The Journals of Gerontology Series A: Biological Sciences and Medical Sciences 58:B984-B991 (2003)
© 2003 The Gerontological Society of America
Dynamics of Postdenervation Atrophy of Young and Old Skeletal Muscles: Differential Responses of Fiber Types and Muscle Types
Eduard I. Dedkov1,
Andrei B. Borisov1 and
Bruce M. Carlson1,2
1 Department of Cell and Developmental Biology
2 Institute of Gerontology, University of Michigan, Ann Arbor.
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Abstract
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We investigated the dynamics of muscle fiber atrophy in denervated fast and slow muscles of young and old rats. Hind limbs of 4-month-old and 24-month-old male rats were denervated, and soleus and tibialis anterior muscles were examined morphometrically 1 and 2 months after denervation. In all denervated muscles, type II muscle fibers underwent rapid atrophy, although muscle-specific differences in rate were observed. In both young and old denervated soleus muscles, the type I fibers underwent a pattern of atrophy closely paralleling that of the type II fibers, but in the tibialis anterior muscle, the mean cross-sectional area of the type I fibers actually increased during the first 2 months postdenervation. This study has shown that, among different muscles and between young and old rats, there is considerable variation in the response of the muscle fibers to denervation and that one cannot generalize from one muscle or one age to another.
THE aging of skeletal muscle is a complex process, with many concurrent themes. Although the dominant trend is toward progressive atrophy, ultimately leading toward sarcopenia at the whole body level (1), the cellular environment of an aging muscle is far from homogeneous. One of the major features of aging in skeletal muscle is the rearrangement of the pattern of innervation, with the death of some of the fast motor neurons leading to populations of denervated fibers within the aging muscle, and the sprouting of remaining neurons (mostly slow) to supply at least some of the physiologically denervated fast muscle fibers (2,3). Other suspected causes of muscle fiber atrophy during aging are reduced overall activity level (1,4,5), decreased levels of hormones and growth factors, such as insulin-like growth factor-1 (IGF-1) (6), or muscle fiber degeneration followed by repair (7).
Our laboratory has been systematically studying models of surgically induced denervation effects on both young and old muscle in an attempt to gain a greater understanding of the responses of old muscle fibers to physiological denervation (8–10). These studies have revealed substantial differences between the responses of fast and slow muscle fibers to denervation, as well as differences among various limb muscles in how the fast and slow muscle fibers react to denervation in both young and old rats.
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METHODS
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Animals
This study was conducted on young adult (4 months old) and old (24 months old) male WI/HicksCar rats maintained at the animal facility at the Department of Biology, University of Michigan. After ether anesthesia, the right sciatic nerve was tightly ligated with silk in two places and the nerve was cut between the sutures. Both proximal and distal nerve stumps were implanted into muscular tissue as far away from each other as possible. This method has produced permanent denervation of the hind limb muscles for as long as 25 months (10). At the University of Michigan, all manipulations and animal care were carried out in accordance with the guidelines of the Unit for Laboratory Animal Medicine. After operations, the rats were treated with oral terramycin for 5 days. Muscle excision and subsequent euthanasia of the animals were done under ether anesthesia 1 and 2 months after denervation. The tibialis anterior (TA) and soleus muscles were harvested from both denervated and intact contralateral legs of 3 experimental animals for each time point. Muscles were used for histological and immunohistochemical studies. The muscles, which were analyzed by more than one method, were cut into equal halves that were processed according to specific protocols. For morphometric analyses, cross-sections were prepared through the equatorial area of muscles and mounted in groups of 4 on histological slides. Three slides per time point were studied from each animal.
Immunohistochemical Analysis
The muscles were fixed in 2% paraformaldehyde in 0.1 M phosphate-buffered saline (PBS) at pH 7.4 for 24 hours and then washed overnight in 0.1 M PBS. To prevent tissue dehydration and the formation of ice crystals during freezing, the muscles were cryoprotected with sucrose. After they were washed in 0.1 M PBS, the muscle segments were immersed in 0.25 M sucrose in PBS for 1 hour, transferred to 0.5 M sucrose in 0.1 M PBS for 45 minutes, and then left in 1.5 M 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., U.K.) 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 (RT) 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 0.1 M PBS for 4 minutes and incubated with 10% normal goat serum at RT. The sections were double-labeled with a mixture of primary antibodies—a mouse antihuman skeletal myosin (slow), clone NOQ7.5.4D (Chemicon International, Inc., Temecula, CA), and a rabbit antimouse laminin (Sigma, St. Louis, MO)—for 3 hours at +37°C. After incubation, the sections were washed in 0.1 M PBS by three 3-minute rinses, incubated with 10% normal goat serum for 10 minutes, and stained with a mixture of secondary antibodies for 45 minutes at RT. This was followed by three 4-minute rinses in 0.1 M PBS. Rhodamine and fluorescein isothiocyanate (FITC)-conjugated goat antimouse and goat antirabbit secondary antibodies (Jackson ImmunoResearch Laboratories, Inc., West Grove, PA), respectively, were used for visualization of primary antibodies. For control, omissions as well as substitution with normal goat serum of one or both primary antibodies were used. The sections were then mounted with Vectashield Mounting Medium for fluorescence with DAPI (4,6-diamidino-2-phenylindole; Vector Laboratories, Burlingame, CA) to counterstain nuclei and were cover-slipped. The sections were examined with Zeiss Axiophot 2 Universal Microscope (Carl Zeiss, Inc., Jena, Germany), and images were captured by using a Zeiss Axiocam digital camera. Figures were prepared from electronic images using Adobe Photoshop software (Adobe Systems, Inc., San Jose, CA).
Morphometry
Immunohistochemically stained sections of both control and denervated 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-labeled and Rhodamine-labeled secondary antibodies (fluorescein-labeled goat antirabbit antibody was used as the secondary antibody against the antilaminin primary antibody, and goat antimouse antibody labeled with Rhodamine was used as the secondary antibody against the antislow myosin primary antibody). The double images of 1 microscopic field, stained by secondary antibodies with 2 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 nontype 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 2 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 cross-sectional areas (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. The mean values (in %) of CSAs of both fast and slow muscle fibers in the tibialis anterior are expressed as the ratio of their CSA to the CSA of control type II fibers (the predominant type of fiber in fast muscles). The mean values (in %) of CSAs of both fast and slow muscle fibers in the soleus muscle are expressed as the ratio of their CSA to the CSA of control type I fibers (the predominant fiber type in slow muscle).
Statistical Analysis
Quantitative data were analyzed with a two-way analysis of variance (ANOVA) followed by the Student's t test (unpaired sample). The values are expressed as means ± SEM (standard error of mean). The level of significance between control and denervation was set at **p 
.01, ***p .001.
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RESULTS
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In all muscles in both young and old rats, the first 2 months after denervation were characterized by a steady decline in the mean cross-sectional areas of the type II muscle fibers (Figure 1). For both young and old TA muscles, the type II fiber area was, respectively, 30.7% ± 1.6% and 28.5% ± 0.9% of control value by the second month after denervation, and in the soleus it had dropped to 22.7% ± 1.3% of control in the young adult and 13.1% ± 0.9% in the old. This was not the case for the type I fibers. Although in the soleus muscles the pattern of decline in cross-sectional area of the type I fibers closely paralleled that of the type II fibers, the pattern in the TA muscles was markedly different. In denervated young adult rat TA muscles, the mean cross-sectional areas of type I muscle fibers rose from 82.9% ± 7.1% of the reference standard in controls to 97.2% ± 6.1% and 95.3% ± 5.6% at 1 and 2 months after denervation (Figure 1). In contrast, in TA muscles from old rats, the type I fibers underwent atrophy at roughly the same rate as the type II fibers, reaching 33.2% ± 3.7% of control values at 2 months of denervation. A previous study of the extensor digitorum longus (EDL) muscle showed a pattern of differential fiber type atrophy similar to that of the TA muscle, except that in old rats the type I muscle fibers underwent atrophy at a slower rate than did the type II fibers (9) (Figure 1). The rate of atrophy of both types I and II fibers in the young adult soleus was considerably more rapid than that in the old soleus muscles.
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Figure 1. Graphical representation of the relative cross-sectional areas (in standard units) of types I and II muscle fibers of the extensor digitorum longus (EDL), tibialis anterior (TA), and soleus muscles after denervation. For the fast EDL and TA muscles, the mean cross-sectional area of control type II fibers was used as the 100% reference point for both type I and II fibers, whereas in the slow soleus muscle, the mean cross-sectional area of control type I fibers was used as the reference point. [Data for EDL muscle fibers are taken from Ref. 9 (Carlson et al., 2002).]
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Frequency distributions of fiber areas in young adult TA muscles show not only a clear-cut atrophy of the type II fibers, but a narrowing of the range of cross-sectional areas during the 2-month denervation period (Figure 2). In contrast, the population of type I fibers was characterized by not only increasing mean cross-sectional areas, but a considerably greater spread of cross-sectional areas than was the case in control muscle. In old rats, the frequency distributions showed a pattern of atrophy of type II fibers that was very similar to that seen in young rats (Figure 2), and the shift in patterns of the type I fibers closely paralleled that of the type II fibers. The patterns of atrophy of types I and II fibers in the denervated young soleus muscle were very similar in terms of both the overall trend of atrophy and the spread of cross-sectional areas (Figure 3). In denervated old soleus muscles, only a minor tailing to the right in the distribution of fiber areas 1 month after denervation distinguished the type I from the type II fibers (Figure 3).
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Figure 2. Frequency distributions (in standard units) of cross-sectional areas of type I and type II fibers from control, 1-month, and 2-month denervated tibialis anterior (TA) muscles from young and old rats
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Figure 3. Frequency distributions (in standard units) of cross-sectional areas of type I and type II fibers from control, 1-month, and 2-month denervated soleus muscles from young and old rats
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The histology of the 2-month denervated TA muscle in young rats clearly shows the reduction in area of the type II muscle fibers in relation to that of the type I fibers (Figure 4A and B). This is in contrast to old denervated muscle, in which the type I muscle fibers are only slightly larger than the type II fibers 2 months after denervation. In the old TA muscles, the slow muscle fibers in control muscles were often bunched together into type groups (11) (Figure 5A and B), but after 2 months of denervation, type groups were not seen (Figure 5C and D). Instead, an increased number over control of slow (type I) muscle fibers was scattered as individual muscle fibers, which at this point differed little in size from the type II fibers (Figure 6).
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Figure 4. Immunohistochemical preparations showing type I muscle fibers (gray) in relation to type II fibers (black, outlined in gray) in control (A and C) and 2-month denervated (B and D) tibialis anterior muscles of young (A and B) and old (C and D) rats. The type I muscle fibers are stained with an antibody against slow myosin, whereas the type II fibers are not directly stained, but are bounded by laminin-stained basal lamina material. Bar = 50 µm
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Figure 5. Immunohistochemical preparations showing the type-grouping of type I fibers (gray) in old control tibialis anterior muscle (A) and the absence of type grouping in 2-month denervated old tibialis anterior muscle (C). Type I muscle fibers are stained with an antibody against slow myosin in A and C. B and D show the same fields as A and C, but the sections were stained with an antilaminin antibody, which outlines the basal lamina around all muscle fibers. Bar = 50 µm
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Figure 6. Low-power survey of the scattered distribution of type I muscle fibers in 2-month denervated old rat tibialis anterior muscle. The type I fibers in A were stained with an antibody against slow myosin, whereas in B the section was stained with antilaminin antibody. Bar = 50 µm
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DISCUSSION
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The results of this study on the denervated TA and soleus muscles, along with the previously reported data on the EDL muscle (9), clearly show that the responses of muscle fiber populations to denervation vary according to both the type of muscle and age of the animal. The greatest differences were seen between the fast and slow muscles, but even between the two fast muscles examined, minor differences in patterns of atrophy were observed. Similarly, Tomanek and Lund (12) noted differences in the rate of atrophy of muscle fibers between the soleus and the red and white portions of the vastus lateralis muscles of the guinea pig. A differential response of individual muscles in response to a broad environmental change is not confined solely to denervation. LeBlanc and colleagues (13) recorded considerable variation in the decline of volumes of a variety of human muscles to bed rest, as measured by magnetic resonance imaging. At the molecular end of the spectrum, Voytik and colleagues (14) and Walters and colleagues (15) recorded variations in the expression of myogenic regulatory factors among several denervated muscles in rodents. Similarly, several investigators have noted that, during aging, levels of gross atrophy differ among different muscles (16–18). Considerable variation at the histological level has been noted among human muscles by Thornell and colleagues (19).
The results noted here and earlier (for the EDL muscle) clearly show that fast and slow muscle fibers are differentially affected by denervation and aging. In the denervated EDL muscle of young rats, type II fibers undergo severe atrophy during the first 2 months following denervation, whereas type I fibers not only maintain, but as a group increase, their mean cross-sectional area (9,20). This same pattern is followed by type I fibers in the TA muscle in young rats.
The young soleus muscle, which is highly reactive to environmental changes, rapidly loses mass during the first month during denervation. This is reflected in the great decreases in cross-sectional area of both types I and II fibers (Figure 1).
The relative loss of fiber area of type II fibers is reduced in EDL muscles of old as compared with young rats (9), and a similar reduction in the rate of atrophy of both types I and II fibers occurs in the denervated old soleus muscle (Figure 1). Ansved (21) noted a similar reduction with age in the rate of fiber atrophy of soleus muscles in limbs immobilized by plaster casts. Similar to our results here, d'Albis and colleagues (22) reported the hypertrophy of many slow muscle fibers in denervated rabbit muscles. It is clear, however, that the reaction of muscle fibers to denervation relates more closely to the type of muscle than to the type of muscle fiber. In the young soleus muscle, the rate of atrophy of the type I fibers paralleled that of the type II fibers, in contrast to the hypertrophy seen in the denervated young fast muscles.
In our old muscles, type-grouping of type I fibers was common in the old fast muscles. This is probably a reflection of the loss of fast motoneurons and the capture of denervated fibers by sprouts emanating from the axons supplying slow motor units, as has been shown by a number of investigators (3). Interestingly, type-grouping was not seen in 2-month denervated muscles in the old rats. One possible explanation is that fast muscle fibers newly captured by slow nerves may have reverted to their original type once the neural input was removed. It is of note that the absolute numbers of type I fibers increased during the 2 months following denervation. This phenomenon was also seen by d'Albis and colleagues (22), who attributed this to the transformation of fast into slow muscle fibers in denervated muscles of rabbits.
Although the nature of the studies reported here does not allow mechanistic explanations for the observed phenomena, our data clearly show significant differences among both muscles and age groups in response to denervation. Such results show that caution should be used in extrapolating results from one muscle to the other and from one age group to the next.
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Acknowledgments
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This research was supported by National Institutes of Health (NIH) grant PO1 AG-10821.
Address correspondence to Dr. Bruce M. Carlson, Institute of Gerontology, 300 N. Ingalls Bldg., Room 913, University of Michigan, Ann Arbor, MI 48109. E-mail: brcarl{at}umich.edu
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Footnotes
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James R. Smith,, PhD, Decision Editor
Received May 30, 2003
Accepted August 12, 2003
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