, Volume 16, Issue 6, pp 747–759 | Cite as

Muscle-specificity of age-related changes in markers of autophagy and sphingolipid metabolism

  • David W. Russ
  • Iva M. Boyd
  • Katherine M. McCoy
  • Katherine W. McCorkle
Research Article


Our previous findings indicate that the gastrocnemius muscle of aging rats exhibits impairments of muscle quality (force/unit muscle tissue) and autophagy and increased sarcoplasmic reticulum stress. The purpose of this study was to examine age-related changes in soleus muscle contractility and in markers of autophagy in the soleus and gastrocnemius muscles. We assessed in situ muscle force and size in the soleus muscle of adult (7–8 months) and aged (24–26 months) male, F344/BN rats. We used immunoblotting to compare abundance of markers of autophagy, sarcoplasmic reticulum (SR) stress and sphingolipid metabolism in the soleus and medial gastrocnemius (MG) muscles of these animals. Relative to adults, aged rats maintained soleus muscle quality and increased muscle size, resulting in increased tetanic force production. Immunoblotting revealed a general pattern of an age-related reduction of basal autophagy, despite increases in indicators of SR stress and upstream autophagic pathway activation in the MG. The MG also exhibited changes in markers of sphingolipid metabolism suggestive of increased muscle ceramide. Minimal age-related changes were observed in the soleus. The soleus maintains muscle mass and quality with age, and exhibits fewer age-related changes in markers of stress and autophagy than the MG. Based on these data, we suggest that maintenance of autophagy may preserve muscle quality by preventing excessive SR stress.


Skeletal muscle Sarcopenia Ceramide Muscle quality 


Muscle function clearly declines with age, including its definitive function—contraction. Loss of contractile function manifests itself as weakness (i.e., loss of force and power production), which negatively influences several aspects of physical function (Bean et al. 2002; Visser et al. 2000) and even predicts mortality (Metter et al. 2002). Moreover, this weakness typically exceeds the loss of muscle mass (Russ et al. 2012a), suggesting that impairment of contractile function accompanies the loss of muscle cell number and size (Degens and Alway 2006; Lexell et al. 1988) that are associated with old age.

One potential mechanism for reduced tissue and organ function that has gained increasing attention in recent years is reduced autophagy (Hubbard et al. 2012; Kim et al. 2013; Wohlgemuth et al. 2010). While excessive autophagy elicited by pathologic stimuli (e.g., ischemia–reperfusion) is associated with certain disease processes and can be maladaptive, a normal level of basal autophagy is essential to the normal turnover of cell constituents and maintenance of cell function (Hubbard et al. 2012). Autophagy is a process of cellular self-degradation, largely characterized by bulk degradation of organelles and protein aggregates. The most-studied form of autophagy, macroautophagy (hereafter referred to as ‘autophagy’), is primarily involved with degradation of organelles (Bernales et al. 2007). It has been suggested that post-mitotic tissues, such as skeletal muscle, are more vulnerable to disruptions of autophagy (Terman and Brunk 2006) as they must maintain cellular function without relying on the proliferation of new cells.

We have previously reported that aging rat gastrocnemius muscles exhibit a proportionately greater loss of force production than mass (i.e., a loss of muscle quality) (Russ et al. 2011, 2014), and reduced abundance of markers of basal autophagy (Russ et al. 2012b), as have others (Wohlgemuth et al. 2010). Thus, it is reasonable to consider impaired autophagy a potential contributing factor to age-related muscle weakness. We have previously suggested that impaired autophagy in muscle may lead to deficits in sarcoplasmic reticulum (SR) function (Russ et al. 2012b, 2014). Release of Ca2+ from the SR is a critical process in muscle excitation–contraction (E–C) coupling, and could impair force generation, independent of muscle size (Ingalls et al. 1998). Indeed, several investigators have suggested that impairments of E–C coupling contribute to weakness in aging muscles, independent of changes in muscle size (Renganathan and Delbono 1998; Weisleder et al. 2006), and our laboratory has found that SR- Ca2+ release is impaired in aging gastrocnemius muscles that also exhibit reduced muscle quality (Russ et al. 2011, 2014).

Our hypothesis that age-related impairments of autophagy are contributing the loss of SR function is based on the idea that there are organelle-specific pathways of autophagy, including an endoplasmic reticulum (ER)-specific pathway (Bernales et al. 2007). Because the SR of skeletal muscle is an abundant specialized form of ER (Volpe et al. 1992), we speculate that there is an SR-specific autophagy in skeletal muscle, and that it is impaired with aging. In support of this concept, we have found increased markers of membrane damage and ER stress, in the SR of aging muscles (Russ et al. 2012b, 2014). Classical ER stress can result from several by-products of reduced autophagy, including: unfolded proteins, glycation, perturbations in redox status or altered membrane lipids (Nikolova-Karakashian and Rozenova 2010), factors which could all affect the aging SR. Indeed, we reported that markers of sphingolipid, specifically ceramide, metabolism are elevated in the SR of aging gastrocnemius muscles (Russ et al. 2014), supporting this latter mechanism. One might expect that increased “SR stress” would stimulate autophagy, in a classic feedback loop, but it has been suggested that aging compromises this adaptive response to stress (Naidoo 2009). We have found that aging gastrocnemius muscles exhibit reduced markers of autophagy, despite increases in stress markers, consistent with the idea of an age-related disruption of stress-autophagy feedback.

In contrast to the gastrocnemius, a reduced or delayed impact of aging on the soleus has been reported (Hagen et al. 2004), though deficits have been observed once the animals are very old (Carter et al. 2010; Rowan et al. 2011). Data from our laboratory are consistent with fewer age-related deficits in the soleus muscle, as we have reported no change in SR function and increased mass in the soleus of aging rats (Horner et al. 2011; Russ et al. 2011). In the present study, we extend these data by assessing contractile function (force and muscle quality) in the aging rat soleus. If problems with basal autophagy are indeed contributing to an age-related decline in SR function, then one would expect: (1) that muscle quality of the soleus would be maintained, and (2) that indices of impaired autophagy would be present in the gastrocnemius, but not the soleus muscle. Although studies have compared the responses of autophagy markers to exercise training and genetic manipulation in slow and fast-phenotype muscles of young mice (Lira et al. 2013; Yamada et al. 2012), such a muscle-specific comparison has not been made for the effect of aging. Accordingly, the present study examines the abundance of several markers of autophagy and related processes (i.e., stress and sphingolipid metabolism) in the gastrocnemius and soleus muscles of adult and aged rats. We hypothesize that markers of autophagy are reduced with aging in the gastrocnemius, but not soleus muscles, and that stress markers are elevated with aging in the gastrocnemius, but not soleus muscles.

Materials and methods

Animals and ethical approval

Adult (n = 5; 7–8 months) and aged (n = 5; 24–25 months), male F344/BN hybrid rats, representing an age of ~60 % maximum lifespan (Turturro et al. 1999), and one at which locomotor deficits have been observed (Horner et al. 2011), were used in the contractile experiments. Stored, frozen muscles from an additional 6 male rats of the same strain and ages (3 adult and 3 aged) that had been used in an earlier experiment testing a different muscle were also used in the immunoblotting experiments. All animals were housed in an environmentally controlled facility (12–12 h light–dark cycle, 22 °C), and had ad libitum access to standard rat chow (Prolab RMH 3000; calories provided by Protein, Fat and Carbohydrate: 26, 15 and 59 %, respectively) and water. Animal use and all procedures were approved by the Ohio University Institutional Animal Care and Use Committee, and the “Principles of laboratory animal care” (NIH publication No. 86–23, revised 1985), were followed throughout the study.

In situ contractile testing of the soleus

Contractile testing and tissue harvest was conducted at roughly the same time of day (early afternoon) for all animals, but food was not withdrawn for any period of time prior to testing. Animals were anesthetized (Ketamine/Xylazine 40:10 mg kg−1 body mass, IP) and mounted in a rigid frame to immobilize the leg and pelvis. Food was not withheld from animals prior to dissection and testing. Muscle dissection and sciatic nerve stimulation were the same as we have described previously (Russ et al. 2014), except that the soleus muscle was clamped in series with the force transducer. Length of the muscle–tendon complex was adjusted to produce optimal twitch force (l0). Muscle quality was calculated by expressing muscle force relative to the cross-sectional area (CSA) of the muscle, as we have previously described (Russ et al. 2014), except that the muscle length/fiber length ratio for the rat soleus was used. Following contractile testing, both simulated and un-stimulated medial gastrocnemius (MG) and soleus muscles were dissected from still-anesthetized animals, rinsed in ice-cold phosphate-buffered saline, blotted dry and weighed, then frozen in liquid N2 and stored at -80º C for later analysis. Animals were sacrificed while still anesthetized by an intracardiac injection of anesthesia (Euthasol, 100 mg kg−1) following tissue collection.

SDS-PAGE and western blotting

The unstimulated, frozen muscles were processed for electrophoresis, then subjected to SDS-PAGE and transferred to transferred to polyvinylidene fluoride (PVDF) membranes, as we have described previously (Russ et al. 2012b). Once transfer was complete, membranes were blocked in buffer (Odyssey, LI-COR, Lincoln, NE) at room temperature for 1 h, then incubated overnight at 4 °C with primary antibodies. Antibodies against Serine Palmitoyltransferase (SPT, ab23696), neutral sphingomyelinase (nSMase, ab131330), Glucose-regulated protein 78 (Grp78, ab108613), Protein tyrosine phosphatase 1b (PTP1b, ab52650), Beclin (ab79937), Autophagy related proteins 3, 4b and 7 (Atg 3, ab108251; Atg 4b, ab154843; Atg 7, ab133528 respectively) were purchased from Abcam (Cambridge, MA). The antibody against microtubule-associated protein 1 light chain 3 beta (LC3b, TA301542) was purchased from OriGene (Rockville, MD). The antibody against neutral ceramidase (ASAH, PA20575) was purchased from EMD Millipore/Calbiochem (Billerica, MA). Antibodies against ubiquitin-binding protein p62/sequestosome1 (p62, P0067) and apoptosis regulator Bcl2 (Bcl2, B9804) were purchased from Sigma-Aldrich (St. Louis, MO). The antibody against acid sphingomyelinase (aSMase, sc9817) was purchased from SCBT (Santa Cruz, CA). All primary antibodies were diluted 1:2,000 in blocking buffer plus Tween 20. After primary incubation, membranes were washed 5 × 5 min in tris-buffered saline plus Tween 20 (TBS-T) and incubated for 1 h at room temperature with appropriate secondary antibodies (LI-COR, Lincoln, NE) which were diluted in blocking buffer (1:10,000–1:20,000). Following secondary incubation, membranes were once again washed 5 × 5 min in TBS-T, then rinsed for 5 min with TBS. Membranes were dried in the dark overnight prior to scanning and densitometric band analysis with a LI-COR Odyssey system. After scanning, the membranes were stained with Coomassie Brilliant Blue R250 and within blot band intensities were normalized to total protein per lane determined from the stained, scanned membrane. The blots were all performed in duplicate. Known amounts of rabbit and mouse IgG were run on each gel as a standard for normalization of bands from different blots.

Myosin heavy chain analysis

Frozen portions of the stimulated MG and soleus muscles were processed to isolate myofibrillar proteins, as described previously (Russ et al. 2014). Myofibrillar proteins (10 µg/lane) were run overnight on 8 % gels containing 30 % glycerol (Mizunoya et al. 2008), then stained with Coomassie Brilliant Blue R250. Gels were scanned on a LI-COR Odyssey system, and bands corresponding to the major myosin heavy chain (MHC) isoforms were analyzed densitometrically, and the relative MHC abundance of each band was expressed as a percent of the total myosin per lane. This method cannot discriminate hybrid fibers; nor can it separate the effects of changes in fiber number from those of fiber size. However, it has been shown to correlate with histochemically-determined percent fiber-type area from muscle sections (Fry et al. 1994).

Statistical analyses

Animal masses were compared using unpaired t tests. Muscle mass and muscle:body mass ratios were compared using 2-way (age × muscle) analyses of variance (ANOVA), with muscle as a repeated factor. Soleus force- and muscle quality-frequency data were compared using a two-way (age × frequency) ANOVA with frequency as a repeated factor. Immunoblot data were compared using 2-way (age × muscle) ANOVAs, with muscle as a repeated factor, and are presented as arbitrary units, normalized to the young MG values. In the event of significant main effects or interactions, post hoc tests were performed using unpaired t tests. Percentages of the MHC isoforms were compared using unpaired t tests. Statistical analyses were conducted with SPSS and the threshold for significance set at P ≤ 0.05.


Animal and muscle mass

There were significant age-related increases in body mass and soleus wet weight (roughly 27 and 17 %, respectively), but not MG wet weight, which showed a reduction of ~10 %. The muscle mass:body mass ratio was reduced for both muscles with aging, but the difference was significant only for the MG (Table 1). Because we determined CSA based on optimal length, and performed contractile testing only on the soleus, we cannot report CSA for the MG. However, the aged soleus CSA was significantly (P = 0.050) greater than that of the adult soleus (0.108 ± 0.006 and 0.089 ± 0.004 cm2, respectively).
Table 1

Body & muscle masses


Adult (n = 8)

Aged (n = 8)

P value

Body mass (g)

426.5 (26.5)

540.3 (39.4)


MG mass (g)

1.150 (0.10)

1.035 (0.081)


MG mass/body mass (%)

0.271 (0.041)

0.192 (0.023)


MG MHC Isoform (%)

 Type IIa

3.7 (1.0)

5.7 (1.4)


 Type IIx

25.9 (4.1)

28.2 (4.4)


 Type IIb

64.8 (5.0)

54.9 (5.8)


 Type I

5.5 (1.2)

11.3 (1.8)


 Sol mass (g)

0.169 (0.010)

0.198 (0.009)


 Sol mass/body mass (%)

0.040 (0.003)

0.037 (0.003)


Sol MHC Isoform (%)

 Type IIa

7.8 (1.3)

5.1 (1.5)


 Type I

92.1 (1.2)

94.9 (1.4)


Soleus muscle contractility: The force-frequency relationships for adult and aged soleus muscles indicated an increase in force production in the aged animals (Fig. 1a), with no difference in muscle quality (Fig. 1b). Thus, the increased soleus mass seen in the aged animals (Table 1) translated to increased muscle force.
Fig. 1

Soleus contractile responses a Mean (±SE) force-frequency data from aging and adult soleus muscles. b Mean (±SE) muscle quality data from aging and adult soleus muscles. *Significant difference between age groups

Markers of autophagy

Abundance of our principal markers of autophagy exhibited marked effects of muscle activity (LC3b-II/I ratio, P = 0.009 and p62, P < 0.001), with lower levels in the soleus (Fig. 2). The LC3b-II/I ratio showed a significant age ×  muscle interaction, and post hoc testing indicated a significant age-related decrease in the ratio for the MG, and a trend for an age-related increase in the soleus muscle (P = 0.059). There were no significant main effects or interactions for abundance of either LC3b-I or -II, though there was a trend for an effect of muscle (P = 0.085) on LC3b-II abundance. For p62, post hoc testing indicated that effects were less robust than for the LC3b-II/I ratio, as there was only a trend (P = 0.087) for an age-related increase in p62 between adult and aged animals in the MG.
Fig. 2

Markers of Autophagy: Mean (± SE) of a LC3b-II/I ratio and p62, b LC3b-I and LC3b-II, c Beclin; d Atg3; e Atg4; f Atg7 and g Bcl2. Data are normalized to adult MG values. a Significant effect of age; m significant effect of muscle; x significant age X muscle interaction; *significant age difference within muscle; #trend for age difference within muscle. h Representative blots

The effects of age and muscle differed for several specific proteins that act upstream of LC3b and p62 in the autophagy pathway (Fig. 2). Significant main effects of muscle were observed both for Beclin and Bcl-2 (P = 0.003 and < 0.001, respectively), and a trend for an effect was present for Atg4b (P = 0.055). A significant main effect of age was present for Atg4b and Bcl-2 (P = 0.028 and 0.010, respectively). Significant age × muscle interactions were observed for Atg3 and Beclin (P = 0.029 and 0.016 respectively). Post-hoc testing for proteins exhibiting significant effects or interactions revealed a general pattern for significant increases in abundance to occur in the MG, but not the soleus, though there was a trend (P = 0.077) for Bcl-2 to increase with age in the soleus. The major exception was Atg7, for which no significant effects or interactions were observed.

Markers of ER/SR stress and sphingolipid metabolism

We examined two established ER stress proteins: Grp78, which we have previously found (Russ et al. 2014) to be elevated in the SR of aging rat MG muscles, and PTP1b. Both proteins exhibited similar patterns of abundance, with increased abundance with age in the MG, but not the soleus, though the soleus tended to have higher overall levels of both proteins. For Grp78, there was a significant age × muscle interaction (P = 0.046) and a trend for an effect of muscle group (P = 0.072). For PTP1b, there was a significant effect of muscle (P = 0.014) and a significant age × muscle interaction (P = 0.026). Post-hoc t tests were significant for age differences in the MG (P = 0.037 and 0.004 for Grp78 and PTP1b, respectively), but not the soleus (P = 0.681 and 0.702 for Grp78 and PTP1b, respectively).

We examined proteins associated with de novo and salvage pathways of ceramide synthesis (SPT, nSMase and aSMase, respectively) and breakdown (Neutral Ceramidase; ASAH). There were no significant effects of age or muscle on SPT, though the age × muscle interaction was significant (P = 0.021), due to an age-related increase in the MG, but not the soleus (Fig. 3). For nSMase, there was a significant effect of muscle (P = 0.010) and a significant age × muscle interaction (P = 0.022), due to the age-related increase in abundance in the MG, but not the soleus. There was also a trend (P = 0.095) for a main effect of age. No significant effect of age or age × muscle interaction was present for ASAH or aSMase, though there was a trend (P = 0.082) for ASAH abundance to increase with age. The effect of muscle was significant both ASAH and ASMase however, with abundance of both proteins significantly less in the soleus than in the MG.
Fig. 3

Markers of ER/SR stress & sphingolipid metabolism: Mean (+SE) abundance of a Grp78, b PTP1b, c Acid Sphingomyelinase, d Neutral Sphingomyelinase, e Serine Palmitoyltransferase and f Neutral Ceramidase. Insets representative blots. Values are normalized to the Adult, MG levels. a Significant effect of age; m significant effect of muscle; x significant age X muscle interaction; *significant age difference within muscle; #trend for age difference within muscle

MHC composition

The soleus exhibited no age-differences in the relative content of the principal MHC isoforms, consistent with previous findings in rats of this strain and age range. In the MG, the content of Type I myosin was significantly greater in the aged animals. No differences were present for any of the major Type II isoforms.


This study of the muscle specificity of aging demonstrates that aging, in general, does not adversely affect contractile function of the soleus and has greater effects on markers of signaling within the autophagic, ER/SR stress and sphingolipid metabolic pathways in the MG than in the soleus muscle. The overall pattern of protein abundance studied here suggests an impairment of autophagy in the MG, despite increases in markers of upstream signaling of autophagy, with far fewer changes in the soleus. Aging was also found to increase markers of ER/SR stress and alter markers of sphingolipid metabolism suggestive of increased muscle ceramides in the MG, but not the soleus.

We found here, as we have previously (Russ et al. 2011), that the soleus exhibits hypertrophy in rats of this strain and age, though the soleus:body mass ratio does decline with aging. Our results indicate that soleus muscle quality is preserved at this early stage of aging and, combined with the increase muscle size, results in an age-related increase in force generation. We speculate, because the age-related changes in soleus and body mass are so similar (Table 1), that the increase in soleus mass is due to a hypertrophic stimulus to this postural muscle from the constant demand of carrying the increased body mass. The soleus muscle in rats older than those studied here does appear to exhibit loss of both mass and function (Carter et al. 2010; Rowan et al. 2011), possibly due to late-life decline in body mass reducing the postural load or a further reduction of physical activity (though all animals studied here had only standard cage activity). However, a trend for increased myofiber hypertrophy has also been reported in 32-month old rats (Garvey et al. 2014). It will be interesting to see in future work if the soleus exhibits impairments in autophagy in animals of ages at which soleus force and mass decline.

Just as the effects of age on the contractile function of the soleus diverged markedly from those we have previously reported for the MG, we found here that the effect of abundance of markers of autophagy also differed between the two muscles. The general pattern was that the aging MG muscle exhibited changes in protein abundance consistent with impairment of basal autophagy, but the aging soleus did not. The intermuscular differences (i.e., MG vs. soleus) observed were similar to those reported by Lira and colleagues (Lira et al. 2013 in some regards (reduced p62 in soleus vs. MG), see Fig. 1), but not others (reduced LC3b-II/I ratio, no difference in Atg7). However, it is important to note that Lira and colleagues studied mice. Although the fiber type of the mixed rat MG samples we studied here is fairly similar to the mouse plantaris studied by Lira et al., the soleus of rat soleus exhibits a much greater overall Type I myosin heavy chain content than does that of mouse (Bloemberg and Quadrilatero 2012). The greater intermuscular difference in fiber type may have accounted for some of the differences in protein abundance between the two studies.

Despite the observed tendency for age to induce changes in LC3b and p62 consistent with a reduction in basal autophagy in the MG, a general age-related increase in abundance of upstream activators of autophagy was seen in the MG, but not the soleus. The major exception was Atg7, which did not exhibit any effects of age or muscle, similar to what has been reported elsewhere in aging, fast-twitch muscles (Wohlgemuth et al. 2010). Interestingly, impaired basal and stress-induced autophagy have been reported in the liver of Atg7-deficient transgenic mice (Komatsu et al. 2005), and young Atg7 knockout animals exhibit reductions in both muscle mass and muscle quality (Masiero et al. 2009). Of note, these animals also manifested SR distension, consistent with our hypothesis that age-associated impairment in autophagy contributes to loss of muscle quality by affecting SR function. Though less pronounced than the knockout of Atg7, it is tempting to speculate that the failure of Atg7 to increase with age may be a “bottleneck” where the link between upstream signaling and autophagic activity is impaired in aging muscle. Perhaps in even older animals, where more extensive loss of muscle function occurs, we would observe a reduction in Atg7.

As ER stress is a factor known to induce autophagy (Madaro et al. 2013), we evaluated a pair of ER stress markers (Grp78 and PTP1b). Although such stress is normally thought to stimulate autophagy to act to relieve the stress, it has been suggested that this normal feedback is disrupted in aging (Naidoo 2009; Wohlgemuth et al. 2010). In the present study, we found that stress markers were elevated in the aging MG, but not soleus (Fig. 3a, b). The age-related responses from the MG are comparable to those we have observed previously in SR samples isolated from the MG, suggesting that SR stress is a factor. It could be that the soleus is subject to fewer age-related stressors, but the overall tendency for stress markers to be higher in the adult soleus suggests this is not the case. It may be that the more consistent loading of the soleus to maintain normal posture maintains a level of stress that continues to stimulate autophagy during aging, maintaining muscle function. Consistent with this hypothesis, we have previously shown that even modest volitional exercise can restore markers of autophagy, at least in aged, but not senescent rats (Russ et al. 2012b). Interestingly, Wohlgemuth and colleagues (Wohlgemuth et al. 2010) have reported that exercise in aging animals, with or without caloric restriction, can increase muscle Atg7, which was the principal upstream regulator of autophagy that did not increase with aging in the sedentary animals studied here.

One factor shown to promote ER/SR stress is perturbation of cellular lipid composition, including increased ceramide (Terman and Brunk 2006; Volmer et al. 2013; Zha et al. 2013). Our results show that 2 key enzymes related to sphingolipid metabolism, serine palmitoyltranferase and neutral sphingomyelinase (nSMase) are elevated with aging in the MG, but not the soleus, although acid sphingomyelinase (aSMase) and neutral ceramidase, which breaks down ceramide, were not (Fig. 3). This pattern of abundance would be expected to increase muscle ceramide levels, and is similar to what we have reported previously from SR vesicles isolated from the MG (Russ et al. 2014). Increased ceramide is associated with increased ER stress (Nikolova-Karakashian and Rozenova 2010) and reduced muscle quality (Ferreira et al. 2010), as is administration of exogenous sphingomyelinase (Bost et al. 2015; Ferreira et al. 2010). Also of note, increased ceramide levels occur following Atg7 inhibition to block autophagy in a cultured cancer cell line (Separovic et al. 2010). These data are all consistent with our observations of reduced muscle quality in the MG, but not the soleus (Fig. 1). Interestingly, aSMase is generally associated with lysosomes (Perrotta and Clementi 2010), whereas nSMase is known to localize at the triad junction in skeletal muscle (Sabbadini et al. 1999), suggesting perhaps that nSMase abundance and activity may preferentially influence SR lipid composition and stress or be influenced by selective SR-specific autophagy. This would support our current hypothesis that impaired SR function is a key factor in the age-related loss of muscle quality. In further support of this working model, byproducts of ceramide metabolism have been shown to directly impair the SR Ca2+-release channel (Sharma et al. 2000). Also of note, the SR of the soleus has been found to contain higher levels of sphingomyelin than fast-twitch muscles (Borchman et al. 1999), perhaps accounting for the higher levels of nSMase observed here in the young soleus.

An age-related shift in MHC composition is often reported in aging muscles (Purves-Smith et al. 2014), though we have previously found that such changes are relatively minor in rats of this strain and age-range (Russ et al. 2011, 2014). In the present study, the MHC composition of the young muscles was comparable to that reported elsewhere (Eng et al. 2008; Staron et al. 1999), and the finding that the MG exhibited only minor shifts in MHC profile, while the soleus exhibited no significant changes is consistent with our earlier studies. In general, it seems unlikely that increased Type I MHC in the MG can account for the muscle-specific differences in protein abundance observed in the present study, given that MHC I accounts for such a small percentage of the total MHC in this muscle. However, the fact that the MG shows changes in MHC with aging, while the soleus does not, is suggestive of differences in intracellular signaling and processing between the two muscles that occurs with aging.

The general pattern of age-related increases in protein abundance occurring in the MG, but not the soleus, is consistent with many of our hypotheses. However, in some instances, this must be reconciled with proteins whose levels of abundance in the soleus muscle, regardless of age, exceeded those in the adult MG. One such protein was nSMase, whereas aSMase abundance was lower in the soleus than the MG. Of note, aSMase is heavily localized in lysosomes, and one study has reported that Type I fibers, which predominate in the soleus, contain less lysosome-associated membrane protein 1 (LAMP1) than type II fibers (Drost et al. 2008), suggesting that differences in the number of lysosomes may be playing a role. It may be that the elevated nSMase in the soleus occurs to maintain an adequate level of the salvage pathway of ceramide synthesis despite the reduced abundance of aSMase. A clearer picture may emerge in future work that examines activity, as well as abundance, of these enzymes.

Both stress-associated proteins we examined (Grp78 and PTP1b) also exhibited elevated abundance in the soleus (Fig. 3). Both of these proteins are associated with unfolded protein response, and it could be that more frequent level of regular contractile activity in the soleus produces more protein modifications that need to be addressed. Further, the continued demand on the soleus throughout aging may maintain these higher levels. Some limited support for this hypothesis comes from our previous work that demonstrated that volitional exercise increased some classic stress markers, though it reduced others and improved autophagy (Russ et al. 2012b). In addition, Grp78 is closely related to heat shock protein 70 (Metskas et al. 2010), and thus may exhibit a similarly increased abundance in slower-phenotype muscles (Locke et al. 1994).

Of the autophagy-related proteins we examined, only Bcl-2 exhibited this pattern of elevated abundance in the soleus (Fig. 2g). Although Bcl-2 has been shown to modify autophagy, it is better known for its anti-apoptotic properties, and the greater abundance in the soleus seen here may reflect a greater resistance to apoptosis in this muscle relative to the MG (Libera et al. 1999). In addition, a previous study has shown a similarly high level of Bcl-2 abundance in slow- vs. fast-phenotype muscles (McMillan and Quadrilatero 2011).

Some limitations of the present study must be acknowledged. We studied animals in early-stage aging both to be consistent with our earlier studies in the MG, and because we have observed impairments in both muscle function and locomotor performance in rats of this strain and age (Horner et al. 2011; Russ et al. 2011, 2014). Other investigators have reported that declines in soleus function occur at more advanced ages (Carter et al. 2010; Rowan et al. 2011). In future work, we plan to examine older animals and determine if we also observe such a decline in force production, and whether or not this decline is associated with age-related changes in markers of autophagy, similar to those we have reported in the MG. In addition, we studied only male animals, again to be consistent with our previous studies. However, the possibility remains open that aging females may not have shown the age-related increase in muscle force seen here in males. Data indicate that aging females may be more vulnerable to loss of muscle mass than males (Akima et al. 2001), and more a recent study suggests that muscle quality is a more important predictor of physical function in aging men than women (Fragala et al. 2012). In addition, significantly higher levels of Grp78 in the biceps brachii of females have been reported in young animals (Metskas et al. 2010), suggesting the possibility of reduced age-related changes in females (similar to what we observed in the soleus here (Fig. 3a)). All rats were tested at a time of day (early afternoon) when, being nocturnal, they may not have eaten for some time. Fasting is known to affect autophagosome production. Though our animals were not fasted prior to testing, we cannot rule out some potential effect of dietary intake on our markers of autophagy, though this effect should have washed out across groups, testing and tissue harvest occurred at roughly the same times. Moreover, skeletal muscle appears to respond to fasting in a much more prolonged manner than other tissues (Sandri 2010), and thus may not exhibit extensive acute fluctuation with normal dietary intake. Finally, though much of our speculation in regards to the effects of age-related impairments in autophagy on muscle quality centers on potential effects on the SR and E–C coupling, it might be that it is contributing to problems at the neuromuscular junction (NMJ) as well. A number of morphological changes occur in the aging NMJ that might well impair its function (for recent review, c.f. (Rudolf et al. 2014)). It has also been shown recently that inhibition of autophagy impairs NMJ function (Carnio et al. 2014).


This study demonstrates that muscle quality in the aging soleus is maintained at adult levels, in contrast to our previous studies of the MG. Because soleus size also increases, its force production is greater in the aged vs. the adult animals. In addition, this study is the first, to our knowledge, to compare signaling within the autophagy pathway in fast and slow muscles of adult and aging animals. These data indicate that markers of autophagy, ER/SR stress and sphingolipid metabolism are better maintained at adult levels in the aging soleus than the aging MG in male rats. The divergent responses of the soleus and MG muscles to increasing age with regard to muscle quality and autophagy are consistent with our proposed mechanism of impaired autophagy contributing to unrelieved SR stress that impairs SR function, subsequently contributing to reduced muscle quality, possibly through impairments of E–C coupling, NMJ dysfunction, or both.



Financial assistance for this project was provided to Dr. Russ by the Ohio Musculoskeletal Neurological Institute. Ms. Boyd and Ms. McCorkle were supported by College of Health Sciences and Professions Graduate Assistantships throughout the project. The authors wish to thank Drs. Sean Garvey and Noah Weisleder for critical reviews of earlier drafts of this manuscript.

Compliance with ethical standards

Conflict of interest

None of the authors is aware of any conflict of interest regarding any aspect of the work reported here.


  1. Akima H et al (2001) Muscle function in 164 men and women aged 20–84 years. Med Sci Sports Exerc 33:220–226CrossRefPubMedGoogle Scholar
  2. Bean JF, Kiely DK, Herman S, Leveille SG, Mizer K, Frontera WR, Fielding RA (2002) The relationship between leg power and physical performance in mobility-limited older people. J Am Geriatr Soc 50:461–467CrossRefPubMedGoogle Scholar
  3. Bernales S, Schuck S, Walter P (2007) ER-phagy: selective autophagy of the endoplasmic reticulum. Autophagy 3:285–287CrossRefPubMedGoogle Scholar
  4. Bloemberg D, Quadrilatero J (2012) Rapid determination of myosin heavy chain expression in rat, mouse, and human skeletal muscle using multicolor immunofluorescence analysis. PLoS ONE 7:e35273. doi:10.1371/journal.pone.0035273 PubMedCentralCrossRefPubMedGoogle Scholar
  5. Borchman D, Tang D, Yappert MC (1999) Lipid composition, membrane structure relationships in lens and muscle sarcoplasmic reticulum membranes. Biospectroscopy 5:151–167. doi:10.1002/(SICI)1520-6343(1999)5:3<151:AID-BSPY5>3.0.CO;2-D CrossRefPubMedGoogle Scholar
  6. Bost ER, Frye GS, Ahn B, Ferreira LF (2015) Diaphragm dysfunction caused by sphingomyelinase requires the p47(phox) subunit of NADPH oxidase. Respir Physiol Neurobiol 205:47–52. doi:10.1016/j.resp.2014.10.011 CrossRefPubMedGoogle Scholar
  7. Carnio S et al (2014) Autophagy impairment in muscle induces neuromuscular junction degeneration and precocious aging. Cell Rep 8:1509–1521. doi:10.1016/j.celrep.2014.07.061 PubMedCentralCrossRefPubMedGoogle Scholar
  8. Carter EE, Thomas MM, Murynka T, Rowan SL, Wright KJ, Huba E, Hepple RT (2010) Slow twitch soleus muscle is not protected from sarcopenia in senescent rats. Exp Gerontol. doi:10.1016/j.exger.2010.04.001 PubMedCentralGoogle Scholar
  9. Degens H, Alway SE (2006) Control of muscle size during disuse, disease, and aging. Int J Sports Med 27:94–99. doi:10.1055/s-2005-837571 CrossRefPubMedGoogle Scholar
  10. Drost MR et al (2008) Both type 1 and type 2a muscle fibers can respond to enzyme therapy in Pompe disease. Muscle Nerve 37:251–255. doi:10.1002/mus.20896 CrossRefPubMedGoogle Scholar
  11. Eng CM, Smallwood LH, Rainiero MP, Lahey M, Ward SR, Lieber RL (2008) Scaling of muscle architecture and fiber types in the rat hindlimb. J Exp Biol 211:2336–2345. doi:10.1242/jeb.017640 CrossRefPubMedGoogle Scholar
  12. Ferreira LF, Moylan JS, Gilliam LA, Smith JD, Nikolova-Karakashian M, Reid MB (2010) Sphingomyelinase stimulates oxidant signaling to weaken skeletal muscle and promote fatigue. Am J Physiol Cell Physiol 299:C552–560. doi:10.1152/ajpcell.00065.2010 PubMedCentralCrossRefPubMedGoogle Scholar
  13. Fragala MS, Clark MH, Walsh SJ, Kleppinger A, Judge JO, Kuchel GA, Kenny AM (2012) Gender differences in anthropometric predictors of physical performance in older adults. Gend Med 9:445–456. doi:10.1016/j.genm.2012.10.004 PubMedCentralCrossRefPubMedGoogle Scholar
  14. Fry AC, Allemeier CA, Staron RS (1994) Correlation between percentage fiber type area and myosin heavy chain content in human skeletal muscle. Eur J Appl Physiol Occup Physiol 68:246–251CrossRefPubMedGoogle Scholar
  15. Garvey SM et al (2014) Metabolomic profiling reveals severe skeletal muscle group-specific perturbations of metabolism in aged FBN rats. Biogerontology 15:217–232. doi:10.1007/s10522-014-9492-5 PubMedCentralCrossRefPubMedGoogle Scholar
  16. Hagen JL, Krause DJ, Baker DJ, Fu MH, Tarnopolsky MA, Hepple RT (2004) Skeletal muscle aging in F344BN F1-hybrid rats: i. Mitochondrial dysfunction contributes to the age-associated reduction in VO2max. J Gerontol A Biol Sci Med Sci 59:1099–1110CrossRefPubMedGoogle Scholar
  17. Horner AM, Russ DW, Biknevicius AR (2011) Effects of early-stage aging on locomotor dynamics and hindlimb muscle force production in the rat. J Exp Biol 214:3588–3595. doi:10.1242/jeb.055087 CrossRefPubMedGoogle Scholar
  18. Hubbard VM, Valdor R, Macian F, Cuervo AM (2012) Selective autophagy in the maintenance of cellular homeostasis in aging organisms. Biogerontology 13:21–35. doi:10.1007/s10522-011-9331-x CrossRefPubMedGoogle Scholar
  19. Ingalls CP, Warren GL, Williams JH, Ward CW, Armstrong RB (1998) E–C coupling failure in mouse EDL muscle after in vivo eccentric contractions. J Appl Physiol 85:58–67PubMedGoogle Scholar
  20. Kim YA, Kim YS, Oh SL, Kim HJ, Song W (2013) Autophagic response to exercise training in skeletal muscle with age. J Physiol Biochem 69:697–705. doi:10.1007/s13105-013-0246-7 CrossRefPubMedGoogle Scholar
  21. Komatsu M et al (2005) Impairment of starvation-induced and constitutive autophagy in Atg7-deficient mice. J Cell Biol 169:425–434. doi:10.1083/jcb.200412022 PubMedCentralCrossRefPubMedGoogle Scholar
  22. Lexell J, Taylor CC, Sjostrom M (1988) What is the cause of the ageing atrophy? Total number, size and proportion of different fiber types studied in whole vastus lateralis muscle from 15- to 83-year-old men. J Neurol Sci 84:275–294CrossRefPubMedGoogle Scholar
  23. Libera LD, Zennaro R, Sandri M, Ambrosio GB, Vescovo G (1999) Apoptosis and atrophy in rat slow skeletal muscles in chronic heart failure. Am J Physiol 277:C982–986PubMedGoogle Scholar
  24. Lira VA et al (2013) Autophagy is required for exercise training-induced skeletal muscle adaptation and improvement of physical performance. Faseb J 27:4184–4193. doi:10.1096/fj.13-228486 PubMedCentralCrossRefPubMedGoogle Scholar
  25. Locke M, Atkinson BG, Tanguay RM, Noble EG (1994) Shifts in type I fiber proportion in rat hindlimb muscle are accompanied by changes in HSP72 content. Am J Physiol 266:C1240–1246PubMedGoogle Scholar
  26. Madaro L, Marrocco V, Carnio S, Sandri M, Bouche M (2013) Intracellular signaling in ER stress-induced autophagy in skeletal muscle cells. Faseb J 27:1990–2000. doi:10.1096/fj.12-215475 CrossRefPubMedGoogle Scholar
  27. Masiero E et al (2009) Autophagy is required to maintain muscle mass. Cell Metab 10:507–515. doi:10.1016/j.cmet.2009.10.008 CrossRefPubMedGoogle Scholar
  28. McMillan EM, Quadrilatero J (2011) Differential apoptosis-related protein expression, mitochondrial properties, proteolytic enzyme activity, and DNA fragmentation between skeletal muscles. Am J Physiol Regul Integr Comp Physiol 300:R531–543. doi:10.1152/ajpregu.00488.2010 CrossRefPubMedGoogle Scholar
  29. Metskas LA, Kulp M, Scordilis SP (2010) Gender dimorphism in the exercise-naive murine skeletal muscle proteome. Cell Mol Biol Lett 15:507–516. doi:10.2478/s11658-010-0020-6 CrossRefPubMedGoogle Scholar
  30. Metter EJ, Talbot LA, Schrager M, Conwit R (2002) Skeletal muscle strength as a predictor of all-cause mortality in healthy men. J Gerontol A Biol Sci Med Sci 57:B359–365CrossRefPubMedGoogle Scholar
  31. Mizunoya W, Wakamatsu J, Tatsumi R, Ikeuchi Y (2008) Protocol for high-resolution separation of rodent myosin heavy chain isoforms in a mini-gel electrophoresis system. Anal Biochem 377:111–113. doi:10.1016/j.ab.2008.02.021 CrossRefPubMedGoogle Scholar
  32. Naidoo N (2009) ER and aging-Protein folding and the ER stress response. Ageing Res Rev 8:150–159. doi:10.1016/j.arr.2009.03.001 CrossRefPubMedGoogle Scholar
  33. Nikolova-Karakashian MN, Rozenova KA (2010) Ceramide in stress response. Adv Exp Med Biol 688:86–108CrossRefPubMedGoogle Scholar
  34. Perrotta C, Clementi E (2010) Biological roles of acid and neutral sphingomyelinases and their regulation by nitric oxide. Physiology 25:64–71. doi:10.1152/physiol.00048.2009 CrossRefPubMedGoogle Scholar
  35. Purves-Smith FM, Sgarioto N, Hepple RT (2014) Fiber typing in aging muscle. Exerc Sport Sci Rev 42:45–52. doi:10.1249/JES.0000000000000012 CrossRefPubMedGoogle Scholar
  36. Renganathan M, Delbono O (1998) Caloric restriction prevents age-related decline in skeletal muscle dihydropyridine receptor and ryanodine receptor expression. FEBS Lett 434:346–350CrossRefPubMedGoogle Scholar
  37. Rowan SL, Purves-Smith FM, Solbak NM, Hepple RT (2011) Accumulation of severely atrophic myofibers marks the acceleration of sarcopenia in slow and fast twitch muscles. Exp Gerontol 46:660–669. doi:10.1016/j.exger.2011.03.005 PubMedGoogle Scholar
  38. Rudolf R, Khan MM, Labeit S, Deschenes MR (2014) Degeneration of neuromuscular junction in age and dystrophy. Front Aging Neurosci 6:99. doi:10.3389/fnagi.2014.00099 PubMedCentralCrossRefPubMedGoogle Scholar
  39. Russ DW, Grandy JS, Toma K, Ward CW (2011) Ageing, but not yet senescent, rats exhibit reduced muscle quality and sarcoplasmic reticulum function. Acta Physiol (Oxf) 201:391–403. doi:10.1111/j.1748-1716.2010.02191.x CrossRefGoogle Scholar
  40. Russ DW, Gregg-Cornell K, Conaway MJ, Clark BC (2012a) Evolving concepts on the age-related changes in “muscle quality”. J Cachexia Sarcopenia Muscle 3:95–109. doi:10.1007/s13539-011-0054-2 PubMedCentralCrossRefPubMedGoogle Scholar
  41. Russ DW, Krause J, Wills A, Arreguin R (2012b) “SR stress” in mixed hindlimb muscles of aging male rats. Biogerontology 13:547–555. doi:10.1007/s10522-012-9399-y CrossRefPubMedGoogle Scholar
  42. Russ DW, Wills AM, Boyd IM, Krause J (2014) Weakness. SR function and stress in gastrocnemius muscles of aged male rats Exp Gerontol 50:40–44. doi:10.1016/j.exger.2013.11.018 PubMedGoogle Scholar
  43. Sabbadini RA, Danieli-Betto D, Betto R (1999) The role of sphingolipids in the control of skeletal muscle function: a review. Ital J Neurol Sci 20:423–430CrossRefPubMedGoogle Scholar
  44. Sandri M (2010) Autophagy in skeletal muscle. FEBS Lett 584:1411–1416. doi:10.1016/j.febslet.2010.01.056 CrossRefPubMedGoogle Scholar
  45. Separovic D, Kelekar A, Nayak AK, Tarca AL, Hanada K, Pierce JS, Bielawski J (2010) Increased ceramide accumulation correlates with downregulation of the autophagy protein ATG-7 in MCF-7 cells sensitized to photodamage. Arch Biochem Biophys 494:101–105. doi:10.1016/ PubMedCentralCrossRefPubMedGoogle Scholar
  46. Sharma C, Smith T, Li S, Schroepfer GJ Jr, Needleman DH (2000) Inhibition of Ca2+ release channel (ryanodine receptor) activity by sphingolipid bases: mechanism of action. Chem Phys Lipids 104:1–11CrossRefPubMedGoogle Scholar
  47. Staron RS, Kraemer WJ, Hikida RS, Fry AC, Murray JD, Campos GE (1999) Fiber type composition of four hindlimb muscles of adult Fisher 344 rats. Histochem Cell Biol 111:117–123CrossRefPubMedGoogle Scholar
  48. Terman A, Brunk UT (2006) Oxidative stress, accumulation of biological ‘garbage’, and aging. Antioxid Redox Signal 8:197–204. doi:10.1089/ars.2006.8.197 CrossRefPubMedGoogle Scholar
  49. Turturro A, Witt WW, Lewis S, Hass BS, Lipman RD, Hart RW (1999) Growth curves and survival characteristics of the animals used in the Biomarkers of Aging Program. J Gerontol A Biol Sci Med Sci 54:B492–501CrossRefPubMedGoogle Scholar
  50. Visser M et al (2000) Change in muscle mass and muscle strength after a hip fracture: relationship to mobility recovery. J Gerontol A Biol Sci Med Sci 55:M434–440CrossRefPubMedGoogle Scholar
  51. Volmer R, van der Ploeg K, Ron D (2013) Membrane lipid saturation activates endoplasmic reticulum unfolded protein response transducers through their transmembrane domains. Proc Natl Acad Sci USA 110:4628–4633. doi:10.1073/pnas.1217611110 PubMedCentralCrossRefPubMedGoogle Scholar
  52. Volpe P, Villa A, Podini P, Martini A, Nori A, Panzeri MC, Meldolesi J (1992) The endoplasmic reticulum-sarcoplasmic reticulum connection: distribution of endoplasmic reticulum markers in the sarcoplasmic reticulum of skeletal muscle fibers. Proc Natl Acad Sci USA 89:6142–6146PubMedCentralCrossRefPubMedGoogle Scholar
  53. Weisleder N et al (2006) Muscle aging is associated with compromised Ca2+ spark signaling and segregated intracellular Ca2+ release. J Cell Biol 174:639–645PubMedCentralCrossRefPubMedGoogle Scholar
  54. Wohlgemuth SE, Seo AY, Marzetti E, Lees HA, Leeuwenburgh C (2010) Skeletal muscle autophagy and apoptosis during aging: effects of calorie restriction and life-long exercise. Exp Gerontol 45:138–148. doi:10.1016/j.exger.2009.11.002 PubMedCentralCrossRefPubMedGoogle Scholar
  55. Yamada E, Bastie CC, Koga H, Wang Y, Cuervo AM, Pessin JE (2012) Mouse skeletal muscle fiber-type-specific macroautophagy and muscle wasting are regulated by a Fyn/STAT3/Vps34 signaling pathway. Cell Rep 1:557–569. doi:10.1016/j.celrep.2012.03.014 PubMedCentralCrossRefPubMedGoogle Scholar
  56. Zha BS et al (2013) HIV protease inhibitors disrupt lipid metabolism by activating endoplasmic reticulum stress and inhibiting autophagy activity in adipocytes. PLoS One 8:e59514. doi:10.1371/journal.pone.0059514 PubMedCentralCrossRefPubMedGoogle Scholar

Copyright information

© Springer Science+Business Media Dordrecht 2015

Authors and Affiliations

  • David W. Russ
    • 1
    • 2
  • Iva M. Boyd
    • 1
  • Katherine M. McCoy
    • 1
  • Katherine W. McCorkle
    • 1
  1. 1.Laboratory for Integrative Muscle Biology, Division of Physical Therapy, School of Rehabilitation & Communication SciencesOhio UniversityAthensUSA
  2. 2.Ohio Musculoskeletal & Neurological Institute (OMNI)Heritage College of Osteopathic MedicineAthensUSA

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