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The Journals of Gerontology Series A: Biological Sciences and Medical Sciences 57:B366-B374 (2002)
© 2002 The Gerontological Society of America

Effects of Long-Term Denervation on Skeletal Muscle in Old Rats

Bruce M. Carlsona,b, Andrei B. Borisovb, Eduard I. Dedkovb, Abdelnaby Khalyfad, Tatiana Y. Kostrominovaa,b, Peter C. D. Macphersonc, Eugenia Wangd and John A. Faulknera

a Institute of Gerontology, University of Michigan, Ann Arbor
b Department of Cell and Developmental Biology, University of Michigan, Ann Arbor
c Mental Health Research Institute, University of Michigan, Ann Arbor
d Department of Biochemistry and Molecular Biology, University of Louisville, Kentucky

Bruce M. Carlson, Institute of Gerontology, University of Michigan, 300 North Ingalls Building, Room 913, Ann Arbor, MI 48109-2007 E-mail: brcarl{at}umich.edu.

Decision Editor: James R. Smith, PhD


   Abstract
Top
 Abstract
Methods
 Results
 Discussion
 References
 
We compared the reactions to denervation of limb muscles between young adult and old rats. After denervation for up to 4 months in 24-month-old rats, limb muscles were removed and analyzed for contractile properties, morphology, and levels of several key molecules, including the peptide elongation factors eEF1A-1 and eEF1A-2/S1, myogenin, {gamma}-subunit of the acetylcholine receptor, and cyclin D3. The principal difference between denervated old and young muscle is a somewhat slower rate of atrophy in denervated older muscle, especially among the type II fibers. Expression levels of certain molecules were higher in old than in young control muscle, but after denervation, levels of these molecules increased to the same absolute values in both young and old rats. Although many aspects of postdenervation reactions do not differ greatly between young and old animals, the lesser degree of atrophy in the old rats may reflect significant age-based mechanisms.

RESEARCH over recent decades has clearly established that changes in the pattern of innervation are a characteristic part of the landscape in the aging of skeletal muscle (1). A typical scenario over time would include the death of alpha motoneurons innervating fast muscle fibers (2)(3)(4). This results in either the temporary or permanent denervation of muscle fibers supplied by a given motoneuron. Considerable evidence suggests that a significant number of the denervated fast muscle fibers are ultimately reinnervated by sprouts extending from other neurons, particularly those supplying slow (type I) muscle fibers (3)(4). Other muscle fibers may remain denervated and undergo denervation atrophy. How long muscle fibers that have been supplied by dying motor axons remain denervated before becoming reinnervated by other axonal sprouts is unknown. The ultimate consequence of this motor unit remodeling in old age is a reduction in the proportion of fast muscle fibers and a corresponding increase in the proportion of slow muscle fibers in a given muscle (1)(3)(5).

Motor unit remodeling in old muscle appears to occur over an extended period of time, rather than as a burst phenomenon. Thus, it is difficult to study the reactions of the muscle fibers that are denervated under these circumstances. Therefore, we have used an entire muscle denervation model to investigate the reactions of skeletal muscle in old rats to prolonged denervation. To date, most published studies have been conducted on long-term denervated muscle in young adult rats (6)(7)(8)(9)(10)(11)(12)(13). The principal question has been the extent to which long-term denervated muscle is capable of restoration through the reversal of atrophy upon reinnervation or by regeneration.

Because motor unit remodeling regularly occurs only in old rats, it is important to determine whether in old rats the reactions of muscle fibers to denervation differ from those occurring in denervated muscle fibers of young adult (4-month-old) rats. The experiments reported here are based on denervation experiments conducted on 24-month-old rats. The experiments were designed to determine whether or not there are significant differences in the reactions to denervation of limb muscles between young adult and old rats.


   Methods
Top
 Abstract
 Methods
Results
 Discussion
 References
 
Animals and Surgical Methods
The experimental portion of this study was conducted on fifteen 24-month-old male rats of the WI/Hicks Car strain. In addition, tissues of normal rats of the same strain from embryonic Day 18 to 34 months of age (at least three muscles per time point) were used for certain control studies. The rats were aged under pathogen-free conditions at Harlan Sprague-Dawley (Indianapolis, IN). Operations and subsequent animal care were carried out in accordance with the guidelines of the University Committee on the Use and Care of Animals at the University of Michigan. At the time of surgery, the rats were anesthetized by ether inhalation. High in the right thigh, the sciatic nerve was exposed and tightly tied off with 7-0 silk suture in two places. The nerve was then severed between the sutures, and each cut end of the nerve was reflected and embedded into muscle masses proximally and distally in the thigh. This method routinely allows for permanent denervation (up to 25 months) without the need for repeat denervations. Postoperatively the rats were placed on oral terramycin for 5 days. All denervations reported in this paper were performed on 24-month-old rats.

The limb muscles were denervated for periods of 10 days and 1, 2, and 4 months. At the end of the experiment, the rats were again anesthetized with ether, and the denervated and control extensor digitorum longus (EDL) muscles were removed and prepared for in vitro analysis of contractile properties (see the following subsection). The rats were then euthanized with an overdose of ether, and the tibialis anterior, soleus, and gastrocnemius muscles were removed and prepared for morphological or molecular studies. Samples of denervated muscles were distributed to different laboratories for their respective analyses.

Contractile Properties
Rats were anesthetized with ether until they were unresponsive to tactile stimuli. Each control and experimental EDL muscle was exposed and dissected free of other tissue. Then 3-0 silk sutures were placed around the proximal and distal tendons, and the tendons were subsequently severed. The muscle to be tested was removed from the rat and secured at resting length in a tissue bath with a Krebs–Ringer bicarbonate solution containing 137mM NaCl, 5mM KCl, 2mM CaCl2, 1mM MgSO4, 1mM NaH2PO4, 24mM NaHCO3, and 0.025mM tubocurarine chloride. The solution was maintained at 25 ± 0°C and gassed with 95% oxygen and 5% carbon dioxide. At this temperature, 86-mg muscles of young rats remain completely stable and viable in structure and function even when required to make maximum contractions every 15 minutes (14). In contrast, muscles in the present study were only required to make a single maximum contraction, and experiments on single muscles were completed within 15 minutes.

One tendon of the muscle was tied to a fixed post and the other end to a Cambridge Technology servomotor–force transducer (Model 305, Cambridge, MA). Platinum electrodes were immersed in the bath on either side of the muscle. Muscles were stimulated by the current flow between the two electrodes by square-wave pulses that were 0.2 milliseconds in duration. The voltage was adjusted to produce a maximum isotonic twitch contraction and was then increased slightly. The length of the muscle was adjusted to the optimum length for the development of twitch force. The fiber length-to-muscle length ratio (Lf /Lo) has been determined for control EDL muscles of 4-month-old male WI/Hicks Car rats at Lf /Lo = 0.40 (15); 2-month-old male Sprague-Dawley rats at Lf /Lo = 0.40 ± 0.05 (14); and 8-month-old male Lewis rats at Lf /Lo = 0.35 ± 0.03 (16). The determination of fiber length is a laborious procedure that requires the total muscle (16). After physiological analysis, the same muscles were subdivided and distributed to various laboratories for subsequent morphological and molecular analysis. Consequently we were unable to determine fiber length on the muscles used in this series. The Lf /Lo does not vary greatly among the various strains of rats, and a value of 0.40 was used for the EDL muscles throughout this study.

Maximum isometric tetanic force (Po) was determined by maintaining the muscle at the length that produced optimal twitch force (17) and then increasing the frequency of stimulation from 80 Hz in increments of 20 Hz until the force plateaued at a maximum value. This value was defined as Po; Po was usually achieved at frequencies of stimulation between 100 and 140 Hz. After the measurement of contractile properties, the muscles were removed from the bath and weighed. Muscle fiber cross-sectional area (CSA) was calculated as CSA = muscle mass (g)/Lf (cm) x 1.06 g/cm2 (muscle density). The specific Po (kN/m2) was calculated by dividing the Po (kN) by the CSA (m2).

Biochemical and Molecular Analysis
RNA isolation and ribonuclease protection assay.-- Total RNA was isolated by homogenizing the muscles in Trizol (GIBCO BRL, Grand Island, NY) followed by the single-step purification method as described by the manufacturer's protocol. Antisense probes used to detect muscle creatine kinase (MCK), myogenin, and the {gamma}-subunit of the acetylcholine receptor were the same as those described by Adams and colleagues (18). Ribonuclease (RNase) protection assays were carried out as previously described (18). The probe for MCK was included in each experiment and served to normalize for differences in the amount of RNA in each of the samples, as has been reported previously. The RNA for MCK was not regulated by any of the conditions used in this report. The myogenin probe was made from mouse DNA and consistently protects two bands when hybridized to rat RNA. RNase-resistant hybrids were analyzed on 6% polyacrylamide, 8M urea gels. After electrophoresis, the gels were dried and exposed to x-ray film. Probe signals were quantified by scanning densitometry and values were normalized to the RNA signal obtained for MCK. The specificity of the protected bands was confirmed by hybridizing probes to tRNA, which resulted in no protected fragments on the gel. Probe integrity was monitored for each experiment by running an aliquot of nonhybridized probe on each gel.

Western blotting.-- Protein was extracted from skeletal muscle as previously described by Khalyfa and colleagues (19) and Kostrominova and colleagues (20). Briefly, rat muscles were dissected, frozen in liquid nitrogen, and stored at –70°C. Muscle tissue was homogenized in solution containing 20mM Tris-HCl (pH 6.8), 4% (wt/vol) sodiumdodecyl sulfate (SDS), 1mM of phenylmethylsulfonyl fluoride (PMSF) and 1µM each of leupeptin and Pepstatin A (for myogenin and cyclin D3 detection) or in buffer containing 300mM sucrose, 150mM KCl, 30mM Tris-HCl (pH 7.4), 5mM MgCl2, 1.5mM dithiothreitol, 1mM ethylenediamine tetra-acetic acid (EDTA), 1.5% Triton X-100, 10 µg/ml aprotinin, 3 µg/ml leupeptin, 2 µg/ml pepstatin, and 2mM PMSF (for eEF1A-1 and eEF1A-2/S1 detection). Protein concentrations were determined by using the Bio-Rad DC protein assay (Bio-Rad Laboratories, Hercules, CA). Protein samples were mixed with loading buffer, subjected to SDS–polyacrylamide gel electrophoresis (10%) and transferred electrophoretically to Immobilon-P membranes (Millipore, Bedford, MA). Gels with identical samples were stained with Coomassie Brilliant Blue dye and used as an additional control of equilibration of protein loading. After transfer, membranes were blocked in buffer containing 5% dry milk in phosphate-buffered saline/0.2% Tween-20 (PBST) or 10mM Tris-HCl (pH 7.5)/150mM NaCl/0.5% Tween-20 (TBST) and then incubated with primary antibody (overnight at 4°C or for 2 hours at room temperature). As previously described by Khalyfa and colleagues (21), highly specific polyclonal antibodies against eEF1A-1 and eEF1A-2/S1 proteins were used for detection. For detection of myogenin and cyclin D3 mouse monoclonal antibodies against myogenin (clone F5D, obtained from the Developmental Studies Hybridoma Bank, The University of Iowa, Iowa City, IA) and against cyclin D3 (NeoMarkers, Union City, CA) were used. Immunodetection was done by using peroxidase-conjugated goat anti-mouse antibody (Jackson ImmunoResearch Laboratory, West Grove, PA) with subsequent chemiluminescence (ECL, Amersham Pharmacia Biotech, Piscataway, NJ). Densitometric scans were taken from x-ray films of Western blots using a Molecular Dynamics densitometer SI (Sunnyvale, CA), and ImageQuaNT 4.1 software (Molecular Dynamics) was used for quantification.

Morphological Analysis
Immunohistochemical analysis.-- Following the measurement of contractile properties and the weighing of each muscle, the muscles were transversely divided into two parts and fixed for subsequent histological, immunocytochemical, and electron microscopic analysis. For histological and immunocytochemical analysis, muscle segments were fixed in 2% paraformaldehyde in 0.1M phosphate-buffered saline (PBS) at pH 7.4 for 24 hours and then washed overnight in 0.1M PBS. To prevent tissue dehydration and the formation of ice crystals during freezing, the muscles were cryoprotected with sucrose. After they were washed in PBS, the muscle segments were immersed in 0.25M sucrose in PBS for 1 hour, transferred to 0.5M sucrose in PBS for 45 minutes, and then left in 1.5M sucrose for 30 minutes. The cryoprotected muscle samples were then placed in specimen molds containing TBS/Tissue Freezing Medium (Triangle Biological Sciences, Durham, NC) and quick frozen by immersing the molds in 2-methylbutane (isopentane) that had been cooled in dry ice. Transverse serial 9-µm sections were cut on a Shandon cryostat (Life Science International Ltd., England) at –28°C, mounted on warm uncoated glass slides, and placed in a freezer at –20°C for storage. Before staining, the sections were rinsed in double-distilled water for 3 minutes at room temperature in order to remove the cryoprotective medium. They were then fixed in 100% methanol at –20°C for 10 minutes and allowed to air dry. The sections were then washed in 0.1M PBS for 4 minutes and incubated with 10% normal goat serum at room temperature.

The sections were double labeled with primary antibodies—a mouse antislow muscle myosin monoclonal antibody, clone NOQ7.5.4D (Chemicon International Inc., Temecula, CA) and a rabbit antilaminin polyclonal antibody (Sigma, St. Louis, MO). The sections were incubated for 3 hours in primary antibody followed by three 3-minute rinses in 0.1M PBS. Fluorescein-labeled goat antirabbit (Jackson ImmunoResearch) was used as the secondary antibody against the antilaminin primary antibody, and goat anti-mouse labeled with Rhodamine was used as the secondary antibody against the antislow muscle primary antibody. The mixture of secondary antibodies was incubated at room temperature for 45 minutes. This was followed by three 4-minute rinses in 0.1M PBS. The sections were then mounted with Vectashield Mounting Medium for fluorescence with DAPI (4,6-diamidino-2-phenylindole; Vector Laboratories, Burlingame, CA) and coverslipped. The sections were examined and photographed with Zeiss Axioplan 2 and Axiophot 2 Universal Microscopes (Carl Zeiss, Inc., Jena, Germany).

Light and transmission electron microscopy.-- For electron microscope analysis, small pieces from the muscles were fixed by immersion in a mixture of buffered formaldehyde/glutaraldehyde (2.5% each in 0.1M sodium cacodylate buffer at pH 7.4) at 4°C for 4 hours, washed with 0.25M sucrose in 0.1M PBS, and postfixed in 2% OsO4 in 0.1M PBS at 4°C for 1 hour. The muscles were then washed with 0.25M sucrose in 0.1M PBS, dehydrated in a graded series of ethanol and absolute acetone and then embedded in a mixture of Epon/Araldite (Eponate 12-Araldite 502 kit from Ted Pella, Inc., Redding, CA). Sections were cut with a Reichert–Jung Ultracut E ultramicrotome at a thickness of 0.5–1.0 µm and were mounted on glass slides. For general muscle structures to be observed at the light microscopic level, the sections were stained with Toluidine Blue. Ultrathin sections were collected on formvar-coated slotted grids and stained with uranyl acetate and lead citrate. The ultrathin sections were then examined under a Philips CM-100 transmission electron microscope (Eindhoven, The Netherlands).

Morphometry.-- Immunohistochemically stained sections of both control and denervated EDL muscles from young and old rats were examined with a Leitz Diaplan microscope, and the images were captured onto a Power Macintosh 8500/120 computer, under the same magnification, using a Pixera camera 1.2.4 (Pixera Corporation, Los Gatos, CA). Two images of the same region of the section were captured through the fluorescence microscope by using different filters for the fluorescein- and Rhodamine-labeled secondary antibodies. The double images of one microscopic field, stained by secondary antibodies with two different colors, were transformed into a single color image by using Adobe Photoshop 5.5 (Adobe Systems Inc., San Jose, CA). As a result, the type I (slow) fibers, which were stained red by Rhodamine, could be accurately recognized. The non-type I (presumably type II fast) muscle fibers were not stained by the antibody against the slow myosin heavy chain, but they were sharply outlined by the fluorescein-stained laminin in the basal laminae that surrounded all the muscle fibers. This technique permitted the two major types of muscle fibers to be distinguished within the same section and greatly facilitated quantitative analysis. The circumferences of both fiber types, delineated by the fluorescein staining of the secondary antibody to laminin that extended along the edge of all muscle fibers, were then electronically traced by using an ArtPad II and a graphics tablet with an Erasing UltraPen (Wacom Technology Co., Vancouver, WA). For each type of muscle fiber, the CSAs were calculated with the help of NIH Image 1.62f software (National Institutes of Health, Bethesda, MD). The distribution by CSA of both fast and slow muscle fibers was laid out in histograms.


   Results
Top
 Abstract
 Methods
 Results
Discussion
 References
 
Contractile Properties
As would be expected from denervation studies on young rats, both muscle mass and maximum tetanic force (Po) in old rats (24 months at the time of denervation) declined markedly during the postdenervation period (Table 1 ). At 1 and 2 months postdenervation, mean muscle mass was approximately half that of the contralateral controls. Mean Po declined much more rapidly than the mass. One month after denervation, the mean Po of the denervated EDL muscle had declined to 14% of the control level. By 4 months of denervation, the Po of the denervated muscle was only 3% of the control value. Specific force was 7% of control value at 4 months. The main difference between the postdenervation responses in young and old rats was the somewhat lesser decline in mass and force in the denervated old muscle (see Table 4 below).


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  		Table 1. Mass and Contractile Property Comparison Between Denervated (Den.) and Contralateral Control EDL Muscles of Old Rats

 

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  		Table 4. Comparison of Responses to Denervation of EDL Muscles Between Young and Old Rats

 
Biochemical and Molecular Analysis
Elongation factors eEF1A-1 and eEF1A-2 ..-- Elongation factor-1a (eEF1A-1) is a highly conserved ubiquitous protein involved in translation. eEF1A-2/S1 is a homologous gene to eEF1A-1, whose transcript and protein are found only in rat and mouse brain, heart, and skeletal muscle (essentially postmitotic tissues). This protein is only minimally present in the embryo (22).

During normal ontogenesis, the eEF1A-1/eEF1A-2 ratios undergo a marked change from a preponderance of eEF1A-1 in the prenatal and early postnatal period to a preponderance of eEF1A-2 during most of adult life (Table 2 ). However, during the latest phases of life, eEF1A-1 becomes increasingly strongly expressed, until by 34 months of age the eEF1A-1/eEF1A-2 ratio is greater than 1.


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  		Table 2. Ratios of eEF1A-1/eEF1A-2 Protein in Rat Skeletal Muscle During Normal Development and Aging

 
Both 10 days and 1 month following denervation in old rats, the eEF1A-1/eEF1A-2 ratio exceeded 1, whereas after 4 months of denervation it had dropped to 0.35 (Table 3 ), a level comparable with that of control muscle in rats of approximately the same age.


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  		Table 3. Ratios of eEF1A-1/eEF1A-2(S1) Protein in Denervated and Contralateral Control Muscles in Old Rats

 
Myogenin and cyclin D3.-- Myogenin is one of the members of the basic helix-loop-helix transcriptional factors that plays a major role in coordinating muscle development (23). Assays of both mRNA and protein showed a similar pattern of myogenin expression. RNase protection assays revealed a nearly 20-fold increase in myogenin mRNA abundance in old versus young control fast muscle (Fig. 1). Four months after denervation, myogenin mRNA increased to approximately the same absolute amount in both young and old rats, but the degree of increase was much greater in denervated muscle of young adult rats. Western blotting with an antibody against myogenin protein showed that unoperated control skeletal muscle from 24-month-old rats had approximately 1.5–2 times higher levels of myogenin protein than 4-month-old unoperated controls. By 10 days after denervation, myogenin protein increased in both young and old muscle to a similar level (approximately 8–9 times higher than young control levels) and stayed high throughout the 4-month duration of the experiment (Fig. 2).



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  		Figure 1. Relative changes in myogenin (Mgn) and nicotinic acetylcholine receptor (nAChR) gamma subunit mRNA during aging and after denervation. Presented are representative ribonuclease protection assays (A) and mean values (B) for Mgn and nAChR gamma subunit in young (Y-Inn) and old (O-Inn) innervated and young (Y-4mDen) and old (O-4mDen) 4-month denervated gastrocnemius muscles. For each protection assay, the levels of the respective RNAs were normalized to the level of expression of muscle-specific creatine kinase (MCK) mRNA. Bars represent means (n = 2) ± SD.

 


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  		Figure 2. Increase in levels of myogenin protein in gastrocnemius muscles from young (4-month-old) and old (24-month-old) rats after denervation. Expression of myogenin was evaluated in innervated and in 10-, 60-, and 120-day denervated muscles by Western blot analysis. A representative blot is shown. Three muscle samples per each time point were analyzed. Coomassie Blue-stained gels were used as an additional control for equal protein loading.

 
The mRNA abundance of the {gamma}-subunit of the acetylcholine receptor complex increased after denervation in a manner parallel to that of myogenin (Fig. 1). The {gamma}-subunit is an embryonic component of the acetylcholine receptor complex, and it is replaced with an {epsilon}-subunit when the embryonic muscle becomes innervated. In control muscle, the {gamma}-subunit is expressed minimally, if at all, whereas it is readily detectable in old control muscle (Fig. 1).

Cyclin D3 is an important regulator of myoblast differentiation (24). Whereas other cyclins (A, D1, D2) are downregulated during myogenic differentiation, cyclin D3 is strongly upregulated in cultured myotubes, in relation to levels in myoblasts (25). Following denervation, levels of cyclin D3 are not greatly increased at 10 days, but they are increased by approximately twofold to fourfold 60 and 120 days after denervation in both young adult and old animals (Fig. 3).



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  		Figure 3. Expression of cyclin D3 in gastrocnemius muscles from young (4-month-old) and old (24-month-old) rats after denervation. Representative Western blot analysis of cyclin D3 expression in innervated and in 10-, 60-, and 120-day denervated muscle and in primary cultures of rat myoblasts and myotubes is shown. Three muscle samples per each time point were analyzed. Coomassie Blue-stained gels were used as an additional control for equal protein loading.

 
Morphological Analysis
Histological cross sections of denervated muscles in young rats revealed a relatively homogeneous picture of atrophy at the tissue level (Fig. 4), whereas muscles in old rats denervated for a corresponding length of time showed considerable variation according to region in both the degree of muscle fiber atrophy and the amount of interstitial connective tissue (Fig. 4).



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  		Figure 4. Photomicrographs of 2-month denervated EDL muscle from 6-month-old (A) and 26-month-old (B–D) rats. Note the greater degree of variation in muscle fiber diameter and the increased amount of interstitial connective tissue in the old animals. Semithin sections were stained with Toluidine Blue. Bar = 25 µm.

 
One significant difference in denervated muscle between old and young rats was seen at the muscle fiber level. In young rats after 1 and 2 months of denervation, muscle fibers staining for slow (type I) myosin have a substantially larger mean CSA than fast (type II) muscle fibers (8). In contrast, the decline in mean CSA of the type I muscle fibers in old rats more closely paralleled that of the type II fibers (Fig. 5). Nevertheless, a small number of type I muscle fibers in 1-month-old denervated EDL muscles in old rats had much larger CSAs than their counterparts in the corresponding young group (Fig. 6).



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  		Figure 5. Relative changes in cross-sectional areas (CSAs) of types I and II fibers in denervated extensor digitorum longus (EDL) muscles of 6-month-old (A) and 26-month-old (B) rats. The basis for comparison of the muscle fiber CSAs is the mean CSA of control type II fibers from the same animals. *p <= .05; **p <= .01.

 


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  		Figure 6. Histograms illustrating changes in profiles of cross-sectional areas (expressed as standard units) of types I and II fibers in control and 1- and 2-month denervated extensor digitorum longus (EDL) muscles in young and old rats.

 
At the ultrastructural level, denervated EDL muscles in old rats showed similar characteristics to denervated muscles in young rats. Clusters of atrophic muscle fibers were often seen in association with larger fibers (Fig. 7), and activated satellite cells were encountered (Fig. 7). Increased interstitial collagen was readily apparent in many areas (Fig. 7, Fig. 7, and Fig. 7). As in denervated muscles in young rats, muscle spindles survived, but the intrafusal fibers underwent atrophy and the capsule loosened (Fig. 7).



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  		Figure 7. Electron micrographs of 2-month denervated extensor digitorum longus muscle in old (26-month-old) rats. A, Juxtaposition of muscle fibers of different cross-sectional areas; B, low-power view of a denervated muscle spindle; C, and D, activated satellite cells (s). Note the large amount of interstitial collagen (c) in D. A and B, bar = 10 µm; C and D, bar = 1 µm.

 

   Discussion
Top
 Abstract
 Methods
 Results
 Discussion
References
 
The principal reason for carrying out this study was to determine whether or not the response of skeletal muscle to denervation in old rats differs in any fundamental respect from that in young rats. The research literature over the past two decades has amply demonstrated that, during the aging process, increasing numbers of muscle fibers are subjected to either temporary or permanent denervation because of the death of alpha motoneurons that supply the motor units (2)(4). In addition, remodeling of individual neuromuscular junctions during the aging process (26) could potentially subject certain muscle fibers to transient denervation. Such changes in the innervation of muscle fibers in aging individuals appear to occur infrequently over extended periods of time, because bursts of denervation do not appear to occur during the normal aging process. Therefore, any denervation effects are likely to be cumulative and to take place against a background of largely innervated muscle fibers. How the muscle fibers of older animals respond to denervation could play a significant role in determining the overall functional role of denervation during the aging process.

The long-term responses of skeletal muscles to denervation in young adult animals have been well characterized in recent years. An initial burst of gene expression and satellite call activation during the first few days after denervation (20)(27)(28)(29) is followed by a period of rapid muscle fiber atrophy, resulting in a steep decline in both muscle mass and contractile force. By the end of the first month of denervation, an adult rat muscle has lost 70% of its mass and over 90% of its maximum tetanic force (7)(30). The first few months of denervation are characterized not only by muscle fiber atrophy, myonuclear death, and capillary loss (10)(11)(12), but by the formation of new muscle fibers in association with both intact and degenerating muscle fibers (12)(13)(31). Following the denervation of EDL muscles in rats, the percentage of satellite cells rises sharply during the first 2 months and then falls steadily over many months (9), whereas in the denervated soleus muscle it falls steadily without a temporary rise (12). Over the course of long-term denervation (many months), a variety of embryonic and fetal genes are reexpressed in the muscles (20)(21)(27)(29)(31).

Over a broad spectrum of analysis, the overall response to denervation of muscles in old rats does not differ substantially from that in young rats. The principal difference is a lesser degree of some measures of atrophy, especially among the type II fibers, in old as compared with young adult animals. The combined data allow interpretation of some of the observed old–young differences. The pattern of relative decline in mass, Po, and specific Po during the 1- and 2-month postdenervation period indicates a slower rate of atrophy in old than in young rats. Some of the lesser decline in mass in old EDL muscles (Table 4 ) might be due to the greater amount of interstitial connective tissue present in the old muscles. The reduced degree of type II muscle fiber atrophy, as measured by CSA in the old muscle fibers (Fig. 5), could also account for the somewhat greater mass and Po in old as compared with young denervated muscle (Table 4 ).

Other than the above-mentioned differences in rates of muscle fiber atrophy, the histological and ultrastructural findings did not reveal major differences in the reactions of young and old muscles to denervation. The somewhat greater variability in the histological picture of old denervated muscle is a reflection of the state of old control muscle, which contains more connective tissue and shows greater variations than young muscle in the spread of muscle fiber areas (32). Even the reaction of muscle spindles to denervation, such as hyperplasia of intrafusal fibers, is similar between old and young animals.

A general pattern of expression of developmentally active molecules in rodent muscle is a decline shortly after birth, low expression during most of adult life, and reexpression or higher levels of expression with increasing old age (20)(33)(34). As the animal approaches the end of its life span, the expression of important regulatory molecules, such as myogenin and MyoD, or their developmental isoforms, in the case of elongation factor and acetylcholine receptor subunits, increases sharply. Similarly, denervation of adult muscle results in an increase in the expression of a wide variety of regulatory molecules (18)(35)(36)(37)(38)(39). Interestingly, the motor unit remodeling that occurs during the aging process (1) could initiate changes in gene expression in muscles of old animals that reflect partial denervation, as well as the aging of innervated muscle fibers.

The molecular data collected in this study reflect this general pattern. Levels of both myogenin mRNA and protein are higher in 24-month-old than in 4-month-old control muscle. After denervation, levels of myogenin mRNA and protein rise sharply in young rats and to a lesser extent in old rats (20). Although myogenin levels do rise following denervation in old rats, the relative difference between control and denervated muscle is less, suggesting that the rise in levels in control muscle already reflects a situation of partial denervation and that the total capacity for myogenin expression in denervated old muscle is not greatly different from that in young muscle.

Levels of cyclin D3 likewise increase after denervation in both young adult and old rats. Contrary to many of the molecules that regulate the cell cycle, cyclin D3 is expressed at only low levels in myoblasts and is upregulated with the postmitotic fusion of myoblasts into myotubes (Fig. 3) (25)(37)(40)(41). On one hand, the increases in cyclin D3 levels in muscle at 2 and 4 months, but not at 10 days, after denervation could be a reflection of the timing of formation of new myotubes in the denervated muscle (31)(42). On the other hand, an increased expression of cyclin D3 in other cellular components of long-term denervated muscle in both young adult and old rats cannot be ruled out.

The reappearance of high levels of eEF1A-1, a developmental isoform of an elongation factor (22), becomes prominent late in the normal life of a rat (Table 2 ). Welle and colleagues (43) found no reduction from young adult levels in the expression of elongation factor-1{alpha} (eEF1A-1) or S1 (eEF1A-2) in a group of human subjects 61–74 years old. However, the age of their human subjects was not great enough to be equivalent to the 34-month-old rats, in which expression of eEF1A-1 increases substantially over normal adult levels (Table 2 ). eEF1A-1 expression also increases following denervation in both young (Khalyfa and Wang, unpublished data, 2002) and old (Table 3 ) rats, as well as during the regeneration of skeletal muscle in young adult rats (19). One explanation for the increased levels of eEF1A-1 in both very old and in denervated muscle is the formation of new muscle fibers, although increased expression in existing muscle fibers cannot be ruled out (31)(42). McGeachie (44) reported considerably increased levels of thymidine incorporation in peripheral nuclei of mouse muscle up to 4 weeks after denervation in the mouse. This could provide the basis for new muscle fiber formation. The formation of new muscle fibers could also account for increased expression of a large variety of molecules, including those reported here, that are normally expressed during various phases of embryonic myogenesis.

More difficult to document in all cases is the degree to which existing muscle fibers may begin to express embryonic markers during aging or after denervation. In the case of myogenin and other muscle regulatory factors, localization of the protein in the muscle fiber could be a reflection of the recent incorporation of a satellite cell that was expressing the protein in question at the time of fusion, or it could represent a change in gene expression within the original muscle fiber itself.

The molecules studied here—elongation factors, a myoregulatory protein, a receptor subunit, and a member of the cyclin family—represent different aspects of developmental control, yet their expression patterns followed a similar pattern. This suggests that both denervation and aging to different degrees call up broad developmental programs, the functions of which still remain poorly defined in these contexts. Of greatest relevance to this communication, denervation during old age does not appear to elicit reactions in the muscle that are substantially different from those that attend denervation in the adult period. Nevertheless, the lesser degree of atrophy, especially among the type II fibers, in denervated muscles of old rats may reflect significant age-based mechanisms.


   Acknowledgments
 
This research was supported by National Institutes of Health (NIH) Grant PO1 AG-10821. T. Kostrominova was supported by NIH Research Training Grant T32-AG00116.

We thank Jean Carlson and Richard Hinckle for technical support.

Received May 7, 2002

Accepted August 6, 2002


   References
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 Abstract
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 References
 

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