Atrophy of skeletal muscle leads to decreases in myofiber size and nuclear number; however, the effects of atrophic conditions on muscle precursor cells (MPC) are largely unknown. MPC lie outside myofibers and represent the main source of additional myonuclei necessary for muscle growth and repair. In the present study, we examined the properties of MPC after hindlimb suspension (HS)-induced atrophy and subsequent recovery of the mouse hindlimb muscles. We demonstrated that the number of MPC in atrophied muscles was decreased. RT-PCR analysis of cells isolated from atrophied muscles indicated that several mRNA characteristic of the myogenic program in MPC were absent. Cells isolated from atrophied muscles failed to properly proliferate and undergo differentiation into multinucleated myotubes. Thus atrophy led to a decrease in MPC and caused dysfunction in those MPC that remained. Upon regrowth of the atrophied muscles, these deleterious effects were reversed. Our data suggest that preventing loss or dysfunction of MPC may be a new pharmacological target during muscle atrophy.
- satellite cells
- hindlimb suspension
skeletal muscle atrophy occurs in response to widely divergent environmental and pathophysiological stimuli. These stimuli range from exposure to a microgravity environment and localized denervation in which selective atrophy of muscles occurs (30) to the general wasting of many muscles in patients with diseases such as cancer (46), acquired immunodeficiency syndrome (17), and cardiac disease (3), as well as in older adults (52).
The majority of studies to date have investigated the effects of atrophic stimuli on myofibers. Myofiber atrophy is associated with the elimination of nuclei from myofibers apparently via an apoptotic mechanism (1, 13, 47, 48). Nuclei may be eliminated from the atrophying myofibers to compensate for the decreased volume and transcriptional/translational requirements of the myofiber. Compensatory myonuclear loss during atrophy may be related to the concept that each myonucleus has a tightly regulated nuclear domain (2). A nuclear domain is the volume of cytoplasm regulated by the gene products of a single myonucleus (18, 31). Because of the postmitotic nature of myonuclei, growth of atrophied myofibers depends on resident muscle precursor cells (MPC) that exist outside myofibers (28). Spatially, MPC can exist as satellite cells that lie underneath the basal lamina of myofibers (25) or as stemlike cells that exist in the interstitial area between myofibers (41), in fat (53), and in blood vessels (27). Satellite cells are the main subpopulation of MPC that contribute to muscle growth and regeneration. In adult skeletal muscle, the majority of MPC are quiescent but, in response to growth stimuli, are activated to reenter the cell cycle. Some MPC terminally differentiate and fuse with one another or with the growing myofiber, whereas a small proportion of MPC instead resume a quiescent state outside the myofiber to provide a “reserve” population of cells for future growth and repair.
To date, few studies have investigated the effects of atrophic stimuli on MPC. In some studies, a decreased number of MPC were observed regardless of the atrophy-inducing stimulus used (20). Increases in apoptotic nuclei outside atrophied fibers (1, 38, 48) could account for these decreases in the number of MPC. In other studies, MPC dysfunction apparently is induced, which adversely affects the ability of MPC to participate appropriately in the myogenic program. For example, prolonged HS-induced atrophy appears to result in long-lived deficits in the MPC of adolescent and aged animals, manifested by an inability of muscle tissue to grow or to regenerate (5, 10, 29). In addition, atrophy can cause changes in MPC, manifested by an increased susceptibility of these cells to proapoptotic stimuli in vitro (21). Atrophic conditions induced by a variety of stimuli are associated with increases in catabolic factors such as myostatin (15, 24, 33) and tumor necrosis factor α (34) as well as decreases in trophic factors (39). The presence or absence of such factors could negatively affect MPC (14, 26, 40). Studies to date have lacked direct molecular and cellular analyses of the ability of MPC from atrophied muscles to proliferate and differentiate.
This report describes experiments that we performed to test the hypothesis that prolonged atrophic conditions lead to MPC dysfunction. We used several techniques to examine the effects of atrophy on the ability of MPC to activate, proliferate, and form multinucleated, differentiated myotubes. These properties also were examined after allowing atrophied muscles to regain their size upon cessation of the atrophic stimulus. MPC from atrophied muscles are impaired in their ability to activate and proliferate and also fail to differentiate properly into multinucleated myotubes. In contrast, MPC from recovered muscle do not differ from control.
MATERIALS AND METHODS
Female C57BL/6 NcrlBR and Balb/c AnNHsd mice (10–12 wk) were purchased from Charles River Laboratories and Harlan Sprague-Dawley, respectively. All mice were housed under a 12:12-h light-dark cycle at room temperature, and food and water were provided ad libitum. All procedures were performed in accordance with Emory University's Institutional Animal Care and Use Committee. Mice were randomly assigned to three groups: hindlimb suspension (HS; group 1), HS followed by recovery (group 2), or control (group 3). In groups 1 and 2, mice were suspended from a tail harness for 2 wk as described previously (28). Mice from group 1 were killed after the 2-wk HS period, whereas the tail harness was removed from mice in group 2 by briefly anesthetizing the mice with xylazine (10 mg/kg body wt) and ketamine (65 mg/kg body wt). Mice were allowed to resume normal cage activity during their period of least activity (light cycle) to minimize damage to the hindlimb muscles during the initial recovery period.
Quantification of Muscle Atrophy
Mice were euthanized by CO2 inhalation, and gastrocnemius (GTN), soleus (SOL), and plantaris (PLAN) muscles, individually or as a group, were collected using standardized dissection methods and cleaned of excess fat, connective tissue, and tendons. Muscles were immediately weighed on an analytical balance and used for subsequent experiments. The SOL, PLAN, and GTN muscles comprised 3.6%, 8.3%, and 88% of the control hindlimb muscle group, respectively. After 2 wk HS, the SOL comprised significantly less (2.5%) of the hindlimb group and the GTN comprised significantly more (89.3%) of the same group. The SOL, PLAN, and GTN muscles atrophied by 52%, 25%, and 24%, respectively, after 2 wk of HS (n = 5; P < 0.05) (Table 1). Accordingly, this hindlimb group, composed of all three muscles, atrophied by 25%. The same degrees of atrophy are attained when muscle weights are normalized to body weight. This is because, in our hands, HS mice do not lose an appreciable amount of body mass. Complete recovery of mass in atrophied muscles was attained after 2 wk in Balb/c mice (28), whereas the C57BL/6 strain required 3 wk (data not shown).
MPC Isolation and Cell Culture
The pooled GTN, SOL, and PLAN muscles of C57BL/6 mice (n = 4 for each group) were subjected to both mechanical and enzymatic dissociation. Muscles were minced into a coarse slurry and digested for 1 h with 0.1% pronase (Calbiochem) in DMEM containing 25 mM HEPES at 37°C with gentle agitation. Subsequently, the digest was mechanically dissociated by triturating the muscle slurry repeatedly and filtered through a 100-μm filter (Millipore). The filtered digest was centrifuged through an isotonic Percoll gradient (60% overlaid with 20%) as described previously (51). Cells were collected from the interface of the Percoll gradient and either counted, lysed in Trizol reagent (Life Technologies) for RT-PCR analysis, or resuspended in primary growth medium (Ham's F-10: 20% FBS, 5 ng/ml bFGF, 100 U/ml penicillin G, and 100 μg/ml streptomycin) and grown on entactin, collagen IV, and laminin (E-C-L; Upstate Biotechnology)-coated tissue culture dishes or Permanox cell culture slides (Nalge Nunc International) in a humidified, 37°C, 5% CO2 incubator. Myogenic differentiation was induced by plating equal numbers of cells and culturing in differentiation medium [DMEM, insulin-transferrin-selenium-A supplement (GIBCO), 100 U/ml penicillin G, and 100 μg/ml streptomycin] for 5–7 days.
Isolation and Culture of Single-Myofiber Explants
Single myofibers were isolated from the GTN muscles of C57BL/6 mice (n = 4 for each group) as described by Rosenblatt et al. (35). Briefly, the GTN muscle was dissected and digested in DMEM containing 0.1% collagenase (type I, 400 U/ml; Worthington Biochemical) at 37°C for 1 h with gentle agitation. Single myofibers were then dissociated by repeated gentle trituration of the digested muscles. Using a dissecting microscope, single myofibers were extracted individually using fire-polished pipettes and transferred serially into fresh plates containing DMEM with 4.5 mg/ml of glucose, 10% FBS, 100 U/ml of penicillin G, and 100 μg/ml of streptomycin. Single myofibers were transferred individually to 24-well plates precoated with growth factor-reduced Matrigel matrix (BD Biosciences), centrifuged for 40 min at 1,100 g to facilitate adhesion to the Matrigel, and incubated for 24 h in a humidified, 37°C, 5% CO2 incubator.
The following antibodies were used: anti-myoD (NCL-MyoD1, 1:30 dilution; NovoCastra), anti-desmin (1:200 dilution; Sigma), anti-embryonic myosin heavy chain (anti-eMyHC; F.1652, neat; Developmental Studies Hybridoma Bank), anti-bromodeoxyuridine (anti-BrdU, BU1/75, 1:500 dilution; Harlan Sera-Lab), and anti-dystrophin (MANDYS8, 1:400 dilution; Sigma). The appropriate isotype control antibodies were purchased from Jackson ImmunoResearch. Primary MPC and single-myofiber explants were rinsed with PBS and fixed in 2% paraformaldehyde for 10 min. All steps were performed at room temperature unless otherwise noted. Images of cells and single-myofiber explants were obtained using a Zeiss Axiovert 200M microscope and Openlab version 3.1.5 software (Improvision). Analyses and photography of muscle sections were performed on a Zeiss Axioplan microscope equipped with a video camera and Scion Image and Adobe Photoshop software.
Counting of nuclei outside myofibers.
Triplicate sections from the belly of the SOL muscles from Balb/c mice (n = 4 for each group) were blocked with 5% goat serum (Sigma) in PBS containing 0.2% Tween 20 (PBST) for 30 min and incubated with anti-dystrophin antibody in blocking buffer for 1 h. Sections were washed with PBST and then incubated with Texas red-conjugated goat anti-mouse IgG (1:50 dilution; Cappel). After further washing in PBST, the sections were fixed in 2% formaldehyde for 20 min. The sections were again washed in PBST and incubated with the nuclear dye 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI, 250 ng/ml; Sigma) for 5 min, washed extensively, and mounted in Vectashield mounting medium (Vector Labs). Nuclei that were outside the myofiber sarcolemma as defined by dystrophin staining were counted as 150–200 fibers per muscle section.
MyoD and desmin costaining.
Endogenous peroxidase activity in primary MPC and single-myofiber explants was quenched by incubating in 3% hydrogen peroxide for 10 min. Cells were blocked with 5% donkey serum (Sigma) in 0.25% Triton X-100 for 1 h and incubated with anti-myoD (mouse) and desmin (rabbit) antibodies in blocking buffer overnight at 4°C. The tyramide signal amplification system (TSA Fluorescein System; NEN) was used to detect myoD according to the manufacturer's protocol. Briefly, cultures were incubated with TNB buffer (0.1 M Tris·HCl, pH 7.5, 0.15 M NaCl, and 0.5% blocking reagent) for 1 h, followed by biotin-conjugated donkey anti-mouse F(ab′)2 fragments (1:500 dilution; Jackson ImmunoResearch) and Texas Red-conjugated donkey anti-rabbit IgG (1:500 dilution; Jackson ImmunoResearch) for 1 h. Cultures were washed with PBST and incubated with streptavidin-horseradish peroxidase (diluted 1:200 in TNB buffer) for 30 min followed by fluorescein-tyramide (diluted 1:250 in amplification diluent) for 5 min. After another series of washes, cultures were incubated with 250 ng/ml DAPI in PBST for 5 min so that nuclei could be visualized.
BrdU and desmin costaining.
Primary MPC were cultured for 48 h and pulsed with 25 μM BrdU (Sigma) for 1 h, rinsed with PBS, and fixed in 2% paraformaldehyde for 10 min. After a series of PBS washes, DNA was denatured with 1 N HCL in PBS for 45 min at 45°C, followed by neutralization in 0.1 M borate buffer for 10 min. Cells were washed with PBS and then incubated with blocking buffer (5% donkey serum, 0.5% bovine serum albumin, and 0.25% Triton X-100 in PBS) for 1 h. Cells were stained for desmin as described above. To detect BrdU, cells were first incubated with anti-BrdU antibody in blocking buffer for 1 h and washed with PBST, followed by secondary antibody [FITC-conjugated donkey anti-rat F(ab)2, 1:500 dilution; Jackson ImmunoResearch] for 1 h. After another series of washes, cells were counterstained with DAPI so that nuclei could be observed as described above.
Primary MPC were blocked with 5% goat serum in PBST for 30 min followed by 1-h incubation with anti-eMyHC antibody. Cells were washed with PBST and incubated with Texas Red goat anti-mouse F(ab′)2 fragments (1:500 dilution; Cappel) for 1 h. After another series of washes in PBST, cells were counterstained with DAPI as described above.
RT-PCR Analysis of mRNA Expression
RT-PCR was performed using 1.0 μg of total RNA per sample. All PCR were performed using 1 μl of the above RT-PCR product, and primers specific for each particular gene and designed to cross intron-exon boundaries (Table 2) were used to generate amplicons in their linear range. 18S rRNA was used as an internal control in each sample using standard QuantumRNA 18S primers (Ambion). PCR products were resolved on 1.5% agarose gels, and the 18S rRNA amplicon was used to verify equal input of RNA to the PCR reaction.
To determine significance between two groups, comparisons were made using Student's t-test. Analysis of multiple groups was performed using one-way ANOVA with Bonferroni's posttest using GraphPad Prism version 4.0a for Macintosh (GraphPad Software, San Diego, CA). For all statistical tests, P < 0.05 was accepted for statistical significance.
Number of Cells Residing Outside Myofibers Decreased After 2 Weeks of HS
To determine whether mononucleated cells were eliminated from muscles in response to atrophic conditions, DAPI-stained nuclei outside SOL myofibers were counted (Fig. 1A). A significant decrease (∼12%) in the number of nuclei outside myofibers was observed after 2 wk of HS (Fig. 1B). We previously demonstrated a 36% decrease in the number of myonuclei with the same atrophic stimulus (28). Upon recovery of muscle mass, the number of nuclei outside myofibers increased 42% and 63% above control and HS values, respectively (Fig. 1B), indicating that marked proliferation of cells occurred during the recovery period to contribute not only to replenishing myonuclei but also to cells outside myofibers.
Atrophic Conditions Adversely Affected Expression of Myogenic mRNA in MPC
To better identify which cells were affected during atrophy and subsequent regrowth of muscle, we isolated mononucleated cells from pooled GTN, SOL, and PLAN muscles and analyzed the expression of a variety of myogenic markers by performing immunohistochemical and RT-PCR analyses. First, total numbers of isolated cells were quantified. Atrophied muscles consistently yielded ∼14% fewer cells than control muscles, whereas recovered muscles yielded ∼50% and ∼75% more cells than control and atrophied muscles, respectively (Fig. 2A). To account for contaminating nonmuscle cells, cells were immunostained for desmin, a muscle-specific intermediate filament protein. Desmin-positive cells composed ∼90%, 70%, and 98% of cultures from control, atrophied, and recovered muscles, respectively (Fig. 2B). The magnitude of the changes in cell number from atrophied and recovered muscles correlates well with the changes in the number of interstitial nuclei observed in cross sections of atrophied and recovered muscles (Fig. 1B). Moreover, the magnitude of the loss of desmin+ cells in atrophied muscles (22% decrease vs. 14% decrease in total cells) suggests a selective loss of myogenic cells.
Because of differences in the numbers of total cells isolated from the different experimental groups, all subsequent analyses were normalized to cell number. Mononucleated cells from control, atrophied, and recovered muscles were harvested for RNA immediately after isolation. Subsequent RT-PCR analyses revealed that although expression of Pax7 mRNA did not appear to be altered, expression of m-cad, c-met, myf5, and syndecan-4 mRNA was absent in cells isolated from atrophied muscle (Fig. 2C). All four of these mRNA were previously shown to be characteristic markers of myogenic cells that reside in the satellite position outside myofibers (4, 6, 7). These data suggest a specific adverse effect on satellite cells in atrophied muscle. Expression of myogenic mRNA markers was upregulated in cells isolated from recovered muscles, indicating that MPC can overcome the adverse effects observed during atrophic conditions (Fig. 2C).
Fewer Satellite Cells Are Associated with Atrophied Single-Myofiber Explants
To assess the prevalence of satellite cells, we isolated single myofibers from control, atrophied, and recovered GTN muscles and determined the number of satellite cells associated with each myofiber after 24 h ex vivo. This time point represents the stage at which the majority of satellite cells have activated and migrated away from the myofiber and cell division has not occurred. Fewer satellite cells emanate from atrophied myofibers; however, recovered myofibers yield an increased number of satellite cells (Fig. 3A). The frequency distribution profile clearly shows that individual myofibers from atrophied muscle had fewer resident satellite cells than did myofibers from control and recovered muscles (Fig. 3B). Similar results were observed when these cultures were immunostained for myoD, a muscle-specific basic helix-loop-helix transcription factor (Fig. 3C). The increased number of satellite cells from recovered myofibers may be due to a compensatory expansion of cells to replenish the myonuclei and satellite cells eliminated during the atrophic period.
Impairment of Activation and/or Proliferation of MPC from Atrophied Muscles
The expression of myoD is upregulated upon MPC entry into the cell cycle and is present throughout the proliferative stage of these cells. To determine whether myogenic cells isolated from atrophied muscles are impaired in activation, we performed immunohistochemical analyses of myoD in total cell isolates from pooled GTN, SOL, and PLAN muscles. After 24 h ex vivo, we immunostained cells for myoD and desmin and expressed the number of myoD+ cells as a function of desmin+ cells to account for differences in the number of desmin-positive cells. In cells from atrophied muscles, desmin+/myoD− cells were often observed (Fig. 4A). Cells that were myoD− are also negative for eMyHC, a marker of differentiated cells (data not shown), thus ruling out the possibility that some cells differentiated with a concomitant decrease in myoD expression. A 16% decrease in the percentage of myoD+ myogenic cells was observed in atrophied cultures compared with control (Fig. 4C). MyoD expression in myogenic cells from recovered muscles was not significantly different from that in control.
Single myofibers from GTN muscles were also immunostained for myoD and desmin (Fig. 4B). Atrophied myofibers produced fewer myoD+/desmin+ cells per myofiber, whereas recovered myofibers consistently produced more (Fig. 4D). To determine whether proliferation of atrophied MPC is affected, cells were isolated from the pooled GTN, SOL, and PLAN muscles, labeled with BrdU, and immunostained for BrdU and desmin. In atrophied muscles, a 25% decrease in the number of BrdU+/desmin+ cells was observed (Fig. 4E). The presence of desmin+/myoD− cells and a decrease in the prevalence of BrdU+/desmin+ cells in cultures from atrophied muscle suggest that activation and/or proliferation of MPC was affected.
MPC Isolated from Atrophied Muscles Form Fewer and Smaller Myotubes
We next compared the ability of control, atrophied, and recovered MPC to differentiate and form multinucleated myotubes. Upon isolation from pooled GTN, SOL, and PLAN muscles, equal numbers of cells from control, atrophied, and recovered muscles were plated directly in myogenic differentiation medium for up to 7 days. Obvious differences were already apparent after 2 days in culture. Growth of control and recovered cells was observed 1–2 days after isolation, whereas growth of atrophied cells was delayed and, on average, was observed 3–4 days after plating (Fig. 5A), consistent with the decreased BrdU incorporation shown in Fig. 4E. Morphologic signs of differentiation (alignment of MPC and formation of nascent myotubes) were observed in control and recovered cells by day 3, compared with day 5 for atrophied cells (Fig. 5A). Formation of large myotubes in control and recovered cultures was complete by day 5. In contrast, atrophied cells must be cultured for a longer period (7 days) for myotube formation to occur, even though confluence was attained by day 5 or 6 (Fig. 5A). Atrophied cells, even when cultured for this longer period, never formed large myotubes. Immunostaining for eMyHC after 7 days in differentiation medium revealed much smaller myotubes with fewer myonuclei than control or recovered cultures (Fig. 5B). Thus cells from atrophied muscles appear to have decreased ability to form myotubes.
We used multiple assays to demonstrate that MPC are adversely affected by HS-induced atrophic conditions. MPC were studied in bulk cultures and derived from single-myofiber explants. The yields of both total mononucleated cells and myogenic cells from atrophied muscle were decreased. Accordingly, analyses of mRNA expression indicated that atrophic conditions adversely affected molecular markers of myogenic cells. In addition, dysfunction of MPC also occurred. The ability of MPC from atrophied muscles to activate or to proliferate normally and form multinucleated myotubes ex vivo was impaired. These deleterious effects on MPC were alleviated in recovered muscles.
Decreases in the number of nuclei outside myofibers, as well as the yield of mononucleated cells after 2 wk of HS, are consistent with the results of previous studies that showed an increase in the number of apoptotic nuclei both inside and outside myofibers in muscles induced to atrophy via different mechanisms (1, 4, 48). In addition, the decreased yield of myogenic cells could arise from cell fusion with atrophying myofibers. However, myonuclear number is decreased during atrophy (2), making this possibility unlikely. The reduction in the number of nonmuscle cells as well as myogenic cells in muscle tissue (Figs. 1 and 3) may be due to a cell-extrinsic mechanism. Atrophy is associated with decreased production of trophic factors in muscle tissue. For example, VEGF expression is downregulated during HS (39) and is an essential survival factor for both endothelial cells in capillaries (42) and myoblasts (14). Alternatively, production of proapoptotic factors may increase during atrophy. For example, TNFα is increased in response to atrophy (9) and can enhance apoptosis of myoblasts in vitro (40). Cell-intrinsic mechanisms may also contribute to the loss of myogenic and nonmyogenic cells in atrophied muscle. Gene array studies of atrophying muscles (39, 44) demonstrated downregulation of genes involved in cell replication as well as upregulation of genes involved in cell cycle arrest. Furthermore, MPC isolated from atrophied muscles were found to be more susceptible to apoptotic stimuli in vitro (21). Previous studies in which HS of rodents was performed demonstrated that osteogenic cells isolated and cultured from unloaded bones retained the deleterious effects of the in vivo unloading conditions (16, 22). Thus cells extracted from the hindlimb retain some memory of their physiological environment, arguing for intrinsic changes in these cells.
That intrinsic changes occur in MPC is supported further by our present results demonstrating that even in the presence of high-serum-containing medium, bulk MPC as well as satellite cells from single-myofiber explants had a diminished ability to activate and express myoD (Fig. 4). Accordingly, a higher proportion of MPC isolated from atrophied muscles was desmin+/myoD−. Proliferation studies showed that fewer cells in these cultures incorporated BrdU (Fig. 4E). The possibility exists that culturing cells ex vivo for longer periods of time may allow these cells to overcome the deleterious atrophic effects.
Not only were early stages of myogenesis impaired by atrophy in MPC but also later stages characterized by differentiation and fusion to form multinucleated myotubes. Specifically, myotube formation occurred later in atrophied cultures, and the number and the size of myotubes formed was smaller than those of controls. Delayed terminal differentiation and subsequent fusion to form myotubes is unlikely to be due to the initial slow expansion of cells from atrophied muscles. We routinely observe no major differences in myotube formation between newly isolated primary MPC and those cultured for several weeks in our lab. Therefore the few extra cell doublings required for cells from atrophied muscle to reach confluence is unlikely to have a significant impact on myotube formation. However, when myotube formation does occur in cultures from atrophied muscles, these cells have reached confluence, arguing against a lack of cells to facilitate fusion to form large myotubes. The prevalence and large size of myotubes observed in control cultures were never observed in cultures from atrophied muscles, even after 2 additional days in differentiation medium. These results suggest that the myogenic capacity to form myotubes is impaired in MPC isolated from atrophied muscles. In support of this idea, more mononucleated, differentiated (eMyHC+) cells were observed in cultures of MPC from atrophied muscles (data not shown). Contributing to problems with myotube formation, however, may be the fact that cultures from atrophied muscle contained 22% more nonmuscle cells (Fig. 2B) than controls. Contamination of myogenic cultures by fibroblasts was previously shown to adversely affect MPC differentiation (50). However, the culture conditions used in our study favored myogenic cells because E-C-L (a matrix containing laminin) promotes adhesion of myogenic cells rather than fibrogenic cells (23). In addition, cells were cultured directly in a low-mitogen-containing medium that favors myogenic differentiation.
A key finding of this study is that myogenic cells isolated from adult muscles that regained normal muscle mass after 2 wk were normal in their numbers and myogenic properties. Therefore no permanent changes in the MPC pool appear to have been induced during the 2-wk HS period, in contrast to other studies of either very young or old rodents. Several mechanisms could account for the recovery of MPC in our study. HS-induced atrophy may lead to a selective reduction in specific MPC populations. MPC are heterogeneous in a number of properties, including proliferation (19, 37) and differentiation (32). Regrowth of myofiber size after atrophy is dependent on MPC (28) and in this model would require expansion of selected pools of MPC. This pool of MPC may be limiting and/or absent at young and old ages and could account for the permanent deficits in muscle growth observed in these age groups. Alternatively, loss of MPC may not be selective, and regrowth of myofiber size may occur through simple recovery of normal MPC numbers and myogenic properties, which in some way is impaired at the two extremes of age. Our data do not distinguish between these two possibilities. In either case, significant expansion of the MPC pool is needed because irradiation impairs the ability of atrophied muscles to regrow (28). Further studies are required to determine whether multiple bouts of an atrophic stimulus lead to permanent adverse effects on the MPC population in adult rodents, even after the recovery period. In addition, prolonged atrophic conditions present in some human myopathies may induce permanent deleterious changes in the properties of MPC.
In Fig. 6, our current and previous work (28) is summarized to illustrate MPC dynamics during muscle atrophy and subsequent recovery. The number of myonuclei (28) and MPC (Figs. 2 and 3) decreased after 2 wk of HS-induced atrophy. The remaining MPC display dysfunction in their ability to activate and/or proliferate (Fig. 4) and differentiate (Fig. 5). Upon cessation of the atrophic stimulus, myofiber size and myonuclear number were restored after 2–3 wk (28). This recovery was dependent, in part, on MPC proliferation, differentiation, and subsequent fusion with myofibers. By the end of the recovery period, the number of MPC (Figs. 2 and 3), as well as their ability to activate and/or proliferate (Fig. 4) and differentiate (Fig. 5), was restored. Thus, in spite of the loss and cellular dysfunction of MPC, muscle growth eventually did occur. However, recovery would likely occur sooner if MPC were not adversely affected by the atrophic conditions. Factors that promote MPC survival and proliferation could be exploited therapeutically. Such factors would act to decrease the number of cell doublings required to replenish the MPC pool. Studies in dystrophic animals and humans have shown that repeated use of myogenic cells for muscle repair leads to exhaustion of the MPC pool (11, 49). Conceivably, preventing loss of MPC and nonmuscle cells may prevent loss of trophic factors produced by these cells and help to diminish myofiber atrophy. Alternatively, enhancing the ability of MPC to differentiate and fuse may promote muscle growth (45). To date, attention has been directed toward preventing apoptosis (8) and protein breakdown in myofibers (12, 36, 43) as a means of attenuating atrophy. Our data suggest that MPC may be a new target of pharmacological approaches to prevent or to treat muscle atrophy.
This work was supported by an American Heart Association predoctoral fellowship (to P. O. Mitchell) and National Institutes of Health Grants AR-47314, AR-48884, and DE-13040 (to G. K. Pavlath).
We thank Roddy O'Connor for assistance with single-myofiber experiments and Todd Mills for assistance with HS experiments.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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