Experimental approach

Throughout this investigation, satellite cells and their progeny were characterized in a variety of culture systems. Typically, each donor mouse was used for several culture systems. Muscles from young or adult mice were always processed in parallel with muscles from old or senile mice.

The C57BL/6 mouse strain was selected due to its long lifespan (Storer, 1966), a prerequisite for measuring gradual age-dependent changes. The age range chosen for this study corresponds with changes in muscle fiber maturation and life span in this strain. Specifically, the youngest age group (3–6 months) aligns with the time of postnatal EDL muscle maturation (Agbulut et al., 2003); the oldest age group (29–33 months) represents the upper limit of this strain’s life span (Goodrick, 1975; Kunstyr and Leuenberger, 1975; Mewissen, 1971). Since age-related attrition can depend on muscle activity, the high level of spontaneous activity exhibited by C57BL/6 mice (Nikulina et al., 1991) is particularly advantageous in studying the consequences of aging on muscle deterioration.

Growth media

Myofibers from EDL and soleus muscles, as well as primary myogenic cultures, were maintained in rich medium that supports both proliferation and differentiation of satellite cell progeny (Yablonka-Reuveni, 2004). This medium consisted of DMEM (Dulbeco’s Modified Eagle Medium [high glucose, with L-glutamine, 110 mg/L sodium pyruvate, and piridoxine hydrochloride] supplemented with 50U/ml penicillin and 50 mg/ml streptomycin; GIBCO-Invitrogen) containing 20% fetal bovine serum (Sigma-Aldrich), 10% horse serum (HyClone), 1% chicken embryo extract prepared from whole 10-day old embryos (Shefer and Yablonka-Reuveni, 2005; Yablonka-Reuveni, 2004).

Myofibers from flexor digitorum brevis (FDB) muscle were maintained in a basal medium (Shefer and Yablonka-Reuveni, 2005; Yablonka-Reuveni and Rivera, 1994; Yablonka-Reuveni et al., 1999a). The basal medium consisted of DMEM, 20% Controlled Process Serum Replacement (CPSR-2, bovine plasma treated with charcoal followed by dialysis, a process which depletes growth promoting factors; Sigma-Aldrich) and 1% horse serum with or without 2 ng/ml FGF2 (human recombinant; provided by Dr. S. Hauschka, University of Washington).

Isolation and culturing of intact myofibers from EDL and soleus muscles

EDL and soleus muscles were digested with collagenase for 60 and 90 minutes, respectively, and released myofibers were cultured individually in Matrigel-coated wells using rich growth medium as previously described (Shefer and Yablonka-Reuveni, 2005; Shefer et al., 2004). Typically, a preparation from a single mouse yielded four to six parallel 24-well tissue culture dishes. Due to the presence of atrophying myofibers in muscles from older mice, more myofibers were released from older animals to achieve the same number of intact myofibers as in younger animals. To detect satellite cells in their in-situ position, EDL and soleus myofibers were allowed to adhere for 3 hours followed by fixation (i.e., freshly isolated myofibers). EDL myofibers were also cultured for either 1 or 2 weeks to study long-term myogenesis. Viable myofibers may influence the performance of satellite cells during early days in culture; because of this, only cultures that contained intact myofibers by culture day 3 were further followed. Clonal cultures were derived from ten to fifteen EDL myofibers per mouse (Shefer et al., 2004). Clonal size was determined as previously described based on the number of microscopic fields occupied by the clones using a 10x objective (Yablonka-Reuveni and Rivera, 1997a). Clones were categorized as small (about 0.75 to 1 field), medium (about 1.5 to 2 fields) and large (more then 2.5 fields).

Isolation and culturing of intact FDB myofibers

To follow the initial steps of satellite cell myogenesis within the myofiber unit, myofiber cultures were prepared from the FDB muscles as previously described (Shefer and Yablonka-Reuveni, 2005; Yablonka-Reuveni et al., 1999a). Muscles were digested with collagenase for 2.5–3 hours and released myofibers were cultured in Vitrogen-coated culture dishes. Typically, a preparation of one mouse resulted in five to six 35-mm tissue culture dishes, with 20–30 myofibers per dish. Myofiber cultures were maintained in basal medium with or without FGF2; medium was replaced every 24 hours to ensure ample supply of FGF2.

Primary myogenic cultures

Cells were dissociated from the fast-twitch tibialis anterior and gastrocnemius muscles using Pronase, following our procedure for adult rodents (Kastner et al., 2000; Yablonka-Reuveni, 2004). Cells were cultured in 24-well dishes (Matrigel pre-coated, as in EDL myofiber cultures) at a density of 5×10³ cells per well. Typically, a preparation from a single mouse was cultured into 12 wells per dish (5–6 dishes, one dish per time point), which permitted multiple parallel cultures per experiment. This protocol yields cultures containing 85–90% myogenic cells, based on the expression of early myogenic markers, such as Pax7 or MyoD, on culture day 4 (Yablonka-Reuveni, 2004; present study).

Immunofluorescent analysis

Freshly isolated and cultured EDL and soleus myofibers, and primary myogenic cultures were fixed with 4% paraformaldehyde followed by immunolabeling and counterlabeling of nuceli with DAPI as previously described (Shefer et al., 2004; Shefer and Yablonka-Reuveni, 2005; Yablonka-Reuveni, 2004). Some cultures were analyzed by single- and double-immunofluorescent staining using mouse, rat and rabbit primary antibodies followed by species-specific secondary antibodies. Other cultures were double-labeled with isotype-specific mouse primary antibodies and reactivity was monitored with isotoype-specific secondary antibodies (Shefer and Yablonka-Reuveni, 2005; Yablonka-Reuveni, 2004). In addition, primary cultures were triple-labeled with two isotype-specific mouse antibodies and a rabbit antibody. For double-immunolabeling, secondary antibodies were AlexaFluor 488 and 568. For triple-immunolabeling, secondary antibodies were AlexaFluor 488, 568 and 647. In order to avoid possible false positive signals due to some overlap in emission spectra of the 568 and 647 chromophores, the latter two were applied together only when demarcating antigens localized to different cellular compartments (i.e., nuclear and cytoplasmic). All secondary antibodies (Molecular Probes) were produced in goat and were used at a 1:1000 dilution. Controls consisted of cultures reacted with one primary antibody followed by 2 or 3 secondary antibodies and of cultures reacted with just secondary antibodies.

FDB myofibers were processed as above except that they were fixed with methanol (Shefer and Yablonka-Reuveni, 2005; Yablonka-Reuveni and Rivera, 1994; Yablonka-Reuveni et al., 1999a). Methanol, but not paraformaldehyde, was used in this set of studies since it maintains good adherence between the myofiber and the Vitrogen matrix throughout the immunostaining procedure.

Primary antibodies

Previously characterized primary antibodies were either monoclonal antibodies developed in mouse and rat, or polyclonal antibodies developed in rabbit. The following primary antibodies were used: anti-Pax7 (mouse IgG1, ascites fluid, Developmental Studies Hybridoma Bank [DSHB], 1:2000 dilution); anti-MyoD (mouse IgG1, clone 5.8A, BD Biosciences, 1:800; and rabbit, M-318, Santa Cruz Biotechnology, 1:400; double immunostaining with both anti-MyoD antibodies showed co-staining of the same cells); anti-myogenin (mouse IgG1, clone F5D, hybridoma supernatant, DSHB, 1:2); anti-β-galactosidase (mouse IgG2a, clone JIE7, supernatant, DSHB, 1:16); anti-sarcomeric myosin (mouse IgG2b, MF20, ascites fluid, DSHB, 1:6000); anti-CD34 (rat, BD Biosciences, 1:100); anti-PCNA (mouse IgG, clone 19F4, Boehringer Mannheim, 1:100); and anti-MAPK (ERK1/ERK2), rabbit, 1:1000; (Yablonka-Reuveni et al., 1999a; Yablonka-Reuveni et al., 1999b).

Cell Quantification

Satellite cells were detected in freshly isolated EDL and soleus myofibers by immunolabeling for Pax7. Cultured FDB myofibers were monitored for the number of myofiber-associated proliferating (PCNA⁺) and differentiating (myogenin⁺) satellite cells at different time points. Positive cells in freshly isolated and in cultured myofibers were identified and quantified by multi-focal level inspection of each myofiber using a 40x objective. In some cases, myofibers were double-labeled with the anti-MAPK antibody to confirm that satellite cells, and not the myofiber nuclei were positive for the nuclear antigens Pax7, PCNA or myogenin (Yablonka-Reuveni et al., 1999a; Yablonka-Reuveni et al., 1999b).

To quantify cells emanating from myofibers, 1- and 2-week old myofiber cultures were double-immunolabeled with mouse anti-Pax7 and rabbit anti-MyoD. Ten arbitrary microscopic-fields were analyzed with a 20x objective. Four images were acquired per field, using phase and fluorescent channels (red [Pax7], green [MyoD], blue [DAPI]). Since arbitrary fields in some of the cultures prepared from old and senile mice did not contain cells, additional images were acquired to accomplish a total of 10 cell-containing fields.

Primary myogenic cultures were quantified in a similar manner to cultures emanating from myofibers with the exception that in addition to double-labeling for Pax7 and MyoD, parallel cultures were triple-labeled with mouse anti-Pax7 (or anti-myogenin) in combination with rabbit anti-MyoD and mouse anti-sarcomeric myosin (red, green and far-red channels, respectively). During early time points, more then 10 arbitrary fields were analyzed to ensure a minimum number of 500 cells per quantification, and results were averaged per 10 fields.

Myogenesis in cultures emanating from EDL myofibers: number of satellite cell progeny, but not their differentiation potential, is reduced with age

Single myofibers were maintained in culture for 1- or 2-weeks. Cells emanating from individual myofibers (referred to below as “emanating cells”) were quantified, using double immunostaining, for the number of Pax7⁺/MyoD⁻, Pax7⁺/MyoD⁺, and Pax7⁻/MyoD⁺ mononucleated cells, and for the number of nuclei fused into myotubes (Fig. 3A–H and Fig. 4; each bar in Fig. 4 depicts the results of an individual myofiber culture). We analyzed cultures at the 1- and 2-week time points because we wished to determine both amplification and differentiation potential of the emanating cells. Satellite cell proliferation begins within the initial days in culture; further proliferation of progeny along with differentiation onset are detected by the first week in culture as shown in the present study and previous investigations (e.g., Zammit et al., 2002; Shefer et al., 2004). We also aimed at analyzing the progression of satellite cell progeny into the more advance phase of differentiation (i.e., fusion into myotubes) and for that we analyzed 2-week old cultures.

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Fig. 3

Immunofluorescent and phase images depicting myogenesis in cultures emanating from single EDL myofibers and in live clones of satellite cells that were dissociated from EDL myofibers. Myofibers from a young mouse were cultured for 1- or 2-weeks (A–D and E–H, respectively) and then double-labeled for Pax7 and MyoD. (A, E) Merged phase and DAPI images; (B, F) Pax7; (C, G) MyoD; (D, H) merged Pax7 and MyoD images demonstrating that all cells in the earlier time point co-express Pax7 and MyoD but at the later time point only residual cells are positive for both antigens. Clones were derived from EDL myofibers of young (I), old (J) and senile (K) mice. Bar, 100 μm.

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Fig. 4

Progression of satellite cell progeny through myogenesis in cultures emanating from individual EDL myofibers. Each stacked bar represents the distribution of cells within a single myofiber culture. The number of mice (n) and the number of myofibers (f) per age group is listed under the respective inset. Data collected by double-immunolabeling and DAPI-counterstaining are shown for 1- and 2-week old cultures (top and bottom panels, respectively). In the inset of each age group, individual myofiber cultures were ranked from left to right according to the total number of myogenic nuclei they encompass. The height of each bar represents the total number of myogenic cells in the corresponding myofiber culture and the different segments in each bar represent the number of cells in each of the four distinct myogenic compartments analyzed. Myofibers that did not give rise to cells are ranked at the left part of each inset. Additional analysis of the data is included in Table 2.

Data show that regardless of age, there was a significant increase in the number of emanating cells between the first and second week in culture (two-way ANOVA, F(1,216)=88.235, p<0.0001) and that emanating cells were able to transit through the myogenic compartments to form myotubes. By the second week in culture there was no significant difference in the proportion of nuclei in each myogenic compartment among all age groups. Table 2 further summarizes the distribution of the myofibers according to the number of emanating cells by week 2 in culture. The data underscore the dramatic difference in the number of cells derived from young myofibers in comparison to the number of cells derived from myofibers of the other age groups. This difference is in agreement with the increased number of satellite cells in the younger age group (Fig. 2 and Table 1).

Notably, in both 1- and 2-week old cultures, the frequency of myofiber cultures containing few or no cells was higher in the old and senile groups than it was in the young and adult groups (Chi square₉=76.588, p<0.001; Fig. 2 and Table 2). This finding is in agreement with the increased number of EDL myofibers lacking satellite cells as shown in Fig. 2. Likewise, there was an age-dependent decline in the number of myofibers giving rise to myogenic clones (data not shown). However, there was no significant difference in clone size among the different age groups (data not shown) and in the degree of morphological differentiation (estimated by the extent of myotubes formed, Fig. 3I–K). In sum, the study of cells emanating from single myofibers shows that the pool size of satellite cell progeny, and not the differentiation capacity of the cells, declined with age.

Myogenesis in primary cultures: initial proliferative phase is retarded in progeny of satellite cells from senile muscle but myogenic differentiation is not impaired

The study of cells emanating from single myofibers indicated an enhanced growth potential in myofiber cultures from the younger age group. This enhanced growth could be due to the larger initial number of satellite cells, to the potential presence of better growth signals from the parent myofibers, and/or to inherent differences in the proliferative capacity of the satellite cells founding the myofiber cultures. To exclusively compare the proliferative capacity, myogenic progenitors were isolated from young and senile muscles and cultured at the same initial density in rich growth medium. We studied young and senile age groups since they represent the best phase of muscle maintenance and the most extreme phase of muscle deterioration, respectively.

Primary cultures were analyzed for growth (total cell numbers), proliferation (number of Pax7⁺/MyoD⁺ cells), differentiation (number of Pax7⁻/MyoD⁺ cells, myogenin⁺ cells, and nuclei in myotubes) and accumulation of reserve cells (Pax7⁺/MyoD⁻). Representative immunostaining images are depicted in Fig. 5 and the data are summarized in Fig. 6. Values in Fig. 6B were calculated based on the total number of DAPI-stained nuclei. The total cell number (Fig 6A) includes remaining cells that did not label with either of the antibodies against the myogenic markers used in this study. These remaining cells have been considered non-myogenic and constitute the extra cells that are not categorized in Fig 6B.

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Fig. 5

Parallel immunofluorescent images depicting myogenesis in primary cultures prepared from the tibialis anterior and gastrocnemius muscles of senile mouse. Cultures shown were fixed on day 7 (A–D) and day 10 (E–H). Cultures were triple-immunolabled for MyoD (A, E), myogenin (B, F) and sarcomeric myosin (D, H; these images merged with parallel DAPI stain). Merged images of MyoD and myogenin are also shown (C–G). The same immunostaining patterns were observed in cultures from young mice, but with an earlier onset of differentiation. Bar, 100 μm.

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Fig. 6

Dynamics of cell growth and myogenesis in primary cultures from young and senile muscle. Cultures were initiated in parallel from one young and one senile mouse. Total cell numbers (Panel A) and myogenic progression (Panel B) were quantified based on DAPI- and immuno-labeling. The total cell number includes remaining cells that did not label with either of the antibodies against the myogenic markers used in this study. These remaining cells were considered non-myogenic. Panel A depicts the mean number (±SEM) of total nuclei based on six parallel cultures per time point. Panel B depicts the kinetics of myogenic progression in the cultures from young (left inset) and senile (right inset) muscles; for each time point, the different segments of the stack bar reflect the percent of nuclei in different myogenic compartments; the residual cells that were not identified by the antibodies are most likely non-myogenic. At each time point, the following antibody combinations were used for double- and triple-immunolabeling of parallel cultures: Pax7/MyoD, Pax7/MyoD/myosin, MyoD/myogenin/myosin (repeating each combination twice). While cells were not co-labeled for Pax7 and myogenin, the combinations of the four markers could be inferred from the direct counts. For example, the proportion of Pax7⁻/MyoD⁺/myogenin⁺/myosin⁺ cells was calculated based on the set of cultures labeled for MyoD/myogenin/myosin (which demonstrated that all cells positive for both MyoD and myosin were also positive for myogenin) and the set of cultures labeled for Pax7/MyoD/myosin (which identified the percent of cells that were Pax7⁻/MyoD⁺/myosin⁺); the latter cells were thus considered to be Pax7⁻/MyoD⁺/myogenin⁺/myosin⁺. The proportion of Pax7⁺/MyoD⁺/myogenin⁺/myosin⁺ cells was deduced in the same manner, based on the number of Pax7⁺/MyoD⁺/myosin⁺ cells. Data shown are representative of three independent experiments.

Between the 6^th and 4^th day after plating, the number of cells (i.e., total DAPI-stained nuclei) in cultures from young mice increased 15-fold whereas numbers in senile cultures increased only 10-fold (Fig. 6A). These data show a more robust growth rate in the young compared to the senile group during early days in culture. The increase in cell numbers between successive time points eventually slowed-down to a 1.8-fold value in cultures from both age groups (along with differentiation and fusion into myotubes), but the decline was more gradual in the senile group. While the decline to the 1.8-fold value occurred between days 6 to 7 in the young group, it occurred between days 7 to 8 in the senile group. Altogether, there was a persistent 24-hour lag before cultures of senile muscle contained the same cell number as those of young cultures. In accordance with the dynamics of cell accumulation, the onset of differentiation in the senile cultures was delayed by 24 hours in comparison to their young counterparts; however, there was no significant difference in the transition through the myogenic compartments (Fig. 6B). Collectively, the study of primary cultures suggests that the delay in differentiation was a result of a reduced proliferative – but not a reduced differentiative – potential of satellite cell progeny from the senile muscle. The intrinsic ability to transit through the myogenic program and yield myotubes was not impaired with age. However, the kinetics of the increase in total cell number over time in culture pointed to a possible inherent impairment in the initial amplification capacity of satellite cells of senile muscle.

The size of the satellite-cell pool declines with age

In this study, satellite cells were quantified along the entire length of individual myofibers, based on monitoring Pax7-expressing cells in a large sample size. Most previous reports on age-related changes in the satellite cell pool have relied on electron microscopy to identify satellite cells in muscle sections (Gibson and Schultz, 1983; Kadi et al., 2004; Roth et al., 2000). In these reports, satellite cell quantification has been represented as ratios between the number of satellite cells and a variety of factors such as the number of myonuclei in the same microscopic fields, myofiber diameter, and myofiber unit length. In the absence of a common denominator, discrepancies in the reported density of satellite cells per myofiber often arise even when analyzing the same muscles. In other studies, the number of progeny emanating from myofibers was used as a measure for the initial number of satellite cells; however, this approach has not taken into consideration the possibility that not all satellite cells emigrated from the myofibers and/or synchronously entered proliferation (Bockhold et al., 1998). Here, we determined the absolute number of satellite cells – an approach that permitted accurate fingerprinting of satellite cell profiles within and among muscles of different ages.

We show that satellite cell numbers in myofibers from both EDL and soleus muscle severely decreased and the frequency of myofibers lacking or having just one satellite cell significantly increased towards the end of the mouse lifespan. This “end of life crisis” in satellite-cell pool size may reflect an overall deterioration in body systems. However, before the onset of this “crisis”, there are clear differences in the age-associated pattern of satellite cell loss between EDL and soleus muscles. In EDL muscle, a major decline in the number of satellite cells occurred by 1 year of age. Conversely, in the soleus muscle, a drastic decrease in satellite cell numbers occurs only near 2.5 years of age. Furthermore, while a significant number of myofibers lacking satellite cells was apparent in the EDL muscle of the old-age group; such myofibers were present in the soleus muscle only at the senile stage. Altogether, the dynamics of age-linked satellite cell loss in EDL and soleus muscles are in accordance with established fiber-type dependent atrophy that occurs during muscle aging, reflecting a potential role for satellite cell abundance in the differential atrophy of fast and slow muscles.

Dual-immunolabeling with an antibody against syndecan-4 and MyoD was recently used as a means to quantify satellite cells in TA myofibers (Brack et al., 2005). Images from that study indicated that many additional entities in freshly-isolated TA myofibers, which seemed to be myofiber nuclei, were dually-labeled for syndecan-4 and MyoD. It is thus unclear how the distinction between satellite cells and myofiber nuclei was achieved. Indeed, we noted syndecan-4 expression associated with myonuclei when studying freshly-isolated myofibers. Nevertheless, many of these nuclei were not co-immunolabeled with Pax7 or within MAPK⁺ cells (G. Shefer and Z. Yablonka-Reuveni, unpublished; antibody was kindly provided by Dr. A. Rapraeger, University of Wisconsin, Madison). Therefore, we did not utilize syndecan-4 expression as a general and unequivocal marker of satellite cells. Also, since only infrequent satellite cells were positive for MyoD in freshly-isolated myofibers, we do not recognize MyoD expression as a general marker of quiescent satellite cells in such freshly-isolated myofibers. In contrast, Pax7-immunolabeling showed an excellent match between our data and those of Collins et al. (2005) where myofibers from the young-age group were studied. Collectively, we conclude that Pax7 immunolabeling provides a highly reliable means for identification of satellite cells in isolated myofibers from limb muscles, regardless the age of the animal being studied.

Satellite cell amplification, but not differentiation potential, declines with age

Based on a comprehensive investigation of satellite cells in several culture systems we conclude that progeny of satellite cells of all ages maintains the potential to proliferate, differentiate and fuse into myotubes, as well as the potential to give rise to reserve (Pax7⁺/MyoD⁻) cells.

In the FDB culture system, where myofibers were maintained in basal medium, proliferation of satellite cells of senile origin was greatly reduced unless the cultures were supplemented with FGF2. Hence, the aging myofiber unit does not produce sufficient growth-promoting agents to sustain satellite cell proliferation. The impaired capacity of aging muscle to support satellite cell recruitment/proliferation was suggested to be a consequence of a decline in IGF-I (Barton-Davis et al., 1998; Chakravarthy et al., 2000; Musaro et al., 2001) or a reduction in notch signaling (Conboy et al., 2003; Conboy and Rando, 2002). The present study introduces a novel insight into the impact of FGF depletion in the aging muscle.

Early studies of satellite cells isolated from 1-year old rats by enzymatic digestion, proposed that a decline in available hepatocyte growth factor (HGF) is responsible for the age-linked proliferative-latency of satellite cells and that this latency could be overcome by the addition of HGF but not FGF. These latter studies suggest that HGF promotes upregulation of FGF receptors (Allen et al., 1995; Johnson and Allen, 1995). We have demonstrated a similar promoting effect of FGF and HGF on satellite cell proliferation in myofibers both from young rats (2–3 month old; Kastner et al., 2000) and from adult rats (9–12 month old (Yablonka-Reuveni et al., 1999b). Hence, both FGF and HGF may regulate satellite cell recruitment in the younger and the older rodents. However, the FGF receptor system may be more sensitive to the enzymatic and physical trituration steps involved in satellite cell isolation, resulting in the reported preferential proliferative response to HGF (compared to FGF) in primary cultures of freshly isolated satellite cells. Differently, the gentler procedure of myofiber isolation may preserve the FGF receptors. Collectively, the present study reinforces our earlier conclusion that the FGF system indeed plays a role in satellite cell recruitment and amplification.

There was a time lag in the accumulation of progeny in primary cultures from senile muscle in comparison to those from young muscle, even though cultures initially had the same cell density and were grown in potent growth medium that contained high levels of growth-promoting agents, including FGF2 (Yablonka-Reuveni, 1995). Conversely, proliferation of satellite cells from the senile muscle occurred in a single bout in the FDB myofiber culture system when supplemented with FGF2. Together, these studies establish that the association with the parent myofiber provides a competent environment for satellite cell activation and cell cycle progression, regardless of age. Such a promoting environment may be a result of paracrine interplay between the parent myofiber and its resident cells. Nevertheless, it is possible that the efficiency of the proliferative machinery is reduced with age, and this decline, combined with the impact of the enzymatic digestion required for satellite cell isolation, may lengthen the recovery time upon cell culturing in the primary culture model. This scenario may account for the reduced synchronization of senile muscle satellite cell proliferation during early days in culture, as observed in this study. Clearly, the present study has not addressed the possibility that the rich growth medium used in the primary culture model may contain a factor that can selectively delay proliferation of satellite cells from senile muscle and this issue is avoided in FDB cultures where a different medium is being used. We do not favor the latter possibility because the proliferative rate is reduced not only when comparing senile to young groups but also when comparing adult to young groups during early days in primary cultures (Allen et al., 1995; our unpublished results); thus, proliferative latency is not unique to the senile age. Notably, myofiber injuries resulting from day-to-day muscle utilization do not involve detachment of satellite cells from their natural niche (as when establishing primary cultures). Therefore, the FDB myofiber system may better represent the proliferative potential of satellite cells in adult and aging muscle.

Present results with the FDB system indicate the proliferation of satellite cells of senile origin can be as efficient as that of satellite cells from young muscle, when proper growth-promoting agents are provided. While the present investigation focused on FGF, other growth factors known to promote satellite cell proliferation may also enhance the mitogenic milieu in aging muscle. A mere analysis of types and levels of growth factor (FGF isoforms included) expressed in old muscle will not necessarily provide a clear insight into the role of growth-promoting factors in aging muscle. For example, while IGF-I overexpression reduces age-linked myofiber atrophy, IGF-I expression level is not reduced in aging muscle; IGF-I signaling can be impaired by cytokines and the latter are known to increase systemically with age (Barton-Davis et al., 1998; Chakravarthy et al., 2000; Edstrom and Ulfhake, 2005; Grounds, 2002; Musaro et al., 2001). Studies with genetic models of inducible growth factor over/under expression are now required to evaluate the effect of growth promoting agents on enhancing satellite cell function during aging.

The present clonal study indicates that clonable satellite cells preserve their amplification and differentiation potential throughout life. Conversely, it has been suggested that the proliferative capacity of satellite cells from old muscle is impaired, based on the smaller number of cells that developed in satellite cell clones from old muscle as compared to clones from young muscle (Schultz and Lipton, 1982). However, the number of cells per clone was studied only during the early days in culture; therefore, a lag in the onset of proliferation, rather than an inherent reduced proliferative capacity, may account for the smaller clones derived from old muscle. Indeed, our analysis of clones at later time points (i.e., 10–14 days in culture) did not reveal apparent differences in clonal growth or in the degree of fusion into myotubes regardless of the age of the donor mouse. Furthermore, RT-PCR analysis of these clones did not reveal any apparent differences in expression levels of muscle-specific transcription factors, structural proteins or various growth factors shown to regulate myogenesis (ongoing studies). In another study reporting on the age-associated decline in clonal size, cultures were expanded long-term, prior to the establishment of clones (Chakravarthy et al., 2000). Therefore, the identity of the subpopulation of cells that founded these clones could not be clearly evaluated since they were not derived directly from satellite cells (Machida et al., 2004). Furthermore, nearly all cell culture studies on the differentiation of satellite cells from aging muscles, have relied on cells that were propagated for long-term (i.e., passaged cells), in order to amplify cell numbers for morphological and biochemical analysis (e.g., Beggs et al., 2004; Chakravarthy et al., 2000; Lees et al., 2005; Taylor-Jones et al., 2002). This propagation process may not only result in reduced myogenicity of satellite-cell progeny (Machida et al., 2004), but also preferentially enhance the growth of non-myogenic cells. The presence of non-myogenic cells can bias conclusions about age-dependent properties of satellite-cell progeny.

Collectively, our results show that, although satellite cell amplification is impaired within the aging muscle, satellite cell functioning can be rejuvenated when the cells receive proper growth-promoting signals.

We express gratitude to Drs. Peter Rabinovitch, Norman Wolf and Warren Ladigen, and their team members (University of Washington) for generously providing or sharing mice throughout the study. We are also grateful to Drs. Michael Rudnicki (Ottawa Health Research Institute) and Shahragim Tajbakhsh (Pasteur Institute) for the Myf5^nlacZ/+ mouse strain.

This work was supported by grants to Z.Y.-R. from the National Institute on Aging (AG21566 and AG13798) and the USDA Cooperative State Research, Education and Extension Service (NRI, 99-35206-7934).