Journal of Applied Physiology Journal of Applied Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

J Appl Physiol 100: 2012-2023, 2006. First published February 23, 2006; doi:10.1152/japplphysiol.01583.2005
8750-7587/06 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
100/6/2012    most recent
01583.2005v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (4)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by McClung, J. M.
Right arrow Articles by Carson, J. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by McClung, J. M.
Right arrow Articles by Carson, J. A.

Estrogen status and skeletal muscle recovery from disuse atrophy

J. M. McClung,1,2 J. M. Davis,2 M. A. Wilson,3 E. C. Goldsmith,4 and J. A. Carson1,2

1Integrative Muscle Biology Laboratory, Division of Applied Physiology, and 2Department of Exercise Science, University of South Carolina; and Departments of 3Pharmacology, Physiology, and Neuroscience and 4Developmental Biology and Anatomy, University of South Carolina Medical School, Columbia, South Carolina

Submitted 16 December 2005 ; accepted in final form 14 February 2006


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Although estrogen loss can alter skeletal muscle recovery from disuse, the specific components of muscle regrowth that are estrogen sensitive have not been described. The primary purpose of this study was to determine the components of skeletal muscle mass recovery that are biological targets of estrogen. Intact, ovariectomized (OVX), and ovariectomized with 17beta-estradiol replacement (OVX+E2) female rats were subjected to hindlimb suspension for 10 days and then returned to normal cage ambulation for the duration of recovery. Soleus muscle mass returned to control levels by day 7 of recovery in the intact animals, whereas OVX soleus mass did not recover until day 14. Intact rats recovered soleus mean myofiber cross-sectional area (CSA) by day 14 of recovery, whereas the OVX soleus remained decreased (42%) at day 14. OVX mean fiber CSA did return to control levels by day 28 of recovery. The OVX+E2 treatment group recovered mean CSA at day 14, as in the intact animals. Myofibers demonstrating central nuclei were increased at day 14 in the OVX group, but not in intact or OVX+E2 animals. The percent noncontractile tissue was also increased 29% in OVX muscle at day 14, but not in either intact or OVX+E2 groups. In addition, collagen 1a mRNA was increased 45% in OVX muscle at day 14 of recovery. These results suggest that myofiber growth, myofiber regeneration, and extracellular matrix remodeling are estrogen-sensitive components of soleus muscle mass recovery from disuse atrophy.

steroid hormone; hypertrophy; muscle regeneration; fibrosis; extracellular matrix

ALTHOUGH THE CAPACITY FOR estrogen synthesis varies throughout a female's lifetime, the consequences these hormonal changes have on skeletal muscle's ability to regulate plasticity and mass are just beginning to be understood. Skeletal muscle myoblasts, myotubes, and mature fibers express functional estrogen receptors, suggesting that skeletal muscle is sensitive to estrogen steroid hormone signaling (33, 55). Estrogens can regulate skeletal muscle mass in developing livestock (60) and rats (37, 58). The depletion of ovarian hormones due to ovariectomy induces an anabolic environment characterized by increased circulating growth factor levels and a positive energy balance (23, 63). This environment manifests itself in pronounced increases in female rat organ weight, skeletal muscle mass, and body mass (23). Estradiol administration attenuates the ovariectomy-induced body mass and muscle protein accretion (54). Estrogen replacement in ovariectomized rats subjected to hindlimb disuse attenuates bone atrophy, but soleus muscle atrophy is not prevented (23, 34). After extended periods of hindlimb disuse, ovariectomy decreases rat skeletal muscle mass recovery, and estrogen replacement benefits atrophied muscle mass recovery (13, 57). These facts suggest an interaction between load-sensing mechanisms and estrogen signaling pathways that benefits skeletal muscle growth. However, estrogen-sensitive components of muscle mass recovery have not been clearly identified.

Skeletal muscle gene regulation is modulated by the load placed on the muscle (6). Several days of disuse can initiate biochemical and morphological changes related to muscle atrophy, protein isoform shifts, and metabolic enzyme alterations (21, 61). Normal weight-bearing activity is a therapeutic treatment for muscle mass restoration after extended periods of disuse (42). During the initial return to normal ambulation processes related to protein accretion, myofiber damage and subsequent regeneration are induced (31, 49). Characteristics of regenerating muscle during recovery from atrophy include centralized nuclei, small sized myofibers, and embryonic myosin heavy chain protein expression (17, 31). Skeletal muscle satellite cells, which become quiescent during hindlimb suspension (19), are activated during recovery from disuse (49) and provide a critical myonuclei source for muscle mass recovery (8). Recovery of muscle mass is an extremely rapid process in the healthy adult and is normally complete after 14 days, even when disuse periods are extended significantly beyond 10 days (10, 11, 49). However, processes related to complete regeneration can extend out to 5 wk (31).

Extracellular matrix remodeling is a critical process related to both skeletal muscle growth and regeneration from injury (36). Extracellular matrix components provide critical structural support, scaffolding for cellular mobility, elasticity, and initiate intracellular signaling cascades in skeletal muscle (36). During periods of extended disuse, the muscle's relative proportion of extracellular matrix volume remains constant (48), whereas the extracellular matrix composition is altered. Atrophying soleus muscle undergoes a collagen isoform shift from type I to type III (48). Reloading after disuse induces alterations in the expression of the extracellular matrix proteins tenascin-C and fibronectin, which serve structural and elastic functions in skeletal muscle (25). Fibrosis, overexpansion of the extracellular matrix, can attenuate skeletal muscle mass recovery and myofiber regeneration after injury-induced damage (56). Fibroblasts contribute to the proportion of mitotically active cells during periods of increased loading and are primarily responsible for extracellular matrix protein synthesis (18, 25). Fibrosis and extracellular matrix remodeling can be regulated by hormone, growth factor, and inflammatory cytokine signaling (38). Additionally, the transforming growth factor-beta (TGF-beta) signaling pathway appears to be critical for regulation of extracellular matrix remodeling (38). Although estrogen is a modulator of TGF-beta signaling (43), estrogen's effect on fibrosis during skeletal muscle regeneration has not been established.

Estrogen has the capability to regulate several processes associated with skeletal muscle mass accretion and regeneration, which point to its potential for modulating muscle recovery from disuse atrophy (17, 39). Components of overall muscle mass recovery that are potentially estrogen sensitive include extracellular matrix remodeling, myofiber regeneration and growth, inflammation, and sarcolemma damage (39). Although estrogen loss can alter skeletal muscle recovery from disuse (13, 57), the components of wet mass recovery that are estrogen sensitive are not known. The primary purpose of this study was to determine the specific components of soleus muscle mass recovery that are biological targets of estrogen. An additional purpose was to determine whether estrogen- sensitive alterations occur specifically during the atrophy or recovery process. We hypothesized that estrogen loss would attenuate myofiber growth, extracellular matrix remodeling, and indexes of muscle regeneration during muscle recovery from disuse. Understanding the targets of estrogen in recovering skeletal muscle may lead to enhanced therapeutic treatments for muscle mass restoration after disuse in many clinical settings.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Animals and housing.   Intact and ovariectomized (OVX) female Sprague-Dawley rats (~200 g) were acquired from Harlan rodent colony (Indianapolis, IN) and randomly divided into four separate day-of-recovery treatment groups (n = 6/group; Fig. 1). These groups included 1) ground control (Con), 2) hindlimb suspension (Sus), 3) suspension with 7 days of reload (Sus+7), and 4) suspension with 14 days of reload (Sus+14). To determine whether OVX female animals exhibit delayed or deficient skeletal muscle regrowth after disuse, a subset of animals were randomly divided into ground control (n = 6) and suspension with 28 days of reloading (n = 6; Sus+28). The role of estrogen (OVX+E2) during both suspension and reload in recovery from atrophy was determined using six groups of OVX females: 1) OVX ground control receiving placebo (Con), 2) OVX ground control with exogenous estrogen for suspension and 14 days of reload (Con All), 3) OVX suspension with estrogen (Sus+E2), 4) OVX suspension with estrogen for suspension and 14 days of reload (Sus+14 All), 5) OVX ground control with estrogen for 14 days of reload (Con Rld), and 6) OVX suspension with estrogen for 14 days of reload (Sus+14 Rld). Animals were housed individually and kept on a 12:12-h light-dark cycle and given ad libitum access to normal rodent chow (intact female treatment groups) or phytoestrogen free rodent chow (Purina Test Diet, Richmond, IN; OVX treatment groups) and water for the duration of the study at the fully accredited animal care facilities at the University of South Carolina, Columbia. At the conclusion of the treatment, animals were killed under anesthesia and the soleus muscle was frozen in liquid nitrogen for analyses. Daily vaginal smears (5 days in length) were performed on all animals at the onset of the study to verify estrous cycling in intact females and a lack of estrous cycling in OVX animals. All procedures were approved by the University of South Carolina Institutional Animal Care and Use Committee.


Figure 1
View larger version (8K):
[in this window]
[in a new window]
  		Fig. 1. Study design. A: intact and ovariectomized (OVX) females were subjected to hindlimb suspension (Sus) and 7, 14, or 28 days of reloading recovery (see METHODS). B: ovariectomized females were administered estrogen for the duration of hindlimb suspension (OVX+E2 Sus), before hindlimb suspension for suspension with 14 days of reloading (OVX+E2 All Days), or at the time of reload from suspension with 14 days of reloading (OVX+E2 Rld Days). Con, ground control.

 
Hindlimb suspension disuse and reload.   Hindlimb suspension-induced disuse and reload recovery were performed as previously described (45), after the initial week of animal monitoring.

Estrogen replacement.   The appropriate OVX treatment groups were implanted with subcutaneous 60-day hormone release pellets (Innovative Research of America) containing 0.25 mg of 17beta-estradiol, the major estrogen in mammalian systems. Appropriate control OVX females were implanted with placebo pellets. The dose of estrogen selected in the present investigation was based on previous work done with physiological estrogen supplementation in the rat model and is designed to provide a constant 40 µg·kg body wt–1·day–1 (52, 59).

Plasma estradiol analysis.   Plasma estradiol was extracted from 500 µl of plasma and measured by use of a commercially available double antibody radioimmunoassay kit (Diagnostic Products, Los Angeles, CA) as per manufacturer's recommendations. Duplicate determinations were made for plasma samples from all treatment groups. The intra- and interassay coefficients of variation were 6.2 and 14.3%, respectively.

Muscle dry weights and water content.   The soleus muscle was analyzed for dry muscle mass and water content as previously described (51) with the following modifications. Briefly, ~30 mg of total soleus muscle wet mass was weighed, subjected to a drying protocol of 70°C, and weighed every 30 min until a stable dry weight was obtained.

Muscle myofibrillar protein.   Total and myofibrillar protein were determined as previously described (1) with the following modifications. Frozen muscle was homogenized in ice-cold buffer A (100 mg/ml) containing 100 mM KCl, 20 mM imidazole, and 5 mM EDTA (pH 6.8) for 20 s by using a Polytron homogenizer on high speed. One-third of the homogenate was stored at –80°C until used for analysis of total muscle protein, and the remaining two-thirds were used for isolation of myofibrils, as follows. The muscle homogenate was centrifuged at 1,000 g for 10 min at 4°C. The supernatant was discarded, and the resulting myofibril pellet was resuspended in 5 ml of buffer B (175 mM KCl containing 0.5% Triton X-100, pH 6.8). After centrifugation at 1,000 g for 10 min at 4°C, the pellet was washed twice with buffer B to remove cellular membranes. The resulting pellet was washed twice in buffer C (150 mM KCl, 20 mM imidazole, pH 7.0) to remove residual Triton-X and then resuspended in 0.5 ml of buffer D, an aqueous solution of 5 mM EDTA, pH 7.4/100 mg original muscle powder. Myofibril suspensions were stored at –80°C until analyzed. The protein content of total muscle and myofibril suspensions was determined using the Bradford assay (Bio-Rad).

Morphological analyses.   For cross-sectional area (CSA; µm2), centralized nuclei, and noncontractile tissue percentage (%NCT) analyses, four distinct digital images from hematoxylin-eosin-stained muscle sections (10 µm) from the midbelly of the soleus muscle at an objective magnification of x20 were obtained and analyzed as previously described (2, 47).

Crude protein extracts.   Crude protein extracts were made from frozen soleus muscles by homogenization in Mueller buffer on ice with a Polytron homogenizer using 3 x 15-s pulses at a low setting as previously described (46).

Western blot analysis.   p70s6k protein and phosphorylation levels were determined in soleus skeletal muscle as previously described (46). After transfer, membranes were probed with p70s6k primary antibody for total protein level (Santa Cruz, C-18) and phosphorylated p70s6k (Thr 389) protein primary antibody (Cell Signaling Technology, no. 9205), diluted in 1% milk-TBS-Tween. The horseradish conjugated anti-rabbit secondary antibody, diluted in 1% milk-TBS-Tween (1:6,500) and conjugated with alkaline phosphatase, was visualized by chemiluminescence (ECL, Amersham Life Sciences) as per manufacturer instructions and quantified by densitometry scanning (Scion Technologies, Frederick, MD).

Total RNA isolation and cDNA synthesis.   Total RNA was isolated by using TRIzol reagent (Life Technologies, Grand Island, NY) as per manufacturer's instructions.

Northern blot analysis.   Northern blot analysis was performed as previously described (10). Briefly, 15 µg of total RNA were fractionated on a denaturing 1% agarose gel (1 x MOPS, 6.7% formaldehyde) and then transferred to a nylon membrane by capillary action. The collagen type Ia probe was a kind gift from Dr. Wayne Carver (Department of Developmental Anatomy and Biology, University of South Carolina School of Medicine). Membranes were then visualized by autoradiography (–80°C, 3–40 h), and quantified by densitometry scanning (Scion Image, Frederick, MD) obtaining an integrated optical density (IOD), which was used to calculate mRNA abundance. mRNA abundances were corrected for 18S IOD calculated from digital gel images before transfer and subsequently normalized to intact female ground control values.

Immunohistochemical analysis.   For the analysis of soleus muscle embryonic myosin heavy chain (eMHC), transverse sections (10 µm) were cut from the midbelly of the soleus muscle on a cryostat at –20°C and mounted on coverslips. Immunohistochemistry staining was performed as previously described (27–29). Briefly, muscle sections were air-dried and fixed in acetone, and endogenous peroxidase activity was quenched by incubation in 0.6% peroxide. Sections were blocked with 4% horse serum and exposed to anti-eMHC (Developmental Studies Hybridoma Bank, Iowa City, IA). Control sections were incubated with 1x PBS only. Sections were washed in 1x PBS and anti-mouse secondary antibody (1:200) (Vector Laboratories) was incubated on all sections (control and treatment) for 90 min at 37°C. Sections were then incubated with the Vectastain ABC reagent (Vector Laboratories) per manufacturer instructions. Sections were then exposed to diaminobenzidine tetrahydrochloride with nickel (Vector Laboratories). Positive fibers were counted under a light microscope and expressed as the total number of eMHC-positive fibers per square millimeter muscle.

Data analysis.   Differences within treatment control groups due to recovery day (10-day Sus, Sus+7, Sus+14, or Sus+28), circulating 17beta-estradiol, and differences in estrogen-replacement controls or reloaded females due to the day of estrogen administration (All days or Rld days) were analyzed by one-way ANOVA. Where no significant effects occurred in treatment controls due to day or in estrogen-replacement controls and reloaded females due to the day of estrogen administration, respective groups were pooled. All other variables were analyzed by two-way ANOVA for main effects (estrogen treatment or recovery day) or interactions (estrogen treatment x recovery day). Where main effects existed devoid of significant interactions, values are presented as pooled. Where significant interactions existed, Bonferroni post hoc analyses were used between groups. Values are presented as means ± SE. Significance was set at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Plasma estradiol.   Ovariectomy (3 ± 1 pg/ml) decreased circulating estradiol 85% compared with intact values (20 ± 7 pg/ml). Ten days of hindlimb suspension did not alter circulating estradiol (32 ± 1 pg/ml) in intact females. Estrogen replacement with 17beta-estradiol administration increased circulating estradiol (67 ± 27 pg/ml) above intact and ovariectomy values.

Analysis of day of controls and estrogen administration timing.   The experimental design of the present study had cage-control rats for each estradiol treatment at each day of reload. In control rats, within intact, ovariectomy, or ovariectomy plus estradiol treatments, there were no effects of recovery day for any morphological or biochemical measurement analyzed in the present study. Control data within treatments were combined for all further analysis.

The experimental design also had two estrogen-replacement treatments given to ovariectomized rats, which varied by the time the estrogen was administered. The first estrogen-replacement treatment had estrogen administration given for the entire experimental time course (disuse and recovery), whereas the second treatment was administered at the time of atrophy recovery after the completion of hindlimb disuse. There were also no observed effects of estrogen treatment timing (entire time course or given at recovery) for any morphological or biochemical measurement analyzed in the present study. As such, data were combined for all further analysis and discussion.

Muscle mass recovery after disuse.   Muscle mass was significantly altered by estrogen manipulation and recovery day, which had a significant interaction on muscle wet weight (Table 1). Ovariectomy alone increased soleus muscle wet weight 28% from intact controls and estrogen replacement attenuated this increase (Table 1). There was significant atrophy of the soleus muscle with hindlimb disuse in all treatments groups. After 10 days of disuse soleus wet weight decreased 27% in intact, 44% in ovariectomy, and 41% with estrogen-replacement treatments (Table 1). Muscle wet weight recovery was accomplished by the 7th day of recovery in intact animals. With ovariectomy, soleus wet weight remained decreased 34% at day 7 and remained 12% less than control values after 14 days of recovery. However, ovariectomized soleus wet weight was fully restored (123 ± 6 mg) by day 28 of recovery.


View this table:
[in this window]
[in a new window]
  		Table 1. Muscle wet weights, wet weights corrected for tibia lengths, myofibrillar protein, water content, and percent noncontractile tissue in intact, ovariectomized, and ovariectomized with estrogen replacement rat soleus muscle recovering from hindlimb suspension disuse

 
Because ovariectomy can induce changes in body mass, tibia lengths (mm) were used to correct for alterations in body size. There was no interaction of estrogen treatment and recovery day on tibia lengths (35.3 ± 1.6 mm). Soleus mass was then corrected for tibia length (mass/tibia). Estrogen treatment and recovery day had a significant interaction on the mass-to-tibia ratio (Table 1). Ovariectomy alone increased muscle mass/tibia length 29% compared with intact animals and estrogen replacement attenuated this increase. Hindlimb disuse decreased the muscle mass-to-tibia ratio 25% in intact, 42% with ovariectomy, and 38% in estrogen-replacement muscle (Table 1). The muscle mass-to-tibia ratio was restored by 7 days of recovery in intact females, but in ovariectomized females it remained decreased 31 and 8% from ground control at days 7 and 14 of recovery, respectively. Ovariectomized rat mass-to-tibia ratio was restored to control values by the 28th day (3.4 ± 0.1 mg/mm) of recovery.

Muscle dry weight, water content, and total muscle myofibrillar protein.   Estrogen status did not alter muscle dry weight (mg). However, there was a main effect of recovery day on muscle dry weight. Suspension (15.9 ± 1.3 mg) decreased dry weight 40% compared with control (26.6 ± 1.3 mg) values. Dry weight remained decreased 39% from control values after 7 (17 ± 4 mg) or 14 days of recovery (20.7 ± 1.2 mg).

Estrogen status altered soleus muscle water content. Soleus muscle from ovariectomized rats had a 39% increase in water content compared with the intact control muscle (Table 1). Estrogen replacement returned muscle water content to intact control levels. Intact muscle water content decreased 21% with suspension but was restored to control values at days 7 and 14 of recovery. In ovariectomized rats, soleus water content decreased 44% with suspension and remained decreased 32% after 7 days of recovery, but it returned to control values by day 14 of recovery. Estrogen replacement did not affect muscle water loss with suspension, as suspension decreased muscle water content 43%.

Estrogen status did not alter the loss or recovery of myofibrillar protein in the present study (Table 1). Suspension decreased myofibrillar protein 60% in the intact animals and 57% in the ovariectomized animals. Myofibrillar protein remained decreased until day 14 of recovery in both treatment groups. Animals receiving estrogen replacement also had a similar loss (50%) of myofibril protein with suspension, which was restored after 14 days of recovery. There was a main effect of treatment day on soleus muscle total protein content. Hindlimb suspension (13 ± 2 mg) decreased muscle total protein content 54% from control (25 ± 2 mg), and it remained decreased 35% with 7 (16 ± 3 mg) and 26% with 14 days (19 ± 1 mg) of recovery. There was no interaction of estrogen status and recovery day on muscle total protein content.

Myofiber cross-sectional area.   Muscle fiber cross-sectional areas were not altered from intact values by OVX or estrogen replacement in control muscle (Fig. 2A). However, there was an interaction of estrogen treatment and recovery day on muscle cross-sectional area. Hindlimb disuse decreased fiber cross-sectional area similarly (40–55%) in all treatment groups (Fig. 2A). After 7 days of recovery, intact and ovariectomized cross-sectional areas remained decreased 42 and 44%, respectively, from control values. After 14 days of recovery, cross-sectional area from intact rats was restored to control values, whereas ovariectomized rats maintained a 42% deficit. Cross-sectional area did return to control values by the 28th day of recovery in ovariectomized females, demonstrating fiber growth after the second week of recovery.


Figure 2
View larger version (27K):
[in this window]
[in a new window]
  		Fig. 2. Soleus muscle myofiber growth alterations with suspension and reloading in ovariectomized female rats. A: soleus fiber mean cross-sectional area (CSA, µm2) in intact (black bars), ovariectomized (OVX, gray bars), and ovariectomized with 17beta-estradiol replacement (OVX+E2, hatched bars) control (Con), 10-day suspension (Sus), 7-day recovery (Sus+7), and 14 days of recovery (Sus+14) treatment groups. Transverse sections (10 µm) were cut from the midbelly of the soleus muscle on a cryostat at –20°C and mounted on coverslips. Hematoxylin-eosin (H and E) staining was performed, and 4 digital images were taken from each muscle section at a x20 magnification to determine the CSA of individual fibers. B: soleus muscle percentage small (<500 µm2) fibers. C: soleus muscle percentage large (>4,000 µm2) fibers. Values are presented as means ± SE. *Significantly (P < 0.05) different from treatment-matched Con. {dagger}Significantly (P < 0.05) different from treatment-matched Sus. {ddagger}Significantly (P < 0.05) different from treatment-matched Sus+7. #Significantly (P < 0.05) different from intact ground control.

 
During remodeling and growth, the distribution of myofiber cross-sectional areas within the muscle can provide important information beyond mean area alone. However, in control muscle, ovariectomy increased small (<500 µm2) fiber incidence 251%, and estrogen replacement attenuated this increase (Fig. 2B). With suspension, intact muscle small myofiber incidence increased 154% and was further elevated after 7 (508%) and 14 days (405%) of recovery. Muscle from ovariectomized rats did not demonstrate a change in small myofiber incidence with suspension or 7 days of recovery. However, after 14 days of recovery small fiber incidence increased 97%, returning to baseline after 28 days of recovery (9.2 ± 1.3%). In controls, large (>4,000 µm2) myofiber incidence was not altered from intact values by ovariectomy or estrogen replacement (Fig. 2C). Regardless of estrogen status, suspension ablated the incidence of large myofibers (Fig. 2C). In intact animals, large fiber incidence was returned to control values at day 14 of recovery, whereas large fiber incidence in ovariectomized muscle remained depressed after 14 days of recovery. After 28 days of recovery large myofiber incidence increased 101% above control values in ovariectomized female muscle.

p70s6kinase expression.   p70s6kinase protein phosphorylation is associated with signaling that increases translational efficiency (45), and estrogen status altered its expression and phosphorylation in control muscles. Ovariectomy reduced control muscle total p70s6k protein abundance by 36%, compared with intact animals (Fig. 3B). Estrogen replacement returned ovariectomized muscle total p70S6kinase abundance to control levels. Muscle from ovariectomized animals (2.1 ± 0.1 IOD) increased Thr389 phosphorylation 115% compared with intact (0.9 ± 0.2 IOD) animals. Estrogen replacement attenuated the ovariectomy-induced increase in phosphorylation (1.0 ± 0.1 IOD). There was an interaction between estrogen treatment and day of recovery on total p70s6k protein expression. Intact muscle did not alter total p70S6kinase expression after suspension, 7 days, or 14 days of recovery (Fig. 3B). However, in ovariectomized animals, total p70s6k decreased 19% with suspension and remained decreased 33% at day 7 of recovery. After 14 days of recovery, total p70s6k protein increased 43% compared with its control. With estrogen-replacement ovariectomized muscle total p70s6k protein abundance was not altered by suspension. Phosphorylation of p70s6k demonstrated no interaction between estrogen treatment and day of recovery from disuse.


Figure 3
View larger version (26K):
[in this window]
[in a new window]
  		Fig. 3. p70s6k protein expression in suspended and reloaded ovariectomized female rat soleus muscle. A: representative Western immunoblot of total p70s6kinase in OVX ground Con, Sus, Sus+7, and Sus+14 muscle. B: soleus total p70s6k protein expression in intact (black bars), OVX (gray bars), and OVX+E2 (hatched bars) Con, 10-day Sus, Sus+7, and Sus+14 treatment groups. Values are presented as means ± SE and normalized to intact female soleus ground control. *Significantly (P < 0.05) different from treatment-matched Con. {dagger}Significantly (P < 0.05) different from treatment-matched 10-day Sus. {ddagger}Significantly (P < 0.05) different from treatment-matched Sus+7. #Significantly (P < 0.05) different from intact ground control. {varepsilon}Significantly (P < 0.05) different from OVX+E2 ground control.

 
Myofiber regeneration.   Centrally localized nuclei (Fig. 4) and positive staining for eMHC (Fig. 5) are indicators of myofiber regeneration in skeletal muscle (17). There was an interaction of estrogen treatment and recovery day on the incidence of myofibers demonstrating central nuclei (Fig. 6A). Suspension decreased central nuclei incidence in intact (76%), ovariectomized (90%), and estrogen-replacement (72%) soleus muscle. Seven days of recovery increased the number of central nuclei containing fibers in intact animals 120%, which returned to control levels at day 14. However, in muscle from ovariectomized rats, central nuclei incidence returned to control values at day 7 of recovery but was then elevated 271% from control values at day 14 of recovery. Estrogen replacement reduced the incidence of myofibers with central nuclei at day 14 of recovery. There was a main effect of recovery day on the incidence of eMHC-positive myofibers. eMHC-positive fibers peaked after 7 days of recovery and remained elevated after 14 days of recovery (Fig. 6B). However, there was no estrogen treatment and recovery day interaction on the occurrence of eMHC positively stained fibers.


Figure 4
View larger version (135K):
[in this window]
[in a new window]
  		Fig. 4. Representative soleus muscle H and E staining of intact ground control (A), intact with 10 days of hindlimb suspension and 14 days of reloading (B), OVX ground control (C), and OVX with 10 days of hindlimb suspension and 14 days of reloading (D).

 

Figure 5
View larger version (160K):
[in this window]
[in a new window]
  		Fig. 5. Representative soleus muscle embryonic myosin heavy chain immunohistochemical staining of negative control (no primary antibody, A), intact ground control (B), intact with 10 days of hindlimb suspension and 14 days of reloading (C), ovariectomized with 10 days of hindlimb suspension and 14 days of reloading (D), ovariectomized with estradiol replacement all days and 10 days of hindlimb suspension and 14 days of reloading (E), and ovariectomized with estradiol replacement reload days only and 10 days of hindlimb suspension and 14 days of reloading (F).

 

Figure 6
View larger version (14K):
[in this window]
[in a new window]
  		Fig. 6. Soleus muscle fiber regeneration with suspension and reloading in ovariectomized female rats. A: occurrence of centralized nuclei in intact (black bars), OVX (gray bars), and OVX+E2 (hatched bars) Con, Sus, Sus+7, and Sus+14 treatment groups. Transverse sections (10 µm) were cut from the midbelly of the soleus muscle on a cryostat at –20°C and mounted on coverslips. H and E staining was performed to determine the percentage of fibers with central nuclei. B: soleus fiber embryonic myosin heavy chain (eMHC) expression. Transverse sections (10 µm) were cut from the midbelly of the soleus muscle on a cryostat at –20°C and mounted on coverslips. Immunohistochemistry staining was performed for eMHC and 4 digital images were taken from each muscle section at a x20 magnification to determine myofiber expression per mm2 of muscle area. There was a main effect of recovery day on eMHC expression per mm2 of soleus muscle area. Values are presented as means ± SE. *Significantly (P < 0.05) different from treatment-matched Con. {dagger}Significantly (P < 0.05) different from treatment-matched Sus. {ddagger}Significantly (P < 0.05) different from treatment-matched Sus+7.

 
Noncontractile tissue.   The composition and volume of the extracellular matrix is thought to contribute to the regulation of skeletal muscle remodeling processes (15, 16). Extracellular matrix expansion can influence muscle mechanical properties and is important in the pathology of fibrosis (17, 40). There was an interaction of estrogen treatment and recovery day on the volume %NCT. Suspension increased soleus muscle %NCT in intact (53%), ovariectomized (74%), and estrogen-replacement (50%) rats, compared with their appropriate control (Table 1). In both intact and ovariectomized animals, the %NCT remained increased 38 and 53%, respectively, from control values at day 7 of recovery. In intact females, %NCT returned to control by day 14 of recovery, whereas ovariectomized rats increased %NCT 29% from control values. Ovariectomized female muscle %NCT returned to control values by day 28 (21 ± 1.3%) of recovery.

Collagen I is highly expressed in postural skeletal muscle and mediates the muscle's resistance to stretch (14). In control muscle, estrogen administration decreased collagen 1a abundance 60 and 76% from intact and ovariectomized values, respectively (Fig. 7). Intact muscle collagen 1a mRNA abundance was not altered by either hindlimb suspension or 7 days of recovery but was decreased 70% at day 14 of recovery. In ovariectomized muscle, suspension decreased collagen 1a mRNA abundance 62% from control and returned to control levels after 7 days of recovery. Collagen 1a mRNA abundance was increased 46% above control values in ovariectomized muscle after 14 days of recovery. In ovariectomized animals administered estrogen, suspension had no effect on collagen 1a mRNA abundance. Interestingly, recovery for 14 days with estrogen administration resulted in increased collagen 1a mRNA abundance compared with treatment control values.


Figure 7
View larger version (21K):
[in this window]
[in a new window]
  		Fig. 7. Alterations in collagen mRNA with suspension and reloading in ovariectomized female rats. Ten to 15 µg of total RNA were fractionated on a denaturing 1% agarose gels (1 x 3 MOPS, 6.7% formaldehyde) and then transferred to a nylon membrane by capillary action. Membranes were visualized by autoradiography (–80°C, 3–40 h) and quantified by densitometry scanning. A: representative Northern blot of collagen 1a mRNA abundance and 18S and 28S mRNA in ovariectomized Con and Sus+14 treatments. B: collagen 1a mRNA expression in intact, OVX, and OVX+E2 Con, 10-day Sus, Sus+7, and Sus+14 treatment groups. Values are presented as means ± SE and normalized to intact female soleus ground control. *Significantly (P < 0.05) different from treatment-matched Con. {dagger}Significantly (P < 0.05) different from treatment-matched 10-day Sus. {ddagger}Significantly (P < 0.05) different from treatment-matched Sus+7. #Significantly (P < 0.05) different from intact ground control. {varepsilon}Significantly (P < 0.05) different from OVX+E2 ground control.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Estrogen loss has been shown to alter skeletal muscle wet weight and protein content during recovery from disuse atrophy (57). A variety of processes within skeletal muscle contribute to disuse mass recovery including edema, myofiber regeneration and growth, and extracellular matrix remodeling. The results of the present study demonstrate that estrogen is an important regulator of soleus muscle fiber growth and extracellular matrix remodeling during recovery from disuse. Loss of ovarian hormones impaired the recovery of soleus skeletal muscle fiber cross-sectional area during the first 2 wk of reload. However, estrogen replacement at physiologically relevant levels restored the ability of muscle fibers to recover cross-sectional area over this same 2-wk period. Another important finding was that the beneficial effect of estrogen replacement on muscle recovery was independent of when the hormone was administered, before disuse or at the time of recovery. Therefore, to our knowledge, this paper demonstrates for the first time that estrogen has a direct effect on the recovery of soleus muscle myofiber size after atrophy. Although ovarian hormone loss dramatically affected myofiber growth during the first 2 wk of recovery, myofiber cross-sectional area was able to fully recover after 28 days, regardless of hormone status. This finding extends previous work with ovarian hormone loss and muscle mass recovery and demonstrates that the soleus muscle can still regain functional muscle mass without ovarian hormones, but the process is significantly delayed.

In the present study, ovarian hormone status induced alterations in soleus muscle wet weight. Fluctuations in muscle wet weight can be accounted for by a variety of muscle components and are not totally dependent on myofiber size (35). This study and others clearly demonstrate that ovariectomy is a potent stimulus for increased rat soleus muscle mass that is independent of increased myofiber cross-sectional area (23, 24). Interestingly, despite this basal increase in soleus muscle mass with ovariectomy, soleus myofibers from these animals had an altered capacity for plasticity related to loading stimuli. The ovariectomized rats in the present study demonstrated a nearly complete restoration of muscle mass during 2 wk of recovery from disuse without a corresponding increase in myofiber area. These findings are in agreement with previous work (13). Initial changes in skeletal muscle mass during disuse recovery are attributed primarily to edema and characterized by deficits in both force production and mean cross-sectional area (11, 31, 45). The time course of gene expression induced early during reloading is a highly coordinated process related to muscle damage and repair, oxidative metabolism, and muscle regeneration (26). In the present study, the time course of skeletal muscle growth after disuse followed a sequential pattern that first involved edema and then was followed by increased dry weight, myofibrillar protein accretion, and myofiber cross-sectional area restoration. Estrogen has the potential to regulate several important processes related to this sequence of muscle recovery from atrophy, and the present study demonstrates a direct effect of estrogen on myofibers. Specifically, estrogen administration rescued soleus mean fiber cross-sectional area and large myofiber incidence in ovariectomized animals during 2 wk of recovery.

Sitnick et al. (57) recently demonstrated that ovariectomy attenuates the recovery of rat gastrocnemius muscle mass during a 14-day recovery period from 28 days of hindlimb suspension and established a role for ovarian hormones in the activation of Akt and p70s6kinase (57). These proteins are critical to the regulation of protein synthesis capacity and/or efficiency (5). The present study also found alterations in p70s6kinase phosphorylation and total protein expression with ovariectomy during a shorter period of hindlimb suspension and during recovery in the postural soleus muscle. We have now expanded on these findings to confirm that the loss of the specific ovarian hormone estrogen is sufficient to induce these alterations. Coinciding with p70s6kinase phosphorylation and total expression, estrogen replacement also rescued ovariectomy-induced alterations in myofiber growth and small myofiber occurrence. Increased p70s6kinase protein expression at day 14 of recovery in ovariectomized animals provides further evidence for protein accretion associated with myofiber growth at later time points of recovery. Soleus myofiber cross-sectional area in ovariectomized rats returned to control levels between the 2nd and 4th weeks of recovery. Myofiber growth is complex and involves many processes related to the myofiber's microenvironment. The delay in fiber growth with ovariectomy may demonstrate the importance of the resolution of early processes related to damage and inflammation so that subsequent myofiber growth can occur. Candidates for estrogen-sensitive manipulation of the fiber environment during muscle recovery from disuse include extracellular matrix remodeling, satellite cell regulation, and inflammatory processes (39).

The extracellular matrix has a critical role in muscle function and homeostatic regulation that is related to both its protein composition and cellularity (36). Skeletal muscle extracellular matrix remodeling can be modulated by mechanical stress, growth factor, hormone, and cytokine signaling (44, 53). The present study demonstrates both disuse and the subsequent recovery dramatically affect extracellular matrix volume. However, the composition, as it relates to collagen 1a mRNA expression, was not sensitive to disuse and recovery. Although not affecting extracellular matrix volume, ovarian hormone status did alter muscle collagen 1a expression. Ovariectomy can induce rat hepatic extracellular matrix expansion, which is characterized by increases in types I and III procollagen mRNA abundance (62). However, in the present study, the regulation of muscle collagen 1a expression was not estrogen sensitive and may be regulated by other ovarian hormones. Ovariectomy may alter extracellular matrix volume indirectly through other hormones or growth factors. IGF-I overabundance is thought to contribute to fibrosis in chronically strained skeletal muscle (20), and estrogen depletion due to ovariectomy increases circulating growth hormone and IGF-I (12, 32).

Muscle loading and ovarian hormone loss had an additive effect on volume of noncontractile tissue and collagen 1a mRNA abundance, suggesting that a hormone and mechanical signaling interaction may underlie extracellular matrix remodeling. Endomysial fibroblasts are the primary source of extracellular matrix protein synthesis (25), and they express estrogen receptors (30). TGF-beta is a profibrotic cytokine, and intracellular signaling related to this pathway has been shown to interact with the estrogen receptor. Specifically, estrogen receptors can physically bind and inhibit the action of Sma and MAD-related protein 3 (Smad-3), a downstream component of TGF-beta signaling (43). Skeletal muscle recovery from either laceration or cardiotoxin injury is inhibited by TGF-beta-induced expression of profibrotic proteins (40). Estrogen effects on TGF-beta signaling may also have implications for satellite cell activity. In myoblasts, Smad-3 functions in TGF-beta-mediated suppression of myoblast differentiation through interaction with myogenic regulatory factor MyoD (41).

There is the possibility that during disuse ovariectomized muscle had a greater depletion of both the myogenic precursor pool and myofiber nuclei, because both of these pools are decreased in atrophying skeletal muscle (49). However, the present study demonstrates that estrogen effects on muscle recovery are more likely related to events during the recovery process rather than during the period of disuse. Estrogen replacement in ovariectomized animals at the time of reloading was as effective for rescuing muscle mass and myofiber size by day 14 of recovery as when estrogen was supplied during both the atrophy and recovery phases. This finding suggests that estrogen's contribution for successful muscle recovery may be related to myogenic activation upon reloading rather than myoblast protection during atrophy. A retarded ability of myogenic precursor cells to activate and differentiate occurs after prolonged atrophic conditions (50). Disuse combined with estrogen loss appears to require longer recovery periods, which would allow precursor cells to reach a critical population necessary for both myofiber growth and successful regeneration. Ovariectomy appears to delay the local expression of cellular markers responsible for differentiation in this population of cells.

The resolution of inflammation and edema may be critical in signaling associated with myofiber growth. Bondesen et al. (9) suggest that the attenuation of inflammation may be critical for myofiber growth owing to its regulation of immune cell phagocytosis of necrotic tissue. When the muscle's inflammatory response is manipulated, recovery from muscle damage is delayed, but not ablated (9). Prolonged or exaggerated edema and inflammation due to ovarian hormone loss could delay regeneration, and ultimately myofiber growth. In the present study, ovariectomy prolonged the demonstration of histological markers for regeneration during reloading. Small myofiber incidence, a morphological marker of ongoing muscle regeneration (17), was induced after 7 days of recovery in the intact female, and ovarian hormone loss suppressed this induction. There is a strong possibility that the estrogen-sensitive mechanisms delaying muscle regeneration may also be attenuating myofiber growth. Estrogen accelerates cutaneous wound healing (3), and hormone replacement therapy prevents chronic wound development in postmenopausal women (7). Ashcroft et al. (4) suggested that estrogen receptor regulation of macrophage inhibitory factor is responsible for the suppression of excessive proteolysis during cutaneous wound healing. A key characteristic of estrogen's action on wound healing appears to be the suppression of extracellular matrix component deposition at the site of injury (3, 43). Immediate early gene expression in muscle can be regulated by growth factor, cytokine, and estrogen receptor signaling interactions (22). Estrogen signaling has the potential to interact with both growth factor and inflammatory cytokine signaling pathways. Estradiol, IGF-I, and the inflammatory cytokine TNF-{alpha} are independently capable of activating serum response element promoter regions on reporter genes in MCF-7 cells, but the addition of constitutively active estrogen receptor-{alpha} markedly enhances this activation. Because inflammatory signaling is critical for the response of muscle precursor cells during regeneration (9), interactions between these pathways could amplify the importance of estradiol availability during regeneration. Further work is needed to determine whether potential alterations in the inflammatory response at the onset of reloading after disuse are sensitive to estrogen.

In summary, successful muscle recovery from disuse atrophy requires the integration of many signaling pathways that serve to first initiate and then sustain muscle regeneration and growth. The present study demonstrates that ovarian hormones can have a significant impact on muscle fiber size and extracellular matrix remodeling during reloading after disuse. Estrogen may aid muscle recovery by suppressing extracellular matrix remodeling. Determining the importance of estrogen regulation of muscle growth and remodeling processes during reloading after atrophy could provide novel therapeutic strategies targeted at the maintenance and recovery of muscle mass.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
ACKNOWLEDGMENTS
 REFERENCES
 
This work supported by National Science Foundation/Experimental Program to Stimulate Competitive Research grant no. EPS-0132573 and National Institutes of Health/Biomedical Research Infrastructure Network (BRIN) grant no. 8-PORR13461A, as well as American College of Sports Medicine/National Aeronautics and Space Administration Student grants awarded to Joseph M. McClung for the 2002–2004 funding periods.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
REFERENCES
 
The authors thank Kristen A. Mehl and Tyrone A. Washington for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. A. Carson, Univ. of South Carolina, Dept. of Exercise Science, 1300 Wheat St., Columbia, SC 29208 (e-mail: carsonj{at}gwm.sc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 

  1. Alt N, Carson JA, Alderson NL, Wang Y, Nagai R, Henle T, Thorpe SR, and Baynes JW. Chemical modification of muscle protein in diabetes. Arch Biochem Biophys 425: 200–206, 2004.[CrossRef][Web of Science][Medline]
  2. Alway SE, Winchester PK, Davis ME, and Gonyea WJ. Regionalized adaptations and muscle fiber proliferation in stretch-induced enlargement. J Appl Physiol 66: 771–781, 1989.[Abstract/Free Full Text]
  3. Ashcroft GS, Greenwell-Wild T, Horan MA, Wahl SM, and Ferguson MW. Topical estrogen accelerates cutaneous wound healing in aged humans associated with an altered inflammatory response. Am J Pathol 155: 1137–1146, 1999.[Abstract/Free Full Text]
  4. Ashcroft GS, Mills SJ, Lei K, Gibbons L, Jeong MJ, Taniguchi M, Burow M, Horan MA, Wahl SM, and Nakayama T. Estrogen modulates cutaneous wound healing by downregulating macrophage migration inhibitory factor. J Clin Invest 111: 1309–1318, 2003.[CrossRef][Web of Science][Medline]
  5. Baar K and Esser K. Phosphorylation of p70(S6k) correlates with increased skeletal muscle mass following resistance exercise. Am J Physiol Cell Physiol 276: C120–C127, 1999.[Abstract/Free Full Text]
  6. Baldwin KM and Haddad F. Skeletal muscle plasticity: cellular and molecular responses to altered physical activity paradigms. Am J Phys Med Rehabil 81: S40–S51, 2002.[CrossRef][Web of Science][Medline]
  7. Berard A, Kahn SR, and Abenhaim L. Is hormone replacement therapy protective for venous ulcer of the lower limbs? Pharmacoepidemiol Drug Saf 10: 245–251, 2001.[CrossRef][Web of Science][Medline]
  8. Bigard XA, Merino D, Serrurier B, Lienhard F, Guezennec YC, Bockhold KJ, and Whalen RG. Role of weight-bearing function on expression of myosin isoforms during regeneration of rat soleus muscles. Am J Physiol Cell Physiol 270: C763–C771, 1996.[Abstract/Free Full Text]
  9. Bondesen BA, Mills ST, Kegley KM, and Pavlath GK. The COX-2 pathway is essential during early stages of skeletal muscle regeneration. Am J Physiol Cell Physiol 287: C475–C483, 2004.[Abstract/Free Full Text]
  10. Booth FW and Seider MJ. Early change in skeletal muscle protein synthesis after limb immobilization of rats. J Appl Physiol 47: 974–977, 1979.[Abstract/Free Full Text]
  11. Booth FW and Seider MJ. Recovery of skeletal muscle after 3 mo of hindlimb immobilization in rats. J Appl Physiol 47: 435–439, 1979.[Abstract/Free Full Text]
  12. Borski RJ, Tsai W, DeMott-Friberg R, and Barkan AL. Regulation of somatic growth and the somatotropic axis by gonadal steroids: primary effect on insulin-like growth factor I gene expression and secretion. Endocrinology 137: 3253–3259, 1996.[Abstract]
  13. Brown M, Foley A, and Ferreria JA. Ovariectomy, hindlimb unweighting, and recovery effects on skeletal muscle in adult rats. Aviat Space Environ Med 76: 1012–1018, 2005.[Medline]
  14. Burgeson RE. The collagens of skin. Curr Probl Dermatol 17: 61–75, 1987.[Medline]
  15. Chan YS, Li Y, Foster W, Fu FH, and Huard J. The use of suramin, an antifibrotic agent, to improve muscle recovery after strain injury. Am J Sports Med 33: 43–51, 2005.[Abstract/Free Full Text]
  16. Chan YS, Li Y, Foster W, Horaguchi T, Somogyi G, Fu FH, and Huard J. Antifibrotic effects of suramin in injured skeletal muscle after laceration. J Appl Physiol 95: 771–780, 2003.[Abstract/Free Full Text]
  17. Charge SB and Rudnicki MA. Cellular and molecular regulation of muscle regeneration. Physiol Rev 84: 209–238, 2004.[Abstract/Free Full Text]
  18. Chiquet M. Regulation of extracellular matrix gene expression by mechanical stress. Matrix Biol 18: 417–426, 1999.[CrossRef][Web of Science][Medline]
  19. Darr KC and Schultz E. Hindlimb suspension suppresses muscle growth and satellite cell proliferation. J Appl Physiol 67: 1827–1834, 1989.[Abstract/Free Full Text]
  20. De Deyne PG, Kinsey S, Yoshino S, and Jensen-Vick K. The adaptation of soleus and edl in a rat model of distraction osteogenesis: IGF-1 and fibrosis. J Orthop Res 20: 1225–1231, 2002.[CrossRef][Web of Science][Medline]
  21. Diffee GM, Caiozzo VJ, Herrick RE, and Baldwin KM. Contractile and biochemical properties of rat soleus and plantaris after hindlimb suspension. Am J Physiol Cell Physiol 260: C528–C534, 1991.[Abstract/Free Full Text]
  22. Duan R, Porter W, and Safe S. Estrogen-induced c-fos protooncogene expression in MCF-7 human breast cancer cells: role of estrogen receptor Sp1 complex formation. Endocrinology 139: 1981–1990, 1998.[Abstract/Free Full Text]
  23. Fisher JS, Hasser EM, and Brown M. Effects of ovariectomy and hindlimb unloading on skeletal muscle. J Appl Physiol 85: 1316–1321, 1998.[Abstract/Free Full Text]
  24. Fisher JS, Kohrt WM, and Brown M. Food restriction suppresses muscle growth and augments osteopenia in ovariectomized rats. J Appl Physiol 88: 265–271, 2000.[Abstract/Free Full Text]
  25. Fluck M, Chiquet M, Schmutz S, Mayet-Sornay MH, and Desplanches D. Reloading of atrophied rat soleus muscle induces tenascin-C expression around damaged muscle fibers. Am J Physiol Regul Integr Comp Physiol 284: R792–R801, 2003.[Abstract/Free Full Text]
  26. Fluck M, Schmutz S, Wittwer M, Hoppeler H, and Desplanches D. Transcriptional reprogramming during reloading of atrophied rat soleus muscle. Am J Physiol Regul Integr Comp Physiol 289: R4–R14, 2005.[Abstract/Free Full Text]
  27. Frenette J, Cai B, and Tidball JG. Complement activation promotes muscle inflammation during modified muscle use. Am J Pathol 156: 2103–2110, 2000.[Abstract/Free Full Text]
  28. Frenette J, Chbinou N, Godbout C, Marsolais D, and Frenette PS. Macrophages, not neutrophils, infiltrate skeletal muscle in mice deficient in P/E selectins after mechanical reloading. Am J Physiol Regul Integr Comp Physiol 285: R727–R732, 2003.[Abstract/Free Full Text]
  29. Frenette J, St-Pierre M, Cote CH, Mylona E, and Pizza FX. Muscle impairment occurs rapidly and precedes inflammatory cell accumulation after mechanical loading. Am J Physiol Regul Integr Comp Physiol 282: R351–R357, 2002.[Abstract/Free Full Text]
  30. Grohe C, Kahlert S, Lobbert K, Stimpel M, Karas RH, Vetter H, and Neyses L. Cardiac myocytes and fibroblasts contain functional estrogen receptors. FEBS Lett 416: 107–112, 1997.[CrossRef][Web of Science][Medline]
  31. Itai Y, Kariya Y, and Hoshino Y. Morphological changes in rat hindlimb muscle fibres during recovery from disuse atrophy. Acta Physiol Scand 181: 217–224, 2004.[CrossRef][Web of Science][Medline]
  32. Jansson JO, Ekberg S, Isaksson OG, and Eden S. Influence of gonadal steroids on age- and sex-related secretory patterns of growth hormone in the rat. Endocrinology 114: 1287–1294, 1984.[Abstract/Free Full Text]
  33. Kahlert S, Grohe C, Karas RH, Lobbert K, Neyses L, and Vetter H. Effects of estrogen on skeletal myoblast growth. Biochem Biophys Res Commun 232: 373–378, 1997.[CrossRef][Web of Science][Medline]
  34. Kawano S, Kanda K, Ohmori S, Izumi R, Yasukawa K, Murata Y, and Seo H. Effect of estrogen on the development of disuse atrophy of bone and muscle induced by tail-suspension in rats. Environ Med 41: 89–92, 1997.[Medline]
  35. Kelso TB, Shear CR, and Max SR. Enzymes of glutamine metabolism in inflammation associated with skeletal muscle hypertrophy. Am J Physiol Endocrinol Metab 257: E885–E894, 1989.[Abstract/Free Full Text]
  36. Kjaer M. Role of extracellular matrix in adaptation of tendon and skeletal muscle to mechanical loading. Physiol Rev 84: 649–698, 2004.[Abstract/Free Full Text]
  37. Kobori M and Yamamuro T. Effects of gonadectomy and estrogen administration on rat skeletal muscle. Clin Orthop: 306–311, 1989.
  38. Leask A and Abraham DJ. TGF-beta signaling and the fibrotic response. FASEB J 18: 816–827, 2004.[Abstract/Free Full Text]
  39. Levin ER. Cell localization, physiology, and nongenomic actions of estrogen receptors. J Appl Physiol 91: 1860–1867, 2001.[Abstract/Free Full Text]
  40. Li Y, Foster W, Deasy BM, Chan Y, Prisk V, Tang Y, Cummins J, and Huard J. Transforming growth factor-beta1 induces the differentiation of myogenic cells into fibrotic cells in injured skeletal muscle: a key event in muscle fibrogenesis. Am J Pathol 164: 1007–1019, 2004.[Abstract/Free Full Text]
  41. Liu D, Black BL, and Derynck R. TGF-beta inhibits muscle differentiation through functional repression of myogenic transcription factors by Smad3. Genes Dev 15: 2950–2966, 2001.[Abstract/Free Full Text]
  42. Lynch GS. Therapies for improving muscle function in neuromuscular disorders. Exerc Sport Sci Rev 29: 141–148, 2001.[CrossRef][Medline]
  43. Matsuda T, Yamamoto T, Muraguchi A, and Saatcioglu F. Cross-talk between transforming growth factor-beta and estrogen receptor signaling through Smad3. J Biol Chem 276: 42908–42914, 2001.[Abstract/Free Full Text]
  44. Mayne R and Sanderson RD. The extracellular matrix of skeletal muscle. Coll Relat Res 5: 449–468, 1985.[Web of Science][Medline]
  45. McClung J, Thompson RW, Lowe LL, and Carson JA. RhoA expression during recovery from skeletal muscle disuse. J Appl Physiol 96: 1341–1348, 2004.[Abstract/Free Full Text]
  46. McClung JM, Lee WJ, Thompson RW, Lowe LL, and Carson JA. RhoA induction by functional overload and nandrolone decanoate administration in rat skeletal muscle. Pflügers Arch 447: 1–8, 2003.[CrossRef][Web of Science][Medline]
  47. McClung JM, Mehl KA, Thompson RW, Lowe LL, and Carson JA. Nandrolone decanoate modulates cell cycle regulation in functionally overloaded rat soleus muscle. Am J Physiol Regul Integr Comp Physiol 288: R1543–R1552, 2005.[Abstract/Free Full Text]
  48. Miller TA, Lesniewski LA, Muller-Delp JM, Majors AK, Scalise D, and Delp MD. Hindlimb unloading induces a collagen isoform shift in the soleus muscle of the rat. Am J Physiol Regul Integr Comp Physiol 281: R1710–R1717, 2001.[Abstract/Free Full Text]
  49. Mitchell PO and Pavlath GK. A muscle precursor cell-dependent pathway contributes to muscle growth after atrophy. Am J Physiol Cell Physiol 281: C1706–C1715, 2001.[Abstract/Free Full Text]
  50. Mitchell PO and Pavlath GK. Skeletal muscle atrophy leads to loss and dysfunction of muscle precursor cells. Am J Physiol Cell Physiol 287: C1753–C1762, 2004.[Abstract/Free Full Text]
  51. Ohira M, Hanada H, Kawano F, Ishihara A, Nonaka I, and Ohira Y. Regulation of the properties of rat hind limb muscles following gravitational unloading. Jpn J Physiol 52: 235–245, 2002.[CrossRef][Web of Science][Medline]
  52. Paroo Z, Dipchand ES, and Noble EG. Estrogen attenuates postexercise HSP70 expression in skeletal muscle. Am J Physiol Cell Physiol 282: C245–C251, 2002.[Abstract/Free Full Text]
  53. Rettig WJ, Erickson HP, Albino AP, and Garin-Chesa P. Induction of human tenascin (neuronectin) by growth factors and cytokines: cell type-specific signals and signalling pathways. J Cell Sci 107: 487–497, 1994.[Abstract]
  54. Richard D. Effects of ovarian hormones on energy balance and brown adipose tissue thermogenesis. Am J Physiol Regul Integr Comp Physiol 250: R245–R249, 1986.[Abstract/Free Full Text]
  55. Saartok T. Steroid receptors in two types of rabbit skeletal muscle. Int J Sports Med 5: 130–136, 1984.[Web of Science][Medline]
  56. Sato K, Li Y, Foster W, Fukushima K, Badlani N, Adachi N, Usas A, Fu FH, and Huard J. Improvement of muscle healing through enhancement of muscle regeneration and prevention of fibrosis. Muscle Nerve 28: 365–372, 2003.[CrossRef][Web of Science][Medline]
  57. Sitnick M, Foley AM, Brown M, and Spangenburg EE. Ovariectomy prevents the recovery of atrophied gastrocnemius skeletal muscle mass. J Appl Physiol 100: 286–293, 2006.[Abstract/Free Full Text]
  58. Suzuki S and Yamamuro T. Long-term effects of estrogen on rat skeletal muscle. Exp Neurol 87: 291–299, 1985.[CrossRef][Web of Science][Medline]
  59. Tapper CM, Greig F, and Brown-Grant K. Effects of steroid hormones on gonadotrophin secretion in female rats after ovariectomy during the oestrous cycle. J Endocrinol 62: 511–525, 1974.[Abstract/Free Full Text]
  60. Trenkle A. The anabolic effect of estrogens on nitrogen metabolism of growing and finishing cattle and sheep. Environ Qual Saf Suppl 5: 79–88, 1976.[Medline]
  61. Tsika RW, Herrick RE, and Baldwin KM. Time course adaptations in rat skeletal muscle isomyosins during compensatory growth and regression. J Appl Physiol 63: 2111–2121, 1987.[Abstract/Free Full Text]
  62. Yasuda M, Shimizu I, Shiba M, and Ito S. Suppressive effects of estradiol on dimethylnitrosamine-induced fibrosis of the liver in rats. Hepatology 29: 719–727, 1999.[CrossRef][Web of Science][Medline]
  63. Yeh JK, Chen MM, and Aloia JF. Effects of estrogen and growth hormone on skeleton in the ovariectomized rat with hypophysectomy. Am J Physiol Endocrinol Metab 273: E734–E742, 1997.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 J EndocrinolHome page
M. Komrakova, C. Werner, M. Wicke, B. T. Nguyen, S. Sehmisch, M. Tezval, K. M. Stuermer, and E. K. Stuermer
Effect of daidzein, 4-methylbenzylidene camphor or estrogen on gastrocnemius muscle of osteoporotic rats undergoing tibia healing period
J. Endocrinol., May 1, 2009; 201(2): 253 - 262.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Endocrinol. Metab.Home page
M. Brown, J. Ning, J. A. Ferreira, J. L. Bogener, and D. B. Lubahn
Estrogen receptor-{alpha} and -{beta} and aromatase knockout effects on lower limb muscle mass and contractile function in female mice
Am J Physiol Endocrinol Metab, April 1, 2009; 296(4): E854 - E861.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
P. M. Tiidus and D. L. Enns
Point:Counterpoint: Estrogen and sex do/do not influence post-exercise indexes of muscle damage, inflammation, and repair
J Appl Physiol, March 1, 2009; 106(3): 1010 - 1012.
[Full Text] [PDF]

Home page
 Am. J. Physiol. Regul. Integr. Comp. Physiol.Home page
J. J. Widrick, G. F. Maddalozzo, H. Hu, J. C. Herron, U. T. Iwaniec, and R. T. Turner
Detrimental effects of reloading recovery on force, shortening velocity, and power of soleus muscles from hindlimb-unloaded rats
Am J Physiol Regulatory Integrative Comp Physiol, November 1, 2008; 295(5): R1585 - R1592.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
B. T. Corona, C. Rouviere, S. L. Hamilton, and C. P. Ingalls
FKBP12 deficiency reduces strength deficits after eccentric contraction-induced muscle injury
J Appl Physiol, August 1, 2008; 105(2): 527 - 537.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Regul. Integr. Comp. Physiol.Home page
J. M. McClung, M. A. Whidden, A. N. Kavazis, D. J. Falk, K. C. DeRuisseau, and S. K. Powers
Redox regulation of diaphragm proteolysis during mechanical ventilation
Am J Physiol Regulatory Integrative Comp Physiol, May 1, 2008; 294(5): R1608 - R1617.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
A. L. Moran, S. A. Nelson, R. M. Landisch, G. L. Warren, and D. A. Lowe
Estradiol replacement reverses ovariectomy-induced muscle contractile and myosin dysfunction in mature female mice
J Appl Physiol, April 1, 2007; 102(4): 1387 - 1393.
[Abstract] [Full Text] [PDF]

Home page
 Exp PhysiolHome page
J. M. McClung, J. M. Davis, and J. A. Carson
Muscle: Ovarian hormone status and skeletal muscle inflammation during recovery from disuse in rats
Exp Physiol, January 1, 2007; 92(1): 219 - 232.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
100/6/2012    most recent
01583.2005v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (4)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by McClung, J. M.
Right arrow Articles by Carson, J. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by McClung, J. M.
Right arrow Articles by Carson, J. A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2006 by the American Physiological Society.