Scand J Med Sci Sports 2011: 21: 773–782 doi: 10.1111/j.1600-0838.2010.01178.x
& 2010 John Wiley & Sons A/S
Myogenic response of human skeletal muscle to 12 weeks of resistance training at light loading intensity A. L. Mackey1,2, L. Holm1,2, S. Reitelseder1,2, T. G. Pedersen1,2, S. Doessing1,2, F. Kadi3, M. Kjaer1,2 1
Department of Orthopaedic Surgery M, Institute of Sports Medicine Copenhagen, Bispebjerg Hospital, University of Copenhagen, Copenhagen, Denmark, 2Centre for Healthy Ageing, Faculty of Health Sciences, University of Copenhagen, Copenhagen, Denmark, 3 School of Health and Medical Sciences, O¨rebro University, O¨rebro, Sweden
Corresponding author: Abigail L. Mackey, Department of Orthopaedic Surgery M, Institute of Sports Medicine Copenhagen, Bispebjerg Hospital, University of Copenhagen, Building 8, Bispebjerg Bakke 23, 2400 Copenhagen NV, Denmark. Tel: 145 35 31 60 90, Fax: 145 35 31 27 33, E-mail:
[email protected] Accepted for publication 22 June 2010
There is strong evidence for enhanced numbers of satellite cells with heavy resistance training. The satellite cell response to very light muscle loading is, however, unknown. We, therefore, designed a 12-week training protocol where volunteers trained one leg with a high load (H) and the other leg with a light load (L). Twelve young healthy men [mean age 25 3 standard deviation (SD) years] volunteered for the study. Muscle biopsies were collected from the m. vastus lateralis of both legs before and after the training period and satellite cells were visualized by CD56 immunohistochem-
istry. A significant main effect of time was observed (Po0.001) for the number of CD561 cells per fiber (L: from 0.11 0.02 to 0.13 0.03; H: from 0.12 0.03 to 0.15 0.05, mean SD). The finding that 12 weeks of training skeletal muscle even with very light loads can induce an increase in the number of satellite cells reveals a new aspect of myogenic precursor cell activation and suggests that satellite cells may play a role in skeletal muscle adaptation over a broad physiological range.
Skeletal muscle hypertrophy is a prime example of tissue plasticity. Expansion of myofibers increases the demand on myonuclei, and according to the myonuclear domain theory, the volume of cytoplasm that each myonucleus can maintain is limited (Cheek, 1985; Allen et al., 1999). It is traditionally thought that further myofiber expansion requires the addition of new myonuclei. The satellite cell pool is considered a renewable source of these new myonuclei (Moss & Leblond, 1971; Rosenblatt et al., 1994). While the other primary role of satellite cells in repairing injured skeletal muscle is well recognized (Grounds et al., 2002), it is not known how satellite cells behave in a working muscle where hypertrophy and muscle injury are absent. It has been reported by many groups that muscle fiber hypertrophy, induced by periods of regular heavy resistance training, is accompanied by increased numbers of satellite cells in young and old men and women (Kadi & Thornell, 2000; Roth et al., 2001; Kadi et al., 2004b; Olsen et al., 2006; Petrella et al., 2006; Mackey et al., 2007; Verney et al., 2008, 2009), although training occasionally fails to elicit a change in satellite cell number in some groups of individuals (Hikida et al., 1998; Petrella et al., 2006). Endurance training has also been documented to
result in increased numbers of satellite cells in elderly men (Charifi et al., 2003; Verney et al., 2008). While it is difficult to compare these studies directly due to dissimilarities in the modes of training as well as gender and age of the participants, it does not appear that there is a strict relationship between the extent of muscle hypertrophy and the magnitude of expansion of the satellite cell pool under these various loading conditions. Furthermore, despite extensive research into the response of satellite cells to different stimuli, the question remains as to whether it is the load placed on the muscle or simply the amount of work performed that is the key determinant for enhancement of the satellite cell pool. We considered this an important question to address, not only to gain further insight into the physiological role of satellite cells but also in the context of conditions where it is not possible to perform heavy resistance training. Heavy resistance training may not be advisable or physically possible in the very early stages of recovery following periods of bed rest due to illness or surgery, particularly in elderly or very frail individuals. In this context, it would be interesting to know if a period of light muscle loading is sufficient to stimulate the satellite cell pool, which, it could be hypothesized, could prepare the muscle for a faster
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Mackey et al. adaptation to heavier resistance training. In order to investigate this, we examined the effects of resistance training with very light loading intensity on satellite cell number in young healthy men. Volunteers performed light training with one leg and as a control, trained the other leg with heavy loading intensity. The two different modes of training were designed such that one involved heavy loads with low numbers of repetitions, while the other involved very light loads but higher repetitions, adjusted so that the total amount of work lifted by both legs was equal. We hypothesized that, in contrast with the heavy resistance training, the light loading protocol would not invoke an expansion of the satellite cell pool. Muscle architecture and morphology were also assessed before and after both modes of training in order to evaluate the extent of muscle remodeling with this form of training and how these changes were associated with changes in satellite cell activity. Materials and methods Subjects and training Twelve young healthy men [mean age 25 3 standard deviation (SD) years; height 1.83 0.06 m; weight 81 14 kg] volunteered for the study, which involved training three times a week for 12 weeks, with the collection of muscle biopsies before and after the training period. All participants gave written informed consent for the study, which was approved by the Ethics Committees of the Municipalities of Copenhagen and Frederiksberg (Ref.: KF 01-171/04) and conformed to the Declaration of Helsinki. An analysis of the activation status of satellite cells in the biopsies obtained during this study is presented as a method validation component in a recent publication (Mackey et al., 2009). Data relating to the myosin heavy chain composition of the muscle biopsies collected during this study as well as results detailing the training adaptations of this study have recently been published, where a detailed description of the training protocol can also be found (Holm et al., 2008). Briefly, unilateral leg extensions were performed three times a week for 12 weeks, with one leg working with a heavy load (H) and the other leg with a lighter load (L). The number of repetitions was adjusted such that the work lifted by the two legs was equal. H was thus calculated to be 10 sets of eight repetitions at 70% of 1-repetition maximum (1 RM), and L was 10 sets of 36 repetitions at 15.5% 1 RM. 1 RM was determined before the 10th, 20th and 30th training sessions and the loads adjusted accordingly.
Muscle fiber pennation angle Pennation angle was measured from sagittal ultrasound images obtained from the mid portion of the vastus lateralis (VL) muscle (Aagaard et al., 2001), using a Siemens Sonoline Sienna scanner (SiemensAG, Erlangen, Germany) with a 7.5 MHz transducer. The subjects were seated in a rigid chair with relaxed leg muscles and the knee flexed to 901. VL fiber pennation angle (yp) was measured as the angle between VL muscle fiber fascicles and the deep aponeurosis of insertion. Five ultrasound images were obtained of each leg at the preand posttraining test sessions. Using a computer software (Scion Image for Windows beta 4.0.2, Scion Corporation,
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Frederick, Maryland, USA), y of the VL was analyzed and the yp was defined manually. To reduce the variation in the yp determination, the pennation angle of eight to 10 fascicles was determined from each of the five recorded images and the mean of these was calculated.
Muscle biopsies Muscle biopsies were obtained from the mid-portion of m. vastus lateralis using a standard needle biopsy technique. The posttraining biopsy was collected 3 days after the last training session. On extraction of the sample, the muscle fibers were aligned, embedded in Tissue-Tek (Sakura Finetek Europe, Zoeterwoude, the Netherlands) and frozen by immersion in isopentane, precooled to approximately 160 1C by liquid nitrogen. Samples were stored at 80 1C pending analyses. Before sectioning, the biopsies were coded by randomly assigning a unique identification number from 1 to 48 to each biopsy such that all samples were blinded with regard to subject, time point and leg. Serial transverse sections (10 mm) were cut at 24 1C using a cryostat and mounted on SuperFrost Plus slides (Menzel-Gla¨ser, Braunschweig, Germany). All subsequent analyses were performed blinded.
Fiber type and size Routine ATPase histochemistry was carried out with preincubation at pH 4.37, 4.60 and 10.30 (Brooke & Kaiser, 1970), and five different fiber types were identified: types I, I/IIa, IIa, IIax and IIx (Andersen & Aagaard, 2000). As described previously in detail (Andersen & Aagaard, 2000), images of the serial sections of the various ATPase stainings were analyzed for proportion of fiber type and fiber area, using a TEMA image analysis system (TEMA, Hadsund, Denmark). For the 48 biopsies analyzed, a mean of 191 41 SD fibers was included in the ATPase analysis.
Capillaries Immunohistochemical staining was carried out to visualize capillaries, as described in detail elsewhere (Qu et al., 1997), using a combination of the primary antibodies, Ulex europaeus agglutinin I Lectin (cat. no. B0279, Dako Norden A/S, Glostrup, Denmark) and collagen type IV (cat. no. M0785, Dako Norden A/S). The secondary biotinylated goat antirabbit (cat. no. E0432, Dako Norden A/S) and goat antimouse (cat. no. E0433, Dako Norden A/S) antibodies were applied subsequently, followed by avidin-biotinylated alkaline phosphatase (ABC-AP, cat. no. K376, Dako Norden A/S). Fuchsin substrate (cat. no. K624, Dako Norden A/S) reacted with the alkaline phosphatase to stain capillaries red. The capillary-to-fiber ratio (Cap f 1) and number of capillaries around each fiber type (CAF) were determined in conjunction with the TEMA-assisted fiber-type analysis. A mean of 390 87 SD capillaries per biopsy was included in this analysis.
Satellite cells Satellite cell counting was facilitated by double-immunofluorescence staining with CD56 and laminin antibodies, as described and validated recently (Lindstro¨m & Thornell, 2009; Mackey et al., 2009). CD56 has been used successfully for the identification of satellite cells on frozen sections of human skeletal muscle in many studies in the literature (Kadi et al., 1999, 2004b; Kadi & Thornell, 2000; Olsen et al., 2006; Petrella
Increased CD56+ cells with low muscle loading
Fig. 1. Example of double immunofluorescent staining for identification of satellite cells using the CD56 antibody (red). Laminin staining (green) delineates the fibers borders (and capillaries) and thus confirms the location of the satellite cell inside (series 1) or outside (series 2 and 3) the fiber basement membrane. Nuclei appear blue. In series 1, three CD561 cells are visible, situated inside the basement membrane. These cells are considered satellite cells and were included in the assessment of satellite cell number in this study. Series 2 and 3 display examples of rare CD56+ cells (red) not situated in the traditional satellite cell position within the confines of the muscle basement membrane (laminin, green). These ‘‘outside’’ cells were excluded from satellite cell counts in this study. Scale bars are 50 mm. et al., 2006; Christov et al., 2007; Mackey et al., 2007, 2009; Murphy et al., 2008; Verney et al., 2008; Lindstro¨m & Thornell, 2009). Briefly, satellite cells were identified with a mouse antiCD56 antibody (CD56, cat. no. 347740; Becton Dickinson, San Jose, California, USA), followed by Alexa Fluor 568 goat antimouse secondary antibody (Molecular Probes cat. no. A11031; Invitrogen A/S, Taastrup, Denmark). The location of the CD561 cells with respect to the basement membrane was determined by subsequent staining of the same sections with a rabbit antilaminin antibody (cat. no. Z0098; Dako Norden A/S) and Alexa Fluor 488 goat anti-rabbit secondary antibody (Molecular Probes cat. no. A11034; Invitrogen A/S), while 40 ,6-diamidino-2-phenylindole (DAPI) in the mounting medium (Molecular Probes ProLong Gold antifade reagent, cat. no. P36935; Invitrogen A/ S) stained the nuclei. This method rendered CD56 staining red, laminin green and nuclei blue (Fig. 1). Counting of CD561 cells was carried out using a 40 objective on 446 219 fibers (mean SD), in line with previous recommendations (Mackey et al., 2009). The same person carried out all counting. Values were expressed in three ways: (1) relative to the number of sublaminar nuclei [(CD561 cells/sublaminar nuclear number) 100], (2) relative to fiber number and (3) relative to the mean fiber crosssectional area (CSA) [(CD561 cells/fiber)/(mean fiber area of all
fiber types, as measured from ATPase analysis)]. Care was taken to only include the CD561 cells located inside the myofiber basement membrane in this assessment. The presence of CD561 cells located outside the basement membrane was recorded separately.
Myonuclei The number of sublaminar nuclei was assessed from sections stained for merosin (Clone Mer3/22B2; Novocastra, Newcastle upon Tyne, UK), a muscle-specific laminin located in the basement membrane (37). Alexa Fluor 568 goat antimouse secondary antibody (Molecular Probes cat. no. A11031; Invitrogen A/S) was used for visualization of antibody binding, while DAPI in the mounting medium (Molecular Probes ProLong Gold antifade reagent, cat. no. P36935; Invitrogen A/S) stained the nuclei.
Myofiber remodeling Using the same double-immunofluorescence staining procedure described above for the identification of satellite cells, myofiber regeneration was assessed from sections double
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Mackey et al. stained for embryonic myosin (F1.652, Developmental Studies Hybridoma Bank, Iowa City, Iowa, USA) and dystrophin (cat. no. ab15277; Abcam, Cambridge, UK). The number of fibers with centrally located nuclei was also determined from these sections. The number of fibers demonstrating cytoplasmic immunoreactivity for CD56, an indication of myofiber remodeling (Winter & Bornemann, 1999), was recorded from sections stained for the assessment of satellite cell number. Myogenin (F5D, Developmental Studies Hybridoma Bank) staining, in combination with a rabbit anti-laminin (cat. no. Z0098; Dako Norden A/S) antibody, was carried out in order to investigate the extent of satellite cell differentiation.
Table 2. The number of capillaries around each fiber type (CAF) and the capillary-to-fiber ratio (Cap f 1), as determined from muscle biopsy cross sections before and after a 12-week period of heavy (H) or light (L) intensity-resistance training
Fiber type
Training mode
Pre
Post
Cap f 1
H L H L H L H L
1.97 0.38 2.05 0.29 4.70 0.66 4.90 0.46 4.56 0.88 4.72 0.73 4.11 0.63 4.22 1.28
2.18 0.49 2.12 0.50 5.19 0.75 4.84 0.96 4.95 1.14 4.47 0.82 4.75 0.96 4.14 1.14
CAF Type I* CAF Type IIa* CAF Type IIx
Statistics Statistical analyses were performed using SigmaPlot version 11 for Windows. Statistical significance was accepted at the 0.05 level. Data are presented as means SDs. Two-way repeated measures ANOVA was carried out on all variables, and where a time training interaction was observed, post hoc pairwise multiple comparisons (Student–Newman–Keuls Method) were performed. Where data failed the normality test (Shapiro–Wilk), one-tailed Wilcoxon signed rank tests were carried out on relative changes to normalize the data for comparisons of the effect of mode of training.
Results Muscle fiber pennation angle Two-way repeated measures ANOVA analysis of VL pennation angle revealed a tendency for a main effect of time (P 5 0.055), but not training or time training interaction. Pennation angle values were observed to increase numerically in eight out of 12 subjects with the H loading protocol (mean group increase of 11%, from 13.91 2.3 to 15.11 2.7), and in nine out of 12 following L training (mean group increase of 6%, from 13.51 1.8 to 14.21 1.9). Fiber type and size Because the number of intermediate fibers (I/IIa and IIa/IIx) was so small, these fibers were distributed among the three main categories (I, IIa and IIx) by
Values are mean SD, n 5 12. *P 0.1, time training interaction.
assigning half of the number to the adjacent slow type and the other half to the adjacent fast fiber-type group, as described previously (Andersen & Aagaard, 2000). The data presented in Table 1 display the relative proportion of each fiber type and the mean area of each fiber-type pre- and posttraining. No statistically significant changes were detected for fiber area. A significant training time interaction (P 5 0.007, two-way repeated measures ANOVA) was observed for the percentage of type IIx fibers, decreasing significantly with H but not with L training (P-values of 0.046 and 0.498, respectively; Student–Newman–Keuls pairwise comparisons). Capillaries The mean number of capillaries around each fiber type (CAF) and the capillary-to-fiber ratio (Cap f 1) data are presented in Table 2. CAF around types I and IIa, but not IIx, fibers demonstrated a tendency for a training time interaction (P-values 0.104 and 0.081, respectively; two-way repeated measures ANOVA). Satellite cells and myonuclei
Table 1. The relative proportion and area of types I, IIa and IIx fiber types in muscle biopsy cross sections before and after a 12-week period of heavy (H) or light (L) intensity-resistance training
Fiber type
Training mode
Pre
Post
Type I (%)
H L H L H L H L H L
55 12 60 14 29 10 30 13 16 17 97 5390 1202 5195 1299 6136 1527 5821 1810
59 10 58 11 33 9 31 9 8 4* 12 8 5265 1504 5264 1722 6100 2635 5542 1979
Type IIa (%) Type IIx (%) Type I (mm2) Type II (mm2)
Values are mean SD, n 5 12. *Po0.05, pairwise comparison compared with baseline.
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Satellite cell data are presented in Fig. 2, expressed relative to the number of sublaminar nuclei, the number of fibers, or relative to the mean fiber area. For all three variables, the two-way repeated measures ANOVA detected a significant effect of time (P 0.005), but not training mode or time training interaction. The software program G*Power was used to carry out post hoc power calculations based on these data and revealed that, at an a level of 0.05, inclusion of 12 subjects resulted in a power level (1 b) of 0.99 with a large effect size (dz) of 1.08 for H, and a power level of 0.64 with a moderate effect size of 0.49 for L. Because all satellite cell data failed the normality test, the relative changes with H and L training were compared using a Wilcoxon test, which
Increased CD56+ cells with low muscle loading Table 3. The number of myonuclei per fiber (mn f 1) and myonuclear domain (mnd) size in muscle biopsy cross sections before and after a 12week period of heavy (H) or light (L) intensity-resistance training
mn f 1 mnd (mm2)
Training mode
Pre
Post
H L H L
2.8 0.6 2.8 0.4 2067 234 1963 314
2.8 0.6 2.8 0.7 2024 419 1928 527
Values are mean SD, n 5 12.
Occasional CD561 cells located outside the basement membrane, i.e. in the interstitial space between the muscle fibers, were observed. Two examples of this are shown in Fig. 1. However, these cells were rare – at baseline an area of a mean (n 5 28) of 250 fibers was required to find one of these cells. Posttraining, this value was 300 fibers for H and 177 fibers for L. Presented in Table 3 are the number of myonuclei per fiber and myonuclear domain size, demonstrating similar numbers before and after training for both modes of training. Differentiation of satellite cells Immunohistochemical staining for myogenin was performed in an attempt to follow differentiation of satellite cells with training. Given the low number of F5D1 cells, the values were expressed per 100 fibers [(F5D1 cells/fiber no.) 100]. The number of F5D1 nuclei expressed per 100 fibers did not change significantly. Positive cells were observed in both groups at baseline (H: 0.20 0.27; L: 0.14 0.13 cells per 100 fibers) and posttraining (H: 0.17 0.28; L: 0.31 0.58 cells per 100 fibers). Myofiber remodeling
Fig. 2. The influence of heavy (white bars) and light (black bars) resistance training on the number of satellite cells (CD561) in the vastus lateralis of young healthy men. The data are presented in three different ways, expressed relative to the number of myonuclei, the number of fibers, or the mean fiber cross-sectional area. Means SD. *Main effect of time, P 0.005, 2-way repeated measures ANOVA.
showed that the relative increase in satellite cell number posttraining in the two legs was similar, for the proportion of CD561 cells (H: 31 32%; L: 18 32%; P 5 0.117), CD561 cells/fiber (H: 30 24%; L: 18 19%; P 5 0.151) and CD561 cells per mm2 (H: 37 39%; L: 23 33%; P 5 0.190).
The presence of fibers immunoreactive for embryonic myosin was only observed in two biopsies at baseline and two biopsies posttraining (not the same individuals as those identified at baseline), both in the L group, where the proportion of positive fibers accounted for from 0.1 to 0.8% of the total number of fibers in the section. A total of 13 embryonic myosin1fibers were detected, 12 of which demonstrated positive cytoplasmic staining for CD56 (identified on a subsequent section stained with CD56). The number of fibers demonstrating positive staining for CD56 was evaluated as a marker of myofiber remodeling (Winter & Bornemann, 1999). Twenty-two of the 28 biopsies at baseline contained fiber cross sections that reacted positively with the CD56 antibody used in this study, accounting for a mean of 2.4% ( 2.9) of fibers in H and 1.3% ( 2.1) in L. Only one biopsy failed to show any positive fibers posttraining (H: 1.2 1.0%; L: 2.5 4.7%).
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Mackey et al. Centrally located nuclei Centrally located nuclei were evident in half of the biopsies, present in H in a mean of 2.3% ( 5.0) of fibers at baseline and 0.1% ( 0.2) posttraining. Values in L were 0.4% ( 0.4) at baseline and 0.5% ( 0.5) posttraining. Statistical analyses did not reveal any significant effect of time, training or training time interaction. Of the total number of 154 fibers observed with internal nuclei in all of the biopsies assessed, 51 of these fibers demonstrated positive cytoplasmic staining for CD56.
Discussion Numerous studies have demonstrated enhanced satellite cell numbers in human skeletal muscle following periods of heavy resistance training. Our results support these previous reports. The novel finding of the present study is an 18% increase in satellite cell (CD561) number with 12 weeks of light intensity (15.5% 1 RM) resistance training. This was in contrast to our hypothesis and contributes further to our knowledge about the stimulus required to enhance the satellite cell pool. Satellite cells The aim of this study was to investigate if very light load-resistance training could enhance the satellite cell pool. This experiment is unique in that we have compared this with heavy load-resistance training, where both modes of training were matched for the total amount of lifted load performed, and each subject performed both modes of training. To our knowledge, this is the first time that the response of satellite cells to such light training has been investigated in young healthy individuals. A possibility that has not hitherto been addressed is that satellite cells are activated simply in response to work performed by the muscle, even under conditions of very low intensity loading. Most of the prior studies investigating the effects of long-term exercise training on satellite cells have been based on periods of heavy resistance training (Kadi & Ponsot, 2010). Two exceptions to this, and possibly the most suitable comparison for the low work performed by the subjects in the present study, are endurance training studies where 14 weeks of training on a bicycle ergometer resulted in increased numbers of satellite cells in the m. vastus lateralis (VL) of elderly men (Charifi et al., 2003; Verney et al., 2008). Significant hypertrophy of type IIa fibers was, however, also observed in these volunteers, which may have driven the increase in satellite cell number. With regard to the H and L resistance training in the present study, we failed to detect any statistically significant inter-
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action between training mode and change in satellite cell number, suggesting that the satellite cell pool responded similarly to both types of training. This is supported by the satellite cell data of Verney et al. (2008), who were unable to detect a difference in response between heavy resistance training and endurance training. Taken together, our finding of enhanced satellite cell number with L intensity resistance training provides new insight into potential stimuli involved in expansion of this resident myogenic precursor cell pool. The presence of occasional CD561 cells or small CD561 fibers outside the muscle basement membrane has been reported previously (Kadi et al., 1999; Doppler et al., 2008; Lindstro¨m & Thornell, 2009), the distinction between the two being difficult if the fibers are very small. In the case of small fibers expressing CD56, this is likely to be an indication of lack of innervation (Winter & Bornemann, 1999), while CD56 expressed on the membrane of a single cell outside the basement membrane could either be a migrating satellite cell or another cell type expressing this marker, such as natural killer cell lymphocytes (Lanier et al., 1989). While these cells were very rare in the biopsies examined in this study, staining for CD56 in combination with a basement membrane marker provides an extra control for not including these cells in the satellite cell counts. Muscle architecture The occurrence of hypertrophy and the associated demand for extra myonuclei would be an obvious explanation for the presence of the increased numbers of satellite cells observed in the present study. Hypertrophy was investigated in two ways – by magnetic resonance imaging (MRI) of the whole quadriceps muscle group and by ATPase histochemical analysis of cross sections of biopsies collected from the VL portion of the quadriceps. These two methods produced different outcomes in the assessment of hypertrophy in our study participants. The MRI data, published elsewhere (Holm et al., 2008), revealed significant increases in quadriceps CSA with both forms of training, although the increase with H training (8%) was statistically greater than the 3% observed with L training (Holm et al., 2008). In contrast to this, mean fiber area (MFA), as assessed from ATPase analysis, did not uncover any evidence of hypertrophy in either group. This discrepancy has been reported in other studies (Holm et al., 2006; Mackey et al., 2007; Verney et al., 2008), and while it is likely to be due to a combination of factors, some possible explanations can be put forward. MRI analysis in this study evaluated the CSA of all of the quadriceps muscles, in contrast to ATPase analysis, which calculated MFA of o200 fibers, sampled
Increased CD56+ cells with low muscle loading from a small part of the VL alone. With regard to muscle architecture, pennation angle displayed a tendency toward a training time interaction (P 5 0.055), with numerical increases observed in eight out 12 for H and nine out of 12 for L, suggesting that the architecture of the VL underwent changes in this study. We cannot rule out the possibility that a remodeling of the muscle architecture would be sufficient to induce a satellite cell response. Alternatively, it is interesting to consider the hypothesis that expansion of the satellite cell pool precedes muscle fiber hypertrophy and represents an attempt to renew and maintain the satellite cell pool, or that satellite cells respond sensitively to any form of work performed by the muscle and that expansion of the satellite cell pool represents a basic physiological adaptation to exercise.
Fate of the new satellite cells Our data on myonuclear number and myogenin staining suggest that the training stimulus was not enough to induce terminal differentiation of satellite cells and ultimately the incorporation of the new satellite cells into the existing myofibers. This is in line with the documentation that significant increases in myonuclear number have only been reported where 426% hypertrophy occurs (Kadi et al., 2004a, b), or where the myonuclear domain expands beyond 2000 mm2 (Petrella et al., 2006). The myonuclear domain size of the biopsies we collected from participants in the present study showed no change with either mode of training and was close to 2000 mm2 for both legs, before and after the training period. Thus, it does not appear that demand for new myonuclei was a factor in the observed increase in satellite cell number in the present study. Similarly, no change was detected in the number of myogenin1 cells with training. Myogenin is a myoregulatory factor involved in cell differentiation and it has been reported previously that the immunohistochemical staining pattern of myogenin in human skeletal muscle is altered in response to acute exercise (Kadi et al., 2004a). Our hypothesis was that an increased myogenin expression would indicate the differentiation of the new satellite cells, although it can be argued that the likelihood of detecting such a change in a single biopsy after 12 weeks of training is small. Furthermore, a limitation of our detection method was the double labeling with laminin and not CD56; hence, we cannot be certain that the stained nuclei are satellite cells and not myonuclei (Kadi et al., 2004a). The lack of change in the number of myogenin1 cells observed in this study although suggests either that the training loads were not sufficient to induce differentiation of satellite cells to a detectable
level at 12 weeks, or that the new satellite cells detected are programmed to remain as satellite cells. Muscle regeneration As well as in response to muscle hypertrophy, satellite cells are known to be important for the repair of damaged fibers or damaged segments of fibers in healthy postnatal skeletal muscle (Grounds et al., 2002; Zammit et al., 2006). Myofiber remodeling was evaluated qualitatively in the present study from immunohistochemical stainings for embryonic myosin and CD56, and the presence of centrally located myonuclei. Embryonic myosin-positive fibers were, however, only present in two of the posttraining biopsies, and although the number of fibers demonstrating CD56 immunoreactivity was higher than those expressing embryonic myosin, there was no difference in the percentage of CD561 fibers when compared with baseline. Examination of the presence of centrally located myonuclei did not reveal any change with training. We therefore cannot conclude that the role of the new satellite cells, visible at 12 weeks, was to repair training-induced damage. Furthermore, it is unlikely that the L training would have induced muscle damage even in the earlier stages of the training period of this study, supporting a broader physiological role for satellite cells in adaptation to exercise. Satellite cell niche One potentially potent factor in activating satellite cells is the response of the surrounding satellite cell niche to the stress placed on the muscle. Besides the parent myofiber, the main component of the satellite cell niche is the surrounding basement membrane, to which it is anchored and through which it can receive mechanical and chemical signals (Kuang et al., 2008). Recently, using a single bout of training identical to the training sessions used in the current study, Holm et al. (2010) showed that the fractional synthesis rate of intramuscular collagen protein was upregulated following both the H and the L modes of exercise (Holm et al., 2010). Therefore, it can be speculated that remodeling of the collagen component of the satellite cell niche may contribute to the observed enhancement in satellite cell content in this study in response to both modes of training. A third element of the satellite cell niche is comprised of neighboring capillaries and vascular endothelial cells (Christov et al., 2007; Kuang et al., 2008; Abou-Khalil et al., 2010) (see Fig. 1 for illustration of the proximity of satellite cells to capillaries). We investigated capillarization of the biopsies collected before and after the training period. Looking at the distribution of capillaries among the individual fiber types, a tendency
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Mackey et al. was observed for a training time interaction regarding the mean number of capillaries (CAF) around type I and type II fibers (Table 2), suggesting that the H training may have resulted in a greater increase in capillarity. Incidentally, satellite cells were counted in the present study on whole muscle cross sections and not with respect to fiber type, a recent development in satellite cell methodology (Verney et al., 2008; Verdijk et al., 2009), and we therefore cannot exclude the possibility that this distinction between satellite cell content of type I and II fibers, as was performed for capillaries, would have uncovered a more nuanced insight into the response of satellite cells in our study. Increases in skeletal muscle capillarization with heavy resistance training or endurance training are well recognized (Andersen & Henriksson, 1977; Schantz et al., 1983; Kadi et al., 1999; Christov et al., 2007) and our tentative findings of enhanced capillarity with H training are in line with these studies. However, the lack of change with L training may be due to the very low intensity and nonexhausting nature of this protocol. Unlike highintensity endurance training, the participants in the present study did not display signs of cardiovascular or local muscle fatigue on completion of the L exercise session. While we cannot exclude the role of endothelial cell-derived factors in satellite cell activation (Abou-Khalil et al., 2010) with our H training protocol, the apparent lack of change in microvascularization with L training in this study does not support this connection. What potential stimuli could then be involved in the observed activation of satellite cells with this type of exercise? Potential stimuli One of the features of satellite cells is that they are self-renewing. For this to happen, they have to leave their quiescent state to reenter the cell cycle and divide. However, it has been documented that very few satellite cells (not 41%) are in the active phase of the cell cycle in muscles of young healthy adults (Mackey et al., 2009), and it is not known what factors are likely to activate them in the absence of muscle disease, injury or strong hypertrophy-inducing stimuli. In light of the recognition of muscle satellite cells as being a heterogeneous population (Conboy & Rando, 2002; Zammit et al., 2004; Kuang et al., 2008), it is possible that weaker physiological stimuli, such as the low loading protocol used in the present study, exclusively activate a subpopulation of satellite cells who only generate self-renewing daughter cells and thus renew and replenish the satellite cell pool. With regard to potential molecular stimuli, our findings on L training at 15.5% of 1 RM are supported at the mRNA level by a recent report on the effect of a single bout of low intensity (20% 1 RM)
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resistance exercise, where increased levels of p21 and MyoD were observed in the hours after exercise (Drummond et al., 2008). Together with a concomitant downregulation of myostatin (Drummond et al., 2008), these results are suggestive of a window of enhanced myogenic activity, coordinated with an attenuation of a potent negative regulator of muscle growth and satellite cell activity, at least in the hours immediately following exercise. While it is not known if this response occurs to the same extent after every training session, it is possible that these factors could be responsible for the enhanced satellite cell number observed in our study with L training. However, further investigations are required to substantiate this connection and to elucidate the mechanisms by which the satellite cell pool is maintained and renewed in healthy adult skeletal muscle in the absence of disease, injury and growth. Perspectives Investigations into the response of the satellite cell pool to exercise have mostly used exercise models involving heavy muscle loading; hence, the finding that 12 weeks of training of the quadriceps muscles by repeated muscle contraction at very low intensity can induce an increase in the number of satellite cells reveals a new aspect of myogenic precursor cell activation. It is unclear from this study whether enhancement of the satellite cell pool was initiated by imminent hypertrophy, by remodeling of the satellite cell niche, or simply in response to the work performed by the muscle. While the functional consequences of the activation remain to be determined for the muscle, it is possible that this knowledge could be useful when considering training regimens where the goal is simply to amplify satellite cell number, or where heavy resistance training is not possible, such as for very frail individuals in the early stages following operation. From a physiological perspective, these data suggest that the role of the satellite cell in adaptation of skeletal muscle to changes in loading could be broadened from their traditional functions in muscle hypertrophy and repair to include taking the opportunity to maintain and renew the satellite cell pool upon registration of weaker physiological stimuli. Furthermore, these data support the hypothesis that expansion of the satellite cell pool represents a basic physiological adaptation to exercise. Key words: neural cell adhesion molecule, satellite cell renewal, hypertrophy, low muscle loading.
Acknowledgements Funding is gratefully acknowledged from the Danish Medical Research Council, Lundbeck Foundation, Novo Nordisk Foundation, Danish H:S and Nordea Foundation (Healthy
Increased CD56+ cells with low muscle loading Ageing grant). The F1.652 monoclonal antibody developed by Helen M. Blau and the F5D monoclonal antibody developed by Woodring E. Wright were obtained from the Develop-
mental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by Department of Biological Sciences, The University of Iowa, Iowa City, Iowa, USA.
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