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The Journals of Gerontology Series A: Biological Sciences and Medical Sciences 61:3-13 (2006)
© 2006 The Gerontological Society of America

Regulation of Skeletal Muscle Mitochondrial Content During Aging

Carrie N. Lyons1, Odile Mathieu-Costello2 and Christopher D. Moyes1,

1 Department of Biology, Queen's University, Kingston, Ontario, Canada.
2 Department of Medicine, University of California San Diego.

Address correspondence to Chris Moyes, PhD, Department of Biology, Queen's University, Kingston, Ontario, Canada, K7L 3N6. E-mail: moyesc{at}biology.queensu.ca


   Abstract
Top
 Abstract
Methods
 Results
 Discussion
 References
 
Mitochondrial content of skeletal muscle varies among fiber types, and changes in complex ways during aging. We evaluated the regulatory origins of differences in mitochondrial content among muscles of varied fiber type in F344xBNF1 rats, and how these regulatory patterns are altered with aging. In adult (12 month) animals we found that units citrate synthase (CS)/g tissue, a marker for mitochondrial content, varied ~3-fold among 10 skeletal muscles. Stoichiometric relationships between CS and isocitrate dehydrogenase, aconitase, and cytochrome c oxidase were generally preserved across fiber types. Among the 10 muscles of adult rats, CS content correlated with nuclear content (R2 = 0.36). Muscles differed widely in CS messenger RNA (mRNA)/DNA (an index of variation in transcriptional regulations) and units CS/CS mRNA (an index of variation in posttranscriptional regulations). All muscles of aged rats (35 months) showed an increase in mg DNA/g, suggestive of atrophy. Age-dependent declines in units CS/DNA were accompanied by reductions in CS mRNA/DNA and/or units CS/CS mRNA, depending on muscle fiber type. Thus, declines in units CS/DNA with age appeared to be due to transcriptional as well as translational variations. Differences in mitochondrial content among muscle fiber types and age groups may arise from variations in nuclear content and posttranscriptional processes, as well as transcriptional regulation.

Mitochondria play a pivotal role in muscle cell physiology, primarily by providing adenosine triphosphate via oxidative phosphorylation. Variations in muscle mitochondrial content are observed throughout the lifetime of an animal. During muscle differentiation, mitochondrial content increases 5-fold (1). When adult phenotypes are established, slow-twitch oxidative muscles contain about 5 times more mitochondria than do fast-twitch glycolytic muscles, as measured by citrate synthase (CS) activity levels (2). Mitochondrial content remains plastic in adult muscle, changing in response to environmental stress (3) and physiological conditions (4,5). The aging process, for example, can cause a decline in mitochondrial content, particularly in muscles that undergo atrophy (6). These metabolic changes, in combination with declines in contractile function, lead to a loss of muscle strength and endurance (7).

Although much is known about the regulatory basis of distinct contractile phenotypes, determinants of the metabolic phenotype in different fiber types and physiological states are less clear. Several lines of evidence suggest that differences in mitochondrial content among muscles of varied fiber type may be due to a transcriptional network including the coactivator PGC-1{alpha} (peroxisome proliferator-activated receptor-{gamma} coactivator) (8). For example, overexpression of PGC-1{alpha} in cultured muscle cells and fat cells leads to mitochondrial proliferation (9,10), and muscles rich in mitochondrial enzymes also possess higher levels of PGC-1{alpha} (11). Despite the research focus on these transcriptional regulators, it is not yet established if steady-state levels of muscle mitochondria are regulated primarily by transcriptional controls.

Understanding the mechanistic basis of variations in mitochondrial content among muscles is complicated by many morphological differences that influence functional relationships. For example, the striated muscles differ in nuclear content. Nuclear content of slow-twitch muscle is twice that of fast-twitch muscle, based on ultrastructural analyses (12–16) or biochemical analyses (mg DNA/g tissue) (17). Changes in the nuclear content of skeletal muscle are also reported to occur during the aging process (18–21). Such variations in nuclear content among fiber types and during aging have important influences on the genetic control of the muscle phenotype (22). Superimposed on these morphological differences are regulatory controls at the level of transcription and translation. Variations in any or all of these parameters (i.e., nuclear content, transcriptional and translational processes) can influence the molecular composition of skeletal muscle, e.g., mitochondrial content.

In this study, we compare mitochondrial enzyme and gene regulatory properties in a spectrum of skeletal muscles in adult (12 months), old (24 months), and very old (35–36 months) rats. We focus on analysis of CS activity as a marker for mitochondrial content, but also analyze activities of other mitochondrial enzymes to assess changes in enzyme stoichiometries, which often occur in muscle with age (23–25). We compare CS activity, CS messenger RNA (mRNA) levels, and DNA content across 10 skeletal muscles that vary in CS activity over a 5-fold range (2). Because variations in muscle mitochondrial content are often thought to arise through transcriptional regulation (e.g., by PGC-1{alpha} and other transcriptional effectors), we investigate whether CS enzyme content parallels CS mRNA levels. However, because multiple factors can influence the molecular composition of skeletal muscle, we also evaluate the importance of nuclear content and posttranscriptional processes on mitochondrial abundance across muscles and during aging. We hypothesize that activity levels of CS, an enzyme widely held to be transcriptionally regulated (26), should covary with CS mRNA levels among fiber types and during aging.


   METHODS
Top
 Abstract
 Methods
Results
 Discussion
 References
 
Animal Care
Male F1 hybrid F344 x Brown Norway rats (Harlan Sprague Dawley Inc., Indianapolis, IN) were housed 2–3 animals per cage in a temperature-controlled room with a 12-h light/dark cycle. Animals had access to standard chow (Harlan Teklad 8604; Madison, WI) and untreated tap water ad libitum. The study was approved by the University of California, San Diego, Animal Subjects Committee and conducted in a pathogen-free facility accredited by the American Association of Accreditation of Laboratory Animal Care. The investigation conforms to the Guide for the Care and Use of Laboratory Animals published by the U.S. National Institutes of Health (NIH Publication No. 85-23, revised 1996).

Tissue Harvest
Tissues were harvested from animals, ages 12 (adult), 24 (old), and 35–36 (very old) months. Because the maximal life span of these rats is 41–42 months (27), the chosen age groups allow for an analysis of the aging process over the last third of the life span, which is the period in humans when the greatest losses in muscle structure and function occur (7). Animals were weighed, anesthetized with 40–70 mg of sodium pentobarbital, and the following muscles were excised as quickly as possible from both sides of the body: adductor longus (Add-Lon), biceps femoris (Bic-Fem), gluteus superficialis (Glu-Sup), plantaris (Plan), extensor digitorum longus (EDL), lateral gastrocnemius (Lat-Gas), medial gastrocnemius (Med-Gas), soleus (Sol), tibialis anterior (Tib-Ant), and diaphragm (Dia). All muscles were harvested within 1.5 hours of administration of anesthetic. Whole muscle samples were weighed immediately, flash-frozen in liquid nitrogen (N2), and stored in a cryovial at –80°C for biochemical and molecular assays. For Lat-Gas, only the deep (fast-twitch oxidative) portion of the muscle was frozen for further analysis. Similarly, only the superficial (fast-twitch glycolytic) portions of the Med-Gas and Tib-Ant were retained.

Enzyme Assays
Frozen muscle samples were powdered using a mortar and pestle under liquid N2 and stored at –80°C until further use. For enzyme assays, powdered tissue was extracted in 20 volumes of a buffer composed of 20 mM HEPES, 1 mM EDTA, 0.1% Triton X-100 (pH 7.2) and homogenized with a glass homogenizer. Specific activities of isocitrate dehydrogenase (IDH), aconitase (ACON), cytochrome c oxidase (COX), and CS were measured on a SpectraMAX Plus spectrophotometer (Molecular Devices, Sunnyvale, CA). IDH, CS, and ACON were assayed according to previously published protocols (1,28). COX was assayed at 550 nm with 50 mM Tris-HCl (pH 8.0) containing 50 µM reduced cytochrome c (1). IDH and ACON activities were measured immediately after homogenization, as activity was found to decline after a 30-minute incubation on ice.

Tissue Protein, DNA, and RNA Content
Protein content (mg/g) of muscle homogenates was measured using a commercial protein assay kit (Bio-Rad, Hercules, CA), based on the Bradford method for protein quantification (29).

DNA content was assayed using DNA-specific fluorescent dyes. Aliquots of muscle homogenate from enzyme assays were reserved and stored at –20°C for assay of mg DNA/g tissue. These measurements are a good estimate of nuclear DNA content because mitochondrial DNA comprises less than 2% of the total DNA content of skeletal muscle (30). Thawed homogenate was diluted 1:6 in DNA digestion buffer and incubated overnight at 55°C. Digestion buffer contained 100 mM NaCl, 10 mM Tris-HCl (pH 8.0), 25 mM EDTA (pH 8.0), 0.5% sodium dodecyl sulfate, and proteinase K at 0.2 mg/ml. Digested homogenate (2 µl) and DNA standards (1.0–60 ng of purified DNA) were loaded onto a 96-well black plate. DNA concentration was quantified according to the manufacturer's instructions with Pico-Green (Molecular Probes, Eugene, OR), a reagent that fluoresces when bound to double-stranded DNA. Fluorescence was detected (excitation 480 nm, emission 520 nm) using a Spectramax Gemini fluorometer (Molecular Devices).

Differences in mg RNA/g tissue of muscles of varied fiber type (17,31,32) must be considered when comparing levels of specific mRNA transcripts across muscles (31). Thus, we measured the total RNA content of all samples to report CS mRNA/g tissue. Frozen powdered tissue was quickly weighed, diluted 10- to 40-fold in GTC solution (4 M guanidium thiocyanate [GTC], 25 mM sodium citrate, 0.5% sarkosyl, 1.0% ß-mercaptoethanol, pH 7.0), and homogenized using a Polytron homogenizer (Brinkmann Instruments, Westbury, NY). Total RNA was extracted from the homogenate using a standard phenol-chloroform procedure according to the methods of Chomczynski and Sacchi (33). After the first phase separation, a small volume (<5%) of the aqueous top layer was reserved for quantification of RNA using the fluorescent reagent Ribo-Green (Molecular Probes). The remainder of the aqueous layer was precipitated and further purified to isolate RNA suitable for northern blot analysis. To quantify mg RNA/g tissue, samples and standards (10–200 ng of purified transfer RNA) were loaded onto a 96-well black plate and measured (excitation 480 nm, emission 536 nm). To account for possible effects of the RNA extraction procedure (i.e., GTC, phenol, chloroform) on fluorescence of Ribo-Green, an equal volume (1 µl) of a mock extraction (i.e., no tissue) was added to each well of the RNA standards.

Northern Blot Analysis
Purified total RNA was denatured and fractionated (5–20 µg/lane) with a standard 1% agarose-formaldehyde gel system. RNA was transferred overnight onto Duralon-UV membrane (Stratagene, La Jolla, CA) and then cross-linked with a UV Crosslinker (Fisher Scientific, Pittsburgh, PA). A complementary DNA (cDNA) probe for COX IV was constructed as previously described (34). CS (GenBank accession no. AF461496) was amplified from rat gastrocnemius cDNA template at 57.5°C using the following primers: 5'-gaaacatcrgttcttgatcc-3' and 5'-gtgtattccagatgtagtcwcgtaa-3'. The resulting 765-bp product was used as a cDNA probe for CS mRNA. Cytochrome c (CYT-C) (GenBank accession no. NM012839) was amplified from rat skeletal muscle cDNA template at 52°C using 5'-cgggacgtctccctaaga-3' and 5'-gctattaggtctgccctttc-3', yielding a 351-bp product that was used as a cDNA probe. Hybridization and phosphorimaging conditions were as described previously (35). Signal intensities for all genes were corrected for loading differences by probing for 18S RNA.

Separate blots were used for every tissue, with each blot containing adult and very old samples. For comparison among tissues, a normalizing blot was generated using samples that appeared on each tissue blot. After probing the normalizing blot with CS, the relative signal strength of CS mRNA across tissues was calculated and used to compare blots.

Statistics
Data are presented as mean ± standard error of the mean (SEM). A two-way analysis of variance (ANOVA) was used to determine significant main effects (p <.05) of age and muscle fiber type on variations in enzyme activity and enzyme stoichiometries. A one-way ANOVA followed by the Tukey-Kramer test was used to detect significant differences (p <.05) within one muscle fiber type across three age groups (adult, old, very old). An independent t test was used to detect differences (p <.05) within one muscle type between the adult and very old age groups. Regression analysis was performed to determine linear relationships between two variables (p <.05). All statistical analyses were performed with JMPIN (Thomson Learning, Florence, KY) statistical software.


   RESULTS
Top
 Abstract
 Methods
 Results
Discussion
 References
 
Regulation of Mitochondrial Content in Different Muscles of Adult Animals
We found that mg DNA/g tissue and CS activity correlate across 10 muscles of varied fiber-type characteristics (Tables 1 and 2) in adult animals, with almost 40% of the variance in mitochondrial content explained by differences in DNA content (Figure 1A, R2 = 0.36, p <.0001). The most parsimonious explanation for this linear relationship is that CS gene expression is regulated primarily by constitutive pathways, and that differences arise from variations in DNA content. If so, then DNA content should also predict CS mRNA levels, which should likewise predict CS enzyme content. CS mRNA/g tissue (Figure 1C) was quantified using paired measurements of CS mRNA/total RNA (via northern blot) and total RNA/g tissue (via Ribo-Green). We found no relationship between CS mRNA/g tissue and mg DNA/g tissue among the muscles (Figure 1C), suggesting that the tissues vary in degree of transcriptional control over CS gene expression. The potential impact of contaminating sources of nonmuscle nuclei in our measurements of mg DNA/g tissue should be considered, but it is expected that these sources also contribute to measurements of mRNA levels and enzyme activity.


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  		Table 1. Changes in Muscle Composition With Age.

 

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  		Table 2. Changes in Specific Activities of Mitochondrial Enzymes With Age.

 

Figure 01
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  		Figure 1. Regulation of mitochondrial content in different muscles of adult and very old animals. Individual paired measurements of citrate synthase (CS) activity and DNA content are compared across 10 skeletal muscles in adult (A) and very old (B) animals. Mean CS messenger RNA (mRNA) content and DNA content of each muscle are compared in adult (C) and very old (D) animals. Mean CS activity and CS mRNA content of each muscle are compared in adult (E) and very old (F) animals. For C–F, each muscle is represented by a unique symbol: adductor longus (black circle), biceps femoris (black triangle), diaphragm (black diamond), extensor digitorum longus (gray triangle), gluteus superficialis (black square), deep lateral gastrocnemius (gray square), superficial medial gastrocnemius (gray circle), plantaris (open triangle), soleus (open circle), and superficial tibialis anterior (open square). N = 4–5 for all measurements. R2 and significance level were determined by linear regression. Standard error of the mean bars that cannot be seen are occluded by the symbol

 
Next, we compared CS activity levels to CS mRNA content across muscles, an index of posttranscriptional variation among tissues. Because CS is not known to be allosterically or covalently regulated, catalytic activity is a good reflection of enzyme content (36). If CS enzyme levels were determined primarily by transcriptional regulation, we would predict a linear relationship between CS mRNA/g tissue and CS activity/g tissue. The lack of relationship between these parameters (Figure 1E) suggests an influence of posttranscriptional controls on CS content in different muscles.

As a result of this analysis, it is clear that some tissues have more CS protein than would be expected based on CS mRNA, and other tissues have more CS mRNA than would be expected based on DNA content. For example, deep Lat-Gas maintains one of the highest levels of CS activity/g tissue with one of the lowest quantities of CS mRNA/g tissue (Figure 1E). Thus, the linear relationship between units CS/g tissue and mg DNA/g tissue is not a simple relationship between constitutive gene expression and nuclear DNA content.

Changes in Muscle Composition With Age
To investigate how the regulation of mitochondrial content is altered with age, we evaluated the same set of muscles in adult (12 months), old (24 months), and very old (35–36 months) rats. In Table 1, aging-related compositional changes are reported for each muscle. With the exception of Add-Lon, considerable atrophy occurred in all tissues, as indicated by an 18%–51% loss in absolute muscle mass. As body mass changed very little with age (~10% higher in old animals, see legend of Table 1), similar declines (17%–49%) in muscle-specific mass were observed. Although there was a trend for decreased protein (mg/g tissue), no significant declines were observed with age except in the superficial Tib-Ant (18%), Bic-Fem (21%), and Dia (13%). DNA levels/g muscle significantly increased in all of the fast-twitch muscles of very old animals (by 33%–75%); no differences were observed in the old age group relative to adults. RNA content (mg/g tissue) also significantly increased in many of the fast-twitch muscles (up to 2.3-fold), except in the superficial Med-Gas and superficial Tib-Ant. No significant changes in DNA or RNA occurred in slow-twitch and mixed muscles, with the exception of a 49% increase in DNA in the deep Lat-Gas.

Regulation of Mitochondrial Content in Different Muscles of Aged Animals
The linear relationship between DNA and CS activity across muscles in adult animals (R2 = 0.36) is preserved in the old age group (R2 = 0.48, p <.001, data not shown). Conversely, in very old animals, only 10% of the variation in CS activity across muscles is attributed to differences in DNA content (Figure 1B, R2 = 0.10, p =.03). Because of the similarity between adult and old animals, the effects of aging on CS mRNA expression were evaluated only in the very old age group. As with the adults, there was no simple relationship between either DNA content and CS mRNA levels (Figure 1D) or CS mRNA levels and CS activity (Figure 1F).

In analyzing specific changes within each muscle with age, we found that units CS/g tissue varied little (Table 2), but units CS/DNA significantly declined (by 19%–50%) in almost all tissues (Figure 2A). This decline signifies major changes in the regulation of nucleus-encoded mitochondrial gene expression. In some tissues that we classified as Group I (superficial Tib-Ant, superficial Med-Gas, deep Lat-Gas, Dia), declines in units CS/DNA are paralleled by declines in CS mRNA/DNA (Figure 2, A and B). These changes suggest that declines in transcription may occur in these muscles. In other tissues that we classified as Group II (Plan, Sol, Bic-Fem, Glu-Sup), decreases in units CS/DNA are paralleled by declines in units CS/CS mRNA (Figure 2, A and C), indicating reductions in posttranscriptional activity. Finally, in two muscles that we classified as Group III, declines in both CS mRNA/DNA and units CS/CS mRNA accompany the decreases in units CS/DNA (Figure 2, A–C). Overall, our results reveal muscle-specific mechanisms for maintaining mitochondrial enzyme levels in aging tissue, with changes occurring at the transcriptional and/or posttranscriptional levels.


Figure 02
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  		Figure 2. Muscle-specific changes in the regulation of mitochondrial content with age. Measurements of citrate synthase (CS) activity, DNA content, and CS messenger RNA (mRNA) content in each tissue are expressed as (A) units CS/DNA (B) CS mRNA/DNA, and (C) units CS/CS mRNA. All values are mean ± standard error of the mean, expressed relative to adult within a tissue (open bars, adult; solid bars, very old). Muscles are grouped according to their pattern of change with age (Groups I–III). N = 5 unless otherwise indicated in parentheses. *Indicates significantly different than adult (p <.05). Add-Lon = adductor longus; Bic-Fem = biceps femoris; Dia = diaphragm; EDL = extensor digitorum longus; Glu-Sup = gluteus superficialis; Lat-Gas = lateral gastrocnemius; Med-Gas = medial gastrocnemius; Plan = plantaris; Sol = soleus; Tib-Ant = tibialis anterior

 
Changes in Transcript Levels of Other Nucleus-Encoded Mitochondrial Genes
To further investigate muscle-specific changes in transcriptional regulation of bioenergetic gene expression, we also evaluated transcript levels for CYT-C and COX IV, two other nucleus-encoded mitochondrial genes. Consistent with Figure 2B, Group I tissues with decreased CS mRNA/DNA showed lower levels of CS mRNA relative to total RNA (Figure 3A), providing further evidence for a down-regulation of CS transcriptional activity in these muscles. However, there were no declines in transcripts for CYT-C or COX IV (Figure 3, B and C); in fact, a trend for increased expression is observed in the deep Lat- and superficial Med-Gas. Group II tissues, which showed no reduction in CS mRNA/DNA with age (Figure 2B), likewise exhibited no decrease in CS mRNA relative to total RNA (Figure 3A). Moreover, Sol and Glu-Sup displayed significant or modest increases in CS, CYT-C, and COX IV transcripts with age (Figure 3, A–C). Two exceptional tissues (EDL, Plan) demonstrated significant declines in CYT-C expression (by 19%–34%, respectively), with no changes in other transcripts. The lack of a consistent trend in mRNA levels for all three nuclear genes indicates that complex regulation by transcriptional activators/suppressors underlies differential expression profiles within and across different muscles.


Figure 03
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  		Figure 3. Muscle-specific changes in nucleus-encoded mitochondrial transcripts with age. Expression of citrate synthase (CS) (A), cytochrome c (CYT-C) (B), and cytochrome c oxidase IV (COX IV) (C) messenger RNA (mRNA) were evaluated by northern blot analyses. All values are the mean hybridization signal for each gene, corrected for loading by 18S and expressed relative to adult within a tissue (open bars, adult; solid bars, very old). Muscles are grouped as in
Figure 2. Add-Lon = adductor longus; Bic-Fem = biceps femoris; Dia = diaphragm; EDL = extensor digitorum longus; Glu-Sup = gluteus superficialis; Lat-Gas = lateral gastrocnemius; Med-Gas = medial gastrocnemius; Plan = plantaris; Sol = soleus; Tib-Ant = tibialis anterior. N = 5 unless otherwise indicated in parentheses. *Indicates significantly different than adult (p <.05). D, Representative northern blot (Glu-Sup tissue)

 
Maintenance of Mitochondrial Enzyme Stoichiometries
Although aging caused significant declines in enzyme activities relative to DNA content, regulatory changes in enzyme synthesis seemed to match reductions in muscle size. Thus, specific activity levels (per g tissue) of CS, IDH, ACON, and COX remained constant with age in most tissues. Of these enzymes, all are encoded entirely by nuclear DNA, except for COX, which is partially encoded by mtDNA. A two-way ANOVA was used to determine main effects of age on enzyme activity. No significant effect of age was found on IDH (F = 1.54, p =.2184) or ACON (F = 2.62, p =.077) activity. A significant main effect of age was found on CS (F = 4.34, p =.015) and COX (F = 5.44, p =.006) activity. However, a Tukey post hoc analysis of Age x Muscle did not reveal any significant differences across age groups for specific muscles. To more closely evaluate the effects of aging on enzyme activity within each muscle, a one-way ANOVA was used (see Table 2). This analysis revealed that a few tissues displayed significant changes in some enzymes with age. Significant losses (20%–34%) occurred in all enzymes in the deep Lat-Gas of very old animals. In Plan, COX and IDH were 20%–30% lower, and although not significant, CS and ACON levels appeared modestly depressed. Although Sol and Add-Lon showed a trend for lower activities of most enzymes with age, the changes were not statistically significant. Interestingly, ACON levels were significantly higher (by 50%) in the Glu-Sup and EDL (by 25%). Although the differences were not statistically significant, other enzymes appeared to be elevated in the Glu-Sup, EDL, and superficial Tib-Ant.

With respect to enzyme stoichiometries, ratios of IDH/CS, ACON/CS, and COX/CS did not significantly change with age in most muscles. We found no main effect of age on IDH/CS (F = 1.95, p =.147) or ACON/CS (F = 0.185, p =.8318). There was a significant main effect of age on COX/CS (F = 5.02, p =.008); however, a Tukey post hoc analysis of Age x Muscle revealed no significant differences among age groups for specific muscles. To more closely evaluate the effects of aging on enzyme ratios within each muscle, a one-way ANOVA was performed. This analysis revealed no significant differences in enzyme ratios among age groups in any muscle, with the exception of a 17% decline in COX/CS in EDL (very old vs adult; Figure 4, A–C).


Figure 04
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  		Figure 4. Mitochondrial enzyme stoichiometries do not change with age. Paired measurements of cytochrome c oxidase (COX), isocitrate dehydrogenase (IDH), aconitase (ACON), and citrate synthase (CS) activities were analyzed in homogenates of 10 muscles from adult (open bars), old (gray bars), and very old (black bars) animals. Enzyme stoichiometries are expressed as ratios of IDH/CS (A), ACON/CS (B), and COX/CS (C). Muscles grouped under the same horizontal line are not significantly different from each other (two-way analysis of variance (ANOVA), main effect of muscle type, p <.05). Add-Lon = adductor longus; Bic-Fem = biceps femoris; Dia = diaphragm; EDL = extensor digitorum longus; Glu-Sup = gluteus superficialis; Lat-Gas = lateral gastrocnemius; Med-Gas = medial gastrocnemius; Plan = plantaris; Sol = soleus; Tib-Ant = tibialis anterior. *Indicates significantly different than adult (one-way ANOVA, p <.05). N = 5 for all enzymes, except IDH in Sol (N = 3)

 
Though enzyme stoichiometries were not largely affected by aging, there was a significant main effect of muscle type on IDH/CS (F = 33.5, p <.0001), ACON/CS (F = 9.25, p <.0001), and COX/CS (F = 12.87, p <.0001). Despite this variation among some muscles (Figure 4), it is important to note that COX/CS and ACON/CS ratios are similar among the majority of the muscles (IDH/CS is more variable).


   DISCUSSION
Top
 Abstract
 Methods
 Results
 Discussion
References
 
Many aspects of the muscle phenotype change in response to development and aging, including cellular morphology (e.g., myonuclear domain), contractile protein profiles (e.g., myosin isoforms), and the bioenergetic machinery (e.g., mitochondrial content). Although these aspects of the remodeling process have been studied in isolation, the extent to which they are co-regulated remains largely unknown. There is a general relationship between these aspects of fiber type, but it is not clear if this relationship is due to shared regulatory pathways or parallel but independent pathways. In discussing these distinct features that contribute to a fiber-type phenotype, it is important to recognize that a change in one parameter (e.g., myosin profile) does not cause a change in another parameter (e.g., myonuclear domain), although they may change in parallel.

Determining the mechanistic basis of differences in mitochondrial content between tissues and physiological states is difficult because of complex controls on transcriptional, translational, and posttranslational processes. For example, differences in DNA content and general turnover rates of RNA, protein, and organelles have major regulatory influences on mitochondrial content. However, recent studies have implicated PGC-1{alpha}, acting through a network of transcription factors, as a master controller of mitochondrial biogenesis. These transcriptional regulators (along with other general transcriptional machinery) could account for patterns of mitochondrial gene expression among muscles and during aging by two separate regulatory mechanisms. First, constitutive expression of nucleus-encoded mitochondrial genes in combination with differences in nuclear DNA content might explain a significant fraction of the variation in mitochondrial content between muscles or physiological states. Thus, a slow oxidative muscle might have more CS enzyme/g tissue simply because it has more nuclear DNA/g tissue (12). Second, superimposed on this constitutive background are tissue-specific variations in transcriptional regulation—that is, myonuclei in oxidative muscles may express more mitochondrial enzyme because of a relatively high activity of transcriptional activators, such as PGC-1{alpha}, on mitochondrial genes. Both of these mechanisms can affect the relationship between DNA, mRNA, and enzyme levels within a tissue.

Impact of Myonuclear Content on CS Activity Across Fiber Type
The initial data of this study, which revealed a significant correlation (R2 = 0.36) between units CS/g tissue and mg DNA/g tissue in adult tissues (Figure 1A), suggested that constitutive expression in combination with differences in DNA content could be a primary explanation for variation in CS activity among muscles of different fiber type. On the basis of this observation, we hypothesized that each myonucleus would produce the same amount of transcript, which in turn would generate the same amount of protein. However, we found that neither CS mRNA/DNA stoichiometries nor units CS/CS mRNA stoichiometries were preserved across tissues (Figure 1, C and E). We conclude that no simple regulatory pattern or process is responsible for the tissue-specific differences in CS enzyme levels. The relative importance of transcriptional and posttranscriptional processes in regulating these changes is specific for each muscle.

Although CS is traditionally considered to be transcriptionally regulated (26), our data in conjunction with other reports (37,38) suggests that differences in CS content among muscles arise through other mechanisms. Consider the patterns in Sol versus deep Lat-Gas of adult animals. Compared to deep Lat-Gas, Sol has 6 times more CS mRNA/DNA (0.436 vs 0.08, Figure 1C), 40% fewer units CS/g tissue (14.1 vs 24.2, Table 2), and 92% fewer units CS/CS mRNA (38 vs 439, Figure 1E). This comparison offers several insights into the regulation of CS enzyme synthesis. First, the difference in CS mRNA/DNA ratios indicates that Sol may have higher rates of transcription for CS gene expression than does deep Lat-Gas, possibly due to greater activity of transcriptional regulators for mitochondrial gene expression. Second, variation in CS enzyme content between the tissues cannot be easily attributed to transcriptional mechanisms, as CS mRNA levels do not predict CS enzyme levels. Finally, different ratios of units CS/CS mRNA between the muscles suggest that deep Lat-Gas has higher rates of posttranscriptional activity for CS enzyme synthesis compared to Sol. Although the mechanism remains to be elucidated, our data provide evidence that posttranscriptional pathways are involved in the regulation of CS enzyme content in different muscles.

Influence of Aging on Myonuclear Content
In our study, muscles of adult and old animals were similar in most respects. With respect to nuclear content, changes in mg DNA/g were observed only in the very old (35–36 months) age group. The same age-dependent pattern is seen in human studies (18–21), where increases in DNA are not seen until after the age of 60. Increases in mg DNA/g tissue with age occurred preferentially in fast-twitch muscles, as is seen in human aging (18–21). It is likely that the increase in DNA can be attributed to a reduction in fiber size, which may arise two ways. First, the changes in muscle morphology may arise from atrophy, where an increase in nuclear content per gram of muscle coincides with a loss of contractile machinery. Second, an increase in nuclear content may also occur when a muscle actively converts a fiber from Type II to Type I, a transition that is known to occur in some models of aging (6,24). The magnitude of the change in myonuclear content associated with a change in fiber type depends on the relative proportion of Type I versus Type II fibers. Because Type I fibers typically have the highest nuclear content (12), the impact of fiber-type switching would be greatest when a predominantly Type II muscle becomes a predominantly Type I muscle. In these animals, it has been shown that aging does not affect the proportion of Type I fibers in Sol, EDL, or Lat-Gas (39); therefore, changes we observed in nuclear content in these muscles are not associated with a change in fiber type. The proportion of Type I fibers does increase in Plan, Med-Gas, and Tib-Ant (39); however, the Type I fibers comprise less than 20% of the total in each of these muscles, even in very old animals. Thus, an increase in proportion of Type I fibers in these muscles (which are predominantly Type II) may contribute to, but does not account for, the observed increase in muscle nuclear content.

Overall, our data add to a growing body of evidence that aging results in a higher DNA content of muscle tissue, which is expected to have profound effects on gene expression. Interestingly, mg RNA/g tissue paralleled mg DNA/g tissue in most muscles (Add-Lon, EDL, Glu-Sup, Plan, superficial Tib-Ant, and Bic-Fem). However, in other muscles, mg RNA/g tissue did not change despite increases in mg DNA/g tissue; this finding may reflect changes in global transcription or RNA stability with age.

Regulation of Enzyme Content in Aging Muscles
It is important to consider how changes in fiber type might influence the patterns in mitochondrial gene expression. As we discussed previously in the context of myonuclear content, remodeling associated with fiber-type shifts could potentially contribute to the metabolic profile of muscles with age. As discussed previously, three of the muscles in our aging model are known to experience an increase in the proportion of Type I fibers: Plan, Med-Gas, and Tib-Ant (39). As Type II fibers are remodeled to Type I fibers, it may be predicted that the mitochondrial content would increase. However, there was no detectible change in mitochondrial content (i.e., units CS/g) in these muscles as a result of aging. As with myonuclei, the fiber-type conversions do not appear to play a major role in determining mitochondrial content.

Although units CS/g tissue remained relatively stable, significant declines in units CS/DNA occurred in almost all muscles (Figure 2A), which signifies major changes in the regulation of CS enzyme synthesis. In Group I muscles (superficial Tib-Ant, superficial Med-Gas, deep Lat-Gas, Dia), these declines may be linked to changes in transcriptional activity, as CS mRNA/DNA ratios declined in older animals (Figure 2B). In the deep Lat-Gas, relatively low levels of CS mRNA/DNA (Figure 1C) in adult animals may predispose this tissue to declines in transcriptional control of CS gene expression with age. The trend of lower CS mRNA/DNA in Group I muscles is paralleled by a decline in specific CS mRNA expression relative to total RNA (Figure 3A). With respect to other nucleus-encoded mitochondrial genes, CYT-C mRNA/total RNA and COX IV mRNA/total RNA did not significantly change (Figure 3, B and C). This finding suggests that gene-specific regulatory mechanisms are differentially affected by aging. In Group II tissues (Plan, Sol, Bic-Fem, Glu-Sup), declines in units CS/DNA may be linked to decreases in posttranscriptional activity, as indicated by decreases in units CS/CS mRNA (Figure 2C). It is unknown whether the declines in units CS/DNA that we observed in Group II muscles with age are linked to changes in elongation, or other points of posttranscriptional control. It is interesting to note that the basis of assignment of muscles to Groups I, II, and III (Figures 2 and 3) does not appear to be linked to the fiber-type profile, as each group is composed of muscles of different fiber type.

Despite much research over the last few decades, it remains equivocal as to whether specific activity levels of mitochondrial respiratory and/or matrix enzymes are affected during aging (40–42). Discrepancies among reports are often due to differences in experimental design including muscle type, species, strain, age group comparisons, and tissue preparation (tissue homogenates vs isolated mitochondria). Several groups have shown that changes in enzyme activity are dependent on muscle fiber type (6,43,44). Our study is one of the few to evaluate multiple mitochondrial enzymes across a broad spectrum of skeletal muscles in the context of aging. We found a significant main effect of age on CS and COX activity with all muscle groups pooled together. However, analysis of individual muscle comparisons across age groups revealed that most enzyme activities were unaltered with age, with a few exceptions. Most notably, the deep Lat-Gas exhibited significant losses (20%–30%) in CS, ACON, IDH, and COX activities/g tissue. A decline in mitochondrial enzyme activity specifically in the deep Lat-Gas (compared to other muscles) has been previously reported (43); however, the mechanism behind this muscle-specific pattern is unknown. Other tissues that showed significant changes in enzyme activity were the Glu-Sup and EDL (each exhibiting a 50% and 25% increase in ACON, respectively) and Plan (34% decline in IDH and 20% decline in COX). In a study of similar design to ours, Hagen and coworkers (45) found that CS activity declined only in very old (36 month) animals. Although this group did not measure COX activity (which decreased with age in our study), they found that flux through Complexes I–III declined with age.

While absolute changes in CS activity are an estimate of variations in mitochondrial content, changes in the stoichiometric relationships between different mitochondrial enzymes may reflect mitochondrial dysfunction. We found that ratios of IDH/CS, ACON/CS, and COX/CS did not significantly change with age in any muscle, except in the EDL (17% decline in COX/CS, very old vs adult). Without further analysis (e.g., mitochondrial adenosine triphosphate production), it is difficult to know whether this modest decline in COX/CS in EDL actually reflects mitochondrial dysfunction. Although there are a few other reports of unchanged enzyme stoichiometries during aging (46–48) and other experimental interventions (49,50), our results contrast with those of several studies that show significant changes in mitochondrial enzyme stoichiometries during aging (23–25,45,51–53). In particular, the study of Hagen and coworkers (described previously) reports a greater decline in respiratory chain enzyme activity relative to CS activity in Plan of aged F344xBNF1 rats (45). This group also showed that these declines in respiratory chain activity were accompanied by increased levels of mtDNA deletions (45), a finding that has been reported by several groups (54–56).

We have evaluated the relative importance of DNA content, transcriptional, and posttranscriptional processes on the regulation of mitochondrial content in muscles of different fiber-type composition and during aging. Although some variability in CS enzyme activity across muscles can be attributed to differences in mg DNA/g tissue, individual muscles demonstrated unique patterns of regulatory controls at the transcriptional and posttranscriptional levels. Although the stoichiometric relationships between mitochondrial enzymes were preserved among the majority of muscle groups, some variability in enzyme ratios between different muscles may reflect these unique patterns of gene regulatory controls. With aging, we found marked declines in CS enzyme activity relative to DNA content. Changes in the relative importance of nuclear content, transcriptional and posttranscriptional regulation for determining CS enzyme levels were muscle-specific. The role of these mechanisms in regulating mitochondrial biogenesis in different muscles must be considered in future studies that focus on the causes and consequences of energetic deficiencies in aging muscle.


   Acknowledgments
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Funding for this research was provided by grants from the Natural Sciences and Engineering Research Council of Canada (to C.D.M.), the American Federation of Aging Research (to C.N.L.), and the National Institutes of Health (to O.M.C.).

We thank Dr. Li Cui, Dr. Yan Ju, and Margarita Trejo-Morales for assistance with tissue harvesting.


   Footnotes
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Decision Editor: James R. Smith, PhD

Received March 4, 2005

Accepted July 1, 2005


   References
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  1. Moyes CD, Mathieu-Costello OA, Tsuchiya N, Filburn C, Hansford RG. Mitochondrial biogenesis during cellular differentiation. Am J Physiol. 1997;272:C1345-C1351.
  2. Delp MD, Duan C. Composition and size of type I, IIA, IID/X, and IIB fibers and citrate synthase activity of rat muscle. J Appl Physiol. 1996;80:261-270.[Abstract/Free Full Text]
  3. Nicholls D, Cunningham S, Wiesinger H. Mechanisms of thermogenesis in brown adipose tissue. Biochem Soc Trans. 1986;14:223-225.[Medline]
  4. Hood DA. Contractile activity-induced mitochondrial biogenesis in skeletal muscle. J Appl Physiol. 2001;90:1137-1157.[Abstract/Free Full Text]
  5. Nelson BD. Thyroid hormone regulation of mitochondrial function. Comments on the mechanism of signal transduction. Biochim Biophys Acta. 1990;1018:275-277.[Medline]
  6. Holloszy JO, Chen M, Cartee GD, Young JC. Skeletal muscle atrophy in old rats: differential changes in the three fiber types. Mech Ageing Dev. 1991;60:199-213.[Medline]
  7. Greenlund LJ, Nair KS. Sarcopenia—consequences, mechanisms, and potential therapies. Mech Ageing Dev. 2003;124:287-299.[Medline]
  8. Puigserver P, Spiegelman BM. Peroxisome proliferator-activated receptor-gamma coactivator 1 alpha (PGC-1 alpha): transcriptional coactivator and metabolic regulator. Endocr Rev. 2003;24:78-90.[Abstract/Free Full Text]
  9. Puigserver P, Wu Z, Park CW, Graves R, Wright M, Spiegelman BM. A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell. 1998;92:829-839.[Medline]
  10. Wu Z, Puigserver P, Andersson U, et al. Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell. 1999;98:115-124.[Medline]
  11. Lin J, Wu H, Tarr PT, et al. Transcriptional co-activator PGC-1 alpha drives the formation of slow-twitch muscle fibres. Nature. 2002;418:797-801.[Medline]
  12. Tseng BS, Kasper CE, Edgerton VR. Cytoplasm-to-myonucleus ratios and succinate dehydrogenase activities in adult rat slow and fast muscle fibers. Cell Tissue Res. 1994;275:39-49.[Medline]
  13. Atherton GW, James NT. Stereological analysis of the number of nuclei in skeletal muscle fibres. Acta Anat (Basel). 1980;107:236-240.[Medline]
  14. Burleigh IG. Observations on the number of nuclei within the fibres of some red and white muscles. J Cell Sci. 1977;23:269-284.[Abstract/Free Full Text]
  15. Matthew CA, Moore MJ. Numbers of myonuclei and satellite cell nuclei in latissimus dorsi muscles of the chicken. Cell Tissue Res. 1987;248:235-238.[Medline]
  16. Allen DL, Monke SR, Talmadge RJ, Roy RR, Edgerton VR. Plasticity of myonuclear number in hypertrophied and atrophied mammalian skeletal muscle fibers. J Appl Physiol. 1995;78:1969-1976.[Abstract/Free Full Text]
  17. Babij P, Booth FW. Alpha-actin and cytochrome c mRNAs in atrophied adult rat skeletal muscle. Am J Physiol. 1988;254:C651-C656.
  18. Manta P, Vassilopoulos D, Spengos M. Nucleo-cytoplasmic ratio in ageing skeletal muscle. Eur Arch Psychiatry Neurol Sci. 1987;236:235-236.[Medline]
  19. Forsberg AM, Nilsson E, Werneman J, Bergstrom J, Hultman E. Muscle composition in relation to age and sex. Clin Sci (Lond). 1991;81:249-256.[Medline]
  20. Kadi F, Charifi N, Denis C, Lexell J. Satellite cells and myonuclei in young and elderly women and men. Muscle Nerve. 2004;29:120-127.[Medline]
  21. Renault V, Thornell LE, Eriksson PO, Butler-Browne G, Mouly V, Thorne LE. Regenerative potential of human skeletal muscle during aging. Aging Cell. 2002;1:132-139.[Medline]
  22. Allen DL, Roy RR, Edgerton VR. Myonuclear domains in muscle adaptation and disease. Muscle Nerve. 1999;22:1350-1360.[Medline]
  23. Aspnes LE, Lee CM, Weindruch R, Chung SS, Roecker EB, Aiken JM. Caloric restriction reduces fiber loss and mitochondrial abnormalities in aged rat muscle. FASEB J. 1997;11:573-581.[Abstract]
  24. Bua EA, McKiernan SH, Wanagat J, McKenzie D, Aiken JM. Mitochondrial abnormalities are more frequent in muscles undergoing sarcopenia. J Appl Physiol. 2002;92:2617-2624.[Abstract/Free Full Text]
  25. Desai VG, Weindruch R, Hart RW, Feuers RJ. Influences of age and dietary restriction on gastrocnemius electron transport system activities in mice. Arch Biochem Biophys. 1996;333:145-151.[Medline]
  26. Annex BH, Kraus WE, Dohm GL, Williams RS. Mitochondrial biogenesis in striated muscles: rapid induction of citrate synthase mRNA by nerve stimulation. Am J Physiol. 1991;260:C266-C270.
  27. Turturro A, Witt WW, Lewis S, Hass BS, Lipman RD, Hart RW. Growth curves and survival characteristics of the animals used in the Biomarkers of Aging Program. J Gerontol A Biol Sci Med Sci. 1999;54:B492-B501.[Abstract]
  28. Gardner P, Nguyen D, White C. Aconitase is a sensitive and critical target of oxygen poisoning in cultured mammalian cells and in rat lungs. Proc Natl Acad Sci U S A. 1994;91:12248-12252.[Abstract/Free Full Text]
  29. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem. 1976;72:248-254.[Medline]
  30. Miller F, Rosenfeldt F, Zhang C, Linnane A, Nagley P. Precise determination of mitochondrial DNA copy number in human skeletal and cardiac muscle by a PCR-based assay: lack of change of copy number with age. Nucleic Acids Res. 2003;31:e61.[Abstract/Free Full Text]
  31. Habets PE, Franco D, Ruijter JM, Sargeant AJ, Pereira JA, Moorman AF. RNA content differs in slow and fast muscle fibers: implications for interpretation of changes in muscle gene expression. J Histochem Cytochem. 1999;47:995-1004.[Abstract/Free Full Text]
  32. Leary SC, Battersby BJ, Moyes CD. Inter-tissue differences in mitochondrial enzyme activity, RNA and DNA in rainbow trout (Oncorhynchus mykiss). J Exp Biol. 1998;201:(Pt 24): 3377-3384.[Abstract]
  33. Chomczynski P, Sacchi N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem. 1987;162:156-159.[Medline]
  34. Leary SC, Battersby BJ, Hansford RG, Moyes CD. Interactions between bioenergetics and mitochondrial biogenesis. Biochim Biophys Acta. 1998;1365:522-530.[Medline]
  35. Leary SC, Hill BC, Lyons CN, et al. Chronic treatment with azide in situ leads to an irreversible loss of cytochrome c oxidase activity via holoenzyme dissociation. J Biol Chem. 2002;277:11321-11328.[Abstract/Free Full Text]
  36. Remington SJ. Structure and mechanism of citrate synthase. Curr Top Cell Regul. 1992;33:209-229.[Medline]
  37. Leary SC, Lyons CN, Rosenberger AG, Ballantyne JS, Stillman J, Moyes CD. Fiber-type differences in muscle mitochondrial profiles. Am J Physiol Regul Integr Comp Physiol. 2003;285:R817-R826.[Abstract/Free Full Text]
  38. Lucassen M, Schmidt A, Eckerle LG, Portner HO. Mitochondrial proliferation in the permanent vs. temporary cold: enzyme activities and mRNA levels in Antarctic and temperate zoarcid fish. Am J Physiol Regul Integr Comp Physiol. 2003;285:R1410-R1420.[Abstract/Free Full Text]
  39. Cui L, Ju Y, Trejo-Morales M, Costello ML, Mathieu-Costello OA. Arteriolar and venular capillary distribution in skeletal muscle of old rats. FASEB J. 2003;17:A1278.
  40. Lenaz G. Role of mitochondria in oxidative stress and ageing. Biochim Biophys Acta. 1998;1366:53-67.[Medline]
  41. Lenaz G, D'Aurelio M, Merlo PM, et al. Mitochondrial bioenergetics in aging. Biochim Biophys Acta. 2000;1459:397-404.[Medline]
  42. Rasmussen UF, Krustrup P, Kjaer M, Rasmussen HN. Experimental evidence against the mitochondrial theory of aging. A study of isolated human skeletal muscle mitochondria. Exp Gerontol. 2003;38:877-886.[Medline]
  43. Barazzoni R, Short KR, Nair KS. Effects of aging on mitochondrial DNA copy number and cytochrome c oxidase gene expression in rat skeletal muscle, liver, and heart. J Biol Chem. 2000;275:3343-3347.[Abstract/Free Full Text]
  44. Houmard JA, Weidner ML, Gavigan KE, Tyndall GL, Hickey MS, Alshami A. Fiber type and citrate synthase activity in the human gastrocnemius and vastus lateralis with aging. J Appl Physiol. 1998;85:1337-1341.[Abstract/Free Full Text]
  45. Hagen JL, Krause DJ, Baker DJ, Fu MH, Tarnopolsky MA, Hepple RT. 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. 2004;59:1099-1110.[Abstract/Free Full Text]
  46. Hansford RG, Castro F. Age-linked changes in the activity of enzymes of the tricarboxylate cycle and lipid oxidation, and of carnitine content, in muscles of the rat. Mech Ageing Dev. 1982;19:191-200.[Medline]
  47. Rooyackers OE, Adey DB, Ades PA, Nair KS. Effect of age on in vivo rates of mitochondrial protein synthesis in human skeletal muscle. Proc Natl Acad Sci U S A. 1996;93:15364-15369.[Abstract/Free Full Text]
  48. Skorjanc D, Traub I, Pette D. Identical responses of fast muscle to sustained activity by low-frequency stimulation in young and aging rats. J Appl Physiol. 1998;85:437-441.[Abstract/Free Full Text]
  49. Reichmann H, Wasl R, Simoneau JA, Pette D. Enzyme activities of fatty acid oxidation and the respiratory chain in chronically stimulated fast-twitch muscle of the rabbit. Pflugers Arch. 1991;418:572-574.[Medline]
  50. Reichmann H, Hoppeler H, Mathieu-Costello OA, von Bergen F, Pette D. Biochemical and ultrastructural changes of skeletal muscle mitochondria after chronic stimulation in rabbits. Pflugers Arch. 1985;404:1-9.[Medline]
  51. Kwong LK, Sohal RS. Age-related changes in activities of mitochondrial electron transport complexes in various tissues of the mouse. Arch Biochem Biophys. 2000;373:16-22.[Medline]
  52. Trounce I, Byrne E, Marzuki S. Decline in skeletal muscle mitochondrial respiratory chain function: possible factor in ageing. Lancet. 1989;1:637-639.[Medline]
  53. Tonkonogi M, Fernstrom M, Walsh B, et al. Reduced oxidative power but unchanged antioxidative capacity in skeletal muscle from aged humans. Pflugers Arch. 2003;446:261-269.[Medline]
  54. Fayet G, Jansson M, Sternberg D, et al. Ageing muscle: clonal expansions of mitochondrial DNA point mutations and deletions cause focal impairment of mitochondrial function. Neuromuscul Disord. 2002;12:484-493.[Medline]
  55. Drew B, Leeuwenburgh C. Ageing and subcellular distribution of mitochondria: role of mitochondrial DNA deletions and energy production. Acta Physiol Scand. 2004;182:333-341.[Medline]
  56. DiMauro S, Tanji K, Bonilla E, Pallotti F, Schon EA. Mitochondrial abnormalities in muscle and other aging cells: classification, causes, and effects. Muscle Nerve. 2002;26:597-607.[Medline]



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