sci Med Sci
. 1995; 50 Spec No: 11–6.
22. Lexell J, Taylor CC, Sjostrom M. What is the cause of the ageing atrophy? Total number, size and proportion of different fiber types studied in whole vastus lateralis muscle from 15- to 83-year-old men. J Neurol Sci
. 1988; 84: 275–94.
23. Mackey AL, Esmarck B, Kadi F, et al.. Enhanced satellite cell proliferation with resistance training in elderly men and women. Scand J Med Sci Sports
. 2007; 17: 34–42.
24. Mackey AL, Kjaer M, Charifi N, et al.. Assessment of satellite cell number and activity status in human skeletal muscle biopsies. Muscle Nerve
. 2009; 40: 455–65.
25. Mari A, Pacini G, Murphy E, Ludvik B, Nolan JJ. A model-based method for assessing insulin sensitivity from the oral glucose tolerance test. Diabetes Care
. 2001; 24: 539–48.
26. Moss FP, Leblond CP. Nature of dividing nuclei in skeletal muscle of growing rats. J Cell Biol
. 1970; 44: 459–62.
27. Olsson MC, Kruger M, Meyer LH, et al.. Fibre type–specific increase in passive muscle tension in spinal cord–injured subjects with spasticity. J Physiol
. 2006; 577: 339–52.
28. Petrella JK, Kim JS, Mayhew DL, Cross JM, Bamman MM. Potent myofiber hypertrophy during resistance training in humans is associated with satellite cell-mediated myonuclear addition: a cluster analysis. J Appl Physiol
. 2008; 104: 1736–42.
29. Rosenblatt JD, Yong D, Parry DJ. Satellite cell activity is required for hypertrophy of overloaded adult rat muscle. Muscle Nerve
. 1994; 17: 608–13.
30. Scelsi R, Marchetti C, Poggi P, Lotta S, Lommi G. Muscle fiber type morphology and distribution in paraplegic patients with traumatic cord lesion. Histochemical and ultrastructural aspects of rectus femoris muscle. Acta Neuropathol
. 1982; 57: 243–8.
31. Schmalbruch H, Lewis DM. Dynamics of nuclei of muscle fibers and connective tissue cells in normal and denervated rat muscles. Muscle Nerve
. 2000; 23: 617–26.
32. Scremin AM, Kurta L, Gentili A, et al.. Increasing muscle mass in spinal cord injured persons with a functional electrical stimulation exercise program. Arch Phys Med Rehabil
. 1999; 80: 1531–6.
33. Singh MA, Ding W, Manfredi TJ, et al.. Insulin-like growth factor I in skeletal muscle after weight-lifting exercise in frail elders. Am J Physiol
. 1999; 277: E135–43.
34. Talmadge RJ. Myosin heavy chain isoform expression following reduced neuromuscular activity: potential regulatory mechanisms. Muscle Nerve
. 2000; 23: 661–79.
35. Urso ML. Disuse atrophy of human skeletal muscle: cell signaling and potential interventions. Med Sci Sports Exerc
. 2009; 41 (10): 1860–8.
36. Verdijk LB, Gleeson BG, Jonkers RA, et al.. Skeletal muscle hypertrophy following resistance training is accompanied by a fiber type–specific increase in satellite cell content in elderly men. J Gerontol A Biol Sci Med Sci
. 2009; 64: 332–9.
37. Verdijk LB, Koopman R, Schaart G, Meijer K, Savelberg HH, van Loon LJ. Satellite cell content is specifically reduced in type II skeletal muscle fibers in the elderly. Am J Physiol Endocrinol Metab
. 2007; 292: E151–7.
38. Verdijk LB, Snijders T, Beelen M, et al.. Characteristics of muscle fiber type are predictive of skeletal muscle mass and strength in elderly men. J Am Geriatr Soc
. 2010; 58: 2069–75.
39. Verney J, Kadi F, Charifi N, et al.. Effects of combined lower body endurance and upper body resistance training on the satellite cell pool in elderly subjects. Muscle Nerve
. 2008; 38: 1147–54.
40. Winter A, Bornemann A. NCAM, vimentin and neonatal myosin heavy chain expression in human muscle diseases. Neuropathol Appl Neurobiol
. 1999; 25: 417–24.