| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
1 Department of Exercise and Nutrition Sciences, University at Buffalo, State University of New York, Buffalo, NY 14214, USA Departments of 2 Pharmacy Practice and Pharmaceutical Sciences, University at Buffalo, State University of New York, Amherst, NY 14260, USA
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
(Received 28 June 2004;
accepted after revision 21 September 2004; first published online 4 October 2004)
Corresponding author K. M. McCormick: Department of Exercise and Nutrition Sciences, University at Buffalo, State University at New York, Buffalo, NY 14214, USA. Email: kmccorm{at}buffalo.edu
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
A limited number of studies have examined the effect of oestrogen replacement on myofibre size and muscle mass in growing animals. Treatment of gonadectomized or sexually immature rats, mice or rabbits with oestrogen reduces muscle mass and fibre size (Ihemelandu, 1980, 1981; Kobori & Yamamuro, 1989; McCormick et al. 2004). Consistent with these findings, muscle protein synthesis and whole body protein gain are also reduced with oestrogen replacement (Denis et al. 1987; Toth et al. 2001). In mature rats, oestrogen replacement had no effect on muscle mass (Fisher et al. 2000). These results suggest that the effect of oestrogen replacement on muscle mass and fibre size may be restricted to immature animals.
The effect of oestrogen on MHC expression has also been examined. Oestrogen clearly alters MHC expression in the heart, but its effects on skeletal muscle are not as clear. Ovariectomy causes a fast to slow shift in cardiac MHC expression that is reversed with oestrogen replacement (Schaible et al. 1984; Scheuer et al. 1987). Similar to the heart, a previous study reported that ovariectomy caused a fast to slow shift in MHC expression in the soleus muscle, a muscle that is composed almost entirely of slow type I fibres (Kadi et al. 2002). The combination of oestrogen replacement and wheel running reversed the effect of ovariectomy, as did wheel running alone, making it difficult to conclude that oestrogen was responsible for changes in MHC expression. Previous work from this laboratory did not find changes in soleus muscle MHC composition with ovariectomy or with oestrogen replacement, suggesting that oestrogen may not alter MHC expression in the slow soleus muscle (McCormick et al. 2004). However, in fast extensor digitorum longus muscle (7% type IIa, 46% type IIx and 47% type IIb), ovariectomy caused a shift from the fastest isoform (IIb) to the slowest isoform (IIa; Kadi et al. 2002). Wheel running alone did not reverse the effect of ovariectomy, but when wheel running was combined with oestrogen replacement in ovariectomized animals, MHC isoform composition was similar to that of control animals, suggesting that oestrogen mediated the changes in MHC expression. These results suggest that the effect of oestrogen on MHC expression may depend on the fibre type composition of the muscle. Further work is needed to clarify this issue.
Regardless of whether oestrogen acts on skeletal muscle through direct and/or indirect mechanisms, scattered evidence suggests that its actions may depend on the maturity and the fibre type composition of the muscle. The purpose of this study was to add to this body of evidence by examining the effect of oestrogen deprivation and replacement on fibre size and MHC expression in the fast plantaris muscle.
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Twenty-five female Sprague–Dawley rats (7 weeks old) were separated into weight-matched groups. We have previously reported on the soleus muscle from these animals (McCormick et al. 2004). All procedures involving animals were approved by the Animal Care and Use Committee at the University at Buffalo in accordance with NIH guidelines.
All animals underwent bilateral ovariectomy or sham operation. Prior to surgery, animals were anaesthetized (75 mg kg–1 ketamine, 10 mg kg–1 xylazine, I.P.) and the skin was prepared for aseptic surgery. A ventral mid-line incision was made and the ovaries were isolated, but not tied off (Sham group), or tied off and removed (OVX group). Ovariectomized animals also received an implant containing either 17ß-oestradiol diluted (1:1) with corn oil (OVX/E2 group) or corn oil alone (OVX/CO group). Immediately after ovariectomy, the animals were placed in dorsal recumbency, the skin prepared, and a small medial incision immediately distal to the scapula was made. A haemostat was used to create a pocket between the skin and muscle layers and a 5 mm long Silastic tube containing oestrogen or corn oil was placed in the pocket. The skin was closed and animals were monitored until they regained consciousness. Two weeks after the initial surgery, animals were anaesthetized as before and the Silastic tube was removed, cleaned and repositioned. This second surgery was necessary to eliminate fatty tissue build-up around the tubing and ensure continued oestrogen delivery.
Groups were pair-fed to prevent the increase in body weight caused by ovariectomy (Davidge et al. 2001). The Sham group was allowed to eat ad libitum and their food intake measured. The amount of food given to the Sham group was given to the OVX/CO and OVX/E2 groups.
Following the 28 day treatment period, animals were killed by exsanguination following anaesthesia (75 mg kg–1 ketamine, 10 mg kg–1 xylazine, I.P.). The uterus was removed and weighed to confirm the effectiveness of the ovariectomy and oestrogen replacement. The plantaris muscles were removed from both limbs. Muscles from the left leg were immediately weighed and frozen in liquid nitrogen for analysis of MHC isoform composition. Muscles from the right leg were mounted for transverse sections and frozen in liquid nitrogen-cooled isopentane. Samples were stored at –70°C until analysis.
Myosin heavy chain composition
MHC isoform composition of the plantaris muscle was determined using sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE). Myofibrils were prepared as previously described (McCormick et al. 2004). Briefly, the muscle was homogenized in rigor buffer (0.1 M NaCl, 10 mM Tris-HCl, 5 mM Tris base and 2 mM EGTA) containing protease inhibitors to create a homogeneous slurry. The slurry was spun for 1 min at 500 g and the supernatant removed. The pellet was washed twice with a rigor buffer containing detergent (2% Triton X 100) to remove membranes. An aliquot of myofibrils was dissolved in SDS sample buffer and total protein measured by the Bradford assay (Bradford, 1976). Myofibrillar protein (0.25 µg per sample) was analysed on SDS-acrylamide gels by modified Laemmli procedure as previously described (McCormick et al. 2004). After electrophoresis, gels were silver stained (Oakley et al. 1980) and an image of each gel captured on a scanner (Hewlett Packard Scan Jet 6100c). The relative amount of each MHC isoform was determined by densitometry (McCormick et al. 2004).
Muscle fibre size
Plantaris muscle fibre size was determined from transverse 10 µm sections. Consecutive sections were cut, mounted on slides, and immunostained with a monoclonal antibody against all adult fast MHC isoforms (Sigma M4276) or type IIa MHC isoform (Developmental Hybridoma Bank of Iowa, Iowa City, IA, USA). Prior to incubation with primary antibody, non-specific antibody binding was blocked by incubating sections in phosphate-buffered saline (PBS) containing 0.5% bovine serum albumin (BSA) and 0.05% Tween-20 for 15 min at room temperature. Sections were then incubated with primary antibody overnight at 4°C. Following overnight incubation in primary antibody, sections were incubated with goat antimouse FITC-conjugated IgG (Jackson ImmunoResearch Laboratories, West Grove, PA, USA) diluted in a solution of 0.5% BSA, 0.05% Tween in PBS for 1 h at room temperature. Slides were mounted in antifade mounting medium (90% glycerol, 0.1 M Tris, pH 9.0, 0.2 M n-propyl gallate). The primary antibody step was omitted for some sections as a negative control. Immunostained sections were examined under epifluorescent illumination and digital images of each section were obtained. Fibres were classified as type I if they did not react with either fast antibody. Fibres were classified as type IIa if they reacted with both fast antibodies. Fibres were classified as type IIx/IIb if they reacted with the antibody against all fast myosins, but not with the antibody against type IIa myosin. The cross-sectional area of each typed fibre was measured using a computerized image analysis program (Image-Pro Plus, Media Cybernetics, Inc., Silver Spring, MD, USA). At least 100 fibres in each plantaris muscle were typed and measured.
Statistical analysis
Data was analysed by a one-factor analysis of variance (ANOVA) to test for differences among groups. Statistical significance was set at P 
0.05. Bonferroni's post hoc test was used to identify which groups were different from one another if a significant main group effect was found by ANOVA.
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Final uterus, body and plantaris muscle weights are presented in Table 1. Ovariectomy was effective, as indicated by the 65% reduction in uterus weight in the OVX/CO group. The final body weight of the OVX/E2 group was approximately 14% less than the other two groups despite all groups having similar food intake (Sham, 13.2 g day–1; OVX/CO, 13.3 g day–1; and OVX/E2, 12.4 g day–1). Similar to body weight, plantaris muscle weight was reduced (by 10%) in the OVX/E2 group. The plantaris-to-body weight ratio was similar in the three groups.
|
Plantaris muscle fibre size was determined on consecutive sections that were immunostained for either fast myosin or type IIa myosin. The immunostaining procedure allowed fibres to be identified as type I, type IIa or type IIb/IIx (Fig. 1). Average fibre size for each group is presented in Table 2. Two Sham animals were omitted from the fibre size analysis because their type I fibre areas were greater than two standard deviations from the mean (4048 and 3933 µm2 versus 2206 ± 753 µm2 for all animals in the study). The reason for the abnormally large type I fibres in these two Sham animals is unknown. Average type II fibre size was significantly reduced in the OVX/E2 group. Of the fast fibres, only the type IIb/IIx fibres were significantly smaller; type IIa fibre size was not different from the Sham group. In the OVX/CO group, the size of all fibre types was similar to that of the Sham group.
|
|
Myosin heavy chain composition was determined by SDS-PAGE. An example of a silver-stained gel appears in Fig. 2. The types I, IIa, IIx and IIb isoforms were clearly resolved. The relative amount of each isoform for the different groups appears in Table 3. With ovariectomy (OVX/CO), the relative amount of type IIx MHC was reduced. This effect was reversed when oestrogen was reintroduced into the system (OVX/E2). The combination of ovariectomy and oestrogen (OVX/E2) caused a reduction in the relative amount of type IIb MHC, although there was no change with ovariectomy alone. There were no significant differences in the relative amount of type IIa MHC among the three groups. There was a trend towards increased type I MHC in the OVX/E2 group (P = 0.07).
|
|
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Ovariectomized and oestrogen-replaced animals were pair-fed with sham-operated animals to control for differences in food intake that could affect growth. Food intake among the groups was similar, but not identical. The oestrogen-replaced animals consumed 6% less food per day than the other two groups. This small difference in food intake may have contributed to the reduced body and plantaris muscle weights of the oestrogen-replaced group, but is not likely to be solely responsible since body weight gain relative to food intake was also reduced in oestrogen-replaced animals (McCormick et al. 2004). The reduction in relative weight gain suggests that oestrogen replacement alters the proportion of calories that are allocated to growth.
We previously reported that oestrogen replacement reduced both fast and slow muscle fibre size in the soleus muscle of these animals (McCormick et al. 2004). In contrast, only the fastest fibres (type IIb/IIx) in the plantaris muscle were affected by oestrogen replacement. Plantaris fibres that were identified as type IIb/IIx based on immunostaining were 15% smaller in oestrogen-replaced animals than in sham-operated and ovariectomized animals. Our immunostaining procedure did not permit us to distinguish between type IIb and IIx fibres, but the decrease in the relative amount of type IIb myosin heavy chain as revealed by SDS-PAGE analysis suggests that the predominate effect of oestrogen replacement was on type IIb fibres. This conclusion is in agreement with a previous study showing that oestrogen treatment caused the greatest reduction in type IIb fibre size in the caudofemoralis muscle from growing, ovariectomized rats (Kobori & Yamamuro, 1989). Similar to the plantaris muscle, the caudofemoralis is composed predominately of fast fibres (Ariano et al. 1973). The cause of fibre type-specific effects cannot be determined from this study. It is possible that the fastest fibres are the most responsive to oestrogen because of differences in oestrogen receptor number and/or sensitivity, but to our knowledge no study has examined this issue.
Findings from the plantaris muscle support our previous conclusion that growing muscle is more responsive to oestrogen than mature muscle. Growing animals show reduced muscle mass and fibre size in response to oestrogen replacement (Ihemelandu, 1980, 1981; Suzuki & Yamamuro, 1985; Kobori & Yamamuro, 1989; McCormick et al. 2004), while muscles in mature animals do not appear to be affected (Fisher et al. 2000). If the results from animals hold true for growing humans, the population most at risk for altered skeletal muscle mass and fibre size might be amenorrhoeic adolescent girls who are prescribed oestrogen replacement therapy. To our knowledge no study has examined the effect of oestrogen replacement therapy on muscle growth in amenorrhoeic girls. However, given the relationship between longitudinal bone growth and lean body mass accretion during the pubertal growth spurt, a relationship between ovarian hormone status and lean body mass is likely. For example, normal girls experience their fastest gain in height (peak height velocity) slightly before they experience their fastest gain in lean body mass (age of peak height velocity, 11.8 years; age of peak lean body mass accretion, 12.2 years; Rauch et al. 2004). A close temporal association between longitudinal bone growth and skeletal muscle growth is consistent with studies from animals showing that longitudinal stretch is a powerful stimulus to skeletal muscle growth (Goldspink, 1977; Goldspink et al. 1986; Loughna et al. 1986). In amenorrhoeic girls, the relationship between longitudinal bone growth and muscle growth may be disrupted because they experience delays in epiphyseal fusion. Treating amenorrhoeic girls with oestrogen could theoretically result in early epiphyseal fusion, which in turn could reduce muscle growth by abolishing the stretch stimulus that comes from longitudinal bone growth.
Oestrogen replacement in ovariectomized animals caused a reduction in the relative amount of the fastest MHC isoform (IIb), but there was no difference in the type IIb MHC isoform in the plantaris with ovariectomy alone. One explanation for these observations is that another ovarian hormone might counteract the effect of oestrogen in an intact animal. Testosterone is one possible candidate. Average serum testosterone levels remain relatively constant from birth to adulthood in female rats and are consistently higher than oestrogen levels (Dohler & Wuttke, 1975). Serum testosterone levels vary with the oestrus cycle. Like oestrogen, testosterone levels are lowest during oestrus (83 ± 17 pg ml–1) and highest during proestrus (160–180 pg ml–1; Dupon & Kim, 1973). There is some evidence that testosterone may affect the expression of type IIb MHC. For example, the appearance of the type IIb isoform in the temporalis muscle is correlated with the increase in testosterone that occurs with sexual maturation in male guinea pigs (Lyons et al. 1986). Castration can prevent the appearance of the type IIb isoform in male guinea-pigs, while treating females with testosterone can increase type IIb expression (Lyons et al. 1986). In addition, testosterone administration increases the relative amount of type IIb MHC in the diaphragm muscle of mature rats (Prezant et al. 1997). The effect of testosterone on the plantaris muscle has not been studied, but it may be that testosterone stimulates type IIb MHC expression, while oestrogen inhibits it, so there is no net effect in an intact or ovariectomized animal. However, in an ovariectomized animal with oestrogen replacement, where oestrogen is present in the absence of testosterone, the effect of oestrogen on MHC expression may dominate.
Ovariectomy decreased the relative amount of type IIx MHC in the plantaris muscle, but did not affect the relative expression of any other isoform. This finding is in conflict with a previous investigation in which ovariectomy reduced the relative amount of type IIb MHC and increased the amount of type IIa MHC in the extensor digitorum longus muscle, another fast muscle (Kadi et al. 2002). The reasons for this conflict are not clear. It may be related to any number of factors, including differences in the plantaris and EDL muscles, rat strain (spontaneously hypertensive versus Sprague–Dawley in the present study) or age (not specified in the previous study versus 8 weeks old in the present study).
Oestrogen replacement countered the effect of ovariectomy on type IIx MHC expression in the present study, suggesting that this effect was oestrogen mediated. It is unknown whether oestrogen affects MHC expression by acting directly on oestrogen receptors in skeletal muscle. Oestrogen receptors (ERs) have been found in rat skeletal muscle (Dahlberg, 1982), so there is potential for receptor-mediated effects. In the classical ER pathway, ligand binding causes the ER to enter the nucleus, where it binds an oestrogen-specific response element (ERE) to regulate transcription. The consensus ERE is a palindrome composed of two half-site motifs separated by three nucleotides (5'-GGTCAnnnTGACC-3'; Klinge, 2001). There is some evidence that the ER can also bind widely spaced direct repeats of the first half-site motif (GGTCA) to regulate transcription (Kato et al. 1995). Interestingly, the rat IIx gene (rat genome database (RGD): 735061) promoter region has two half-site ERE motifs, making it possible that oestrogen regulates type IIx MHC expression in this way.
Alternatively, oestrogen may alter MHC expression by modulating levels of other hormones that regulate myosin expression. For example, ovariectomy reduces serum thyroid hormone concentration (triiodothyronine, T3), a reduction that is reversed with oestrogen replacement (Thomas et al. 1986). Thyroid deficiency reduces type IIx MHC expression in growing rats (Adams et al. 2000).
In conclusion, this study supports the notion that skeletal muscle is an oestrogen-responsive tissue. The response appears to vary depending on the fibre type composition of the muscle. In the predominantly slow soleus muscle, oestrogen replacement reduced the size of all fibres, both fast and slow, but MHC expression was not affected. In contrast, oestrogen replacement reduced the size of only the fastest fibres (IIx/IIb) and reduced type IIb MHC expression in the predominantly fast plantaris muscle. Additionally, ovariectomy caused a reduction in type IIx MHC expression that was reversed with oestrogen replacement. Further work is needed to distinguish effects that are mediated by the oestrogen receptors on skeletal muscle and those that are secondary to the interactions of oestrogen with other hormones.
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Ariano MA, Armstrong RB & Edgerton VR (1973). Hindlimb muscle fiber populations of five mammals. J Histochem Cytochem 21, 51–55.[Abstract]
Bradford MM (1976). A rapid and sensitive method for quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72, 248–254.[CrossRef][Medline]
Dahlberg E (1982). Characterization of the cytosolic estrogen receptor in rat skeletal muscle. Biochim Biophys Acta 717, 65–75.[Medline]
Davidge ST, Zhang Y & Stewart KG (2001). A comparison of ovariectomy models for estrogen studies. Am J Physiol 280, R904–R907.
Denis R, Rochon L, Deshaies Y & Denis R (1987). Effects of exercise training on energy balance of ovariectomized rats. Am J Physiol 253, R740–R745.
Dohler KD & Wuttke W (1975). Changes with age in levels of serum gonadotrophins, prolactin, and gonadal steroids in prepubertual male and female rats. Endocrinology 97, 898–907.[Abstract]
Dupon C & Kim MH (1973). Peripheral plasma levels of testosterone, androstenedione, and oestradiol during the rat oestrous cycle. J Endocrinol 59, 653–654.[Medline]
Fisher JS, Kohrt WM & Brown M (2000). Food restriction suppresses muscle growth and augments osteopenia in ovariectomized rats. J Appl Physiol 88, 265–271.
Goldspink DF (1977). The influence of immobilization and stretch on protein turnover of rat skeletal muscle. J Physiol 264, 267–283.
Goldspink DF, Morton AJ, Loughna P & Goldspink G (1986). The effect of hypokinesia and hypodynamia on protein turnover and the growth of four skeletal muscles of the rat. Pflugers Arch 407, 333–340.[CrossRef][Medline]
Ihemelandu EC (1980). Effect of oestrogen on muscle development of female rabbits. Acta Anat 108, 310–315.[Medline]
Ihemelandu EC (1981). Comparison of effect of oestrogen on muscle development of male and female mice. Acta Anat 110, 311–317.[Medline]
Jansson JO, Ekberg S, Isaksson OG & Eden S (1984). Influence of gonadal steroids on age- and sex-related secretory patterns of growth hormone in the rat. Endocrinology 114, 1287–1294.[Abstract]
Kadi F, Karlsson C, Larsson B, Eriksson J, Larval M et al. (2002). The effects of physical activity and estrogen treatment on rat fast and slow skeletal muscles following ovariectomy. J Muscle Res Cell Motil 23, 335–339.[CrossRef][Medline]
Kalu DN, Arjmandi BH, Liu CC, Salih MA & Birnbaum RS (1994). Effects of ovariectomy and estrogen on the serum levels of insulin-like growth factor-I and insulin-like growth factor binding protein-3. Bone Miner 25, 135–148.[Medline]
Kato S, Sasaki H, Suzawa M, Masushige S, Tora L et al. (1995). Widely spaced, directly repeated PuGGTCA elements act as promiscuous enhancers for different classes of nuclear receptors. Mol Cell Biol 15, 5858–5867.[Abstract]
Klinge CM (2001). Estrogen receptor interaction with estrogen response elements. Nucl Acids Res 29, 2905–2919.
Kobori M & Yamamuro T (1989). Effects of gonadectomy and estrogen administration on rat skeletal muscle. Clin Orthop 243, 306–311.
Lemoine S, Granier P, Tiffoche C, Rannou-Bekono F, Thieulant ML & Delamarche P (2003). Estrogen receptor alpha mRNA in human skeletal muscles. Med Sci Sports Exerc 35, 439–443.[CrossRef][Medline]
Loughna PT, Goldspink G & Goldspink DF (1986). Effect of inactivity and passive stretch on protein turnover in phasic and postural rat muscles. J Appl Physiol 61, 173–179.
Lyons GE, Kelly AM & Rubinstein NA (1986). Testosterone induced changes in contractile protein isoforms in the sexually dimorphic temporalis muscle of the guinea pig. J Biol Chem 261, 13278–13284.
McCormick KM, Burns KL, Piccone CM, Gosselin LE & Brazeau GA (2004). Effects of ovariectomy and estrogen on skeletal muscle function in growing rats. J Muscle Res Cell Motil 25, 21–27.[CrossRef][Medline]
Oakley BR, Kirsch DR & Morris NR (1980). A simplified ultrasensitive silver stain for detecting proteins in polyacrylamide gels. Anal Biochem 105, 361–363.[CrossRef][Medline]
Prezant DJ, Karwa ML, Kim HH, Maggiore D, Chung V & Valentine DE (1997). Short- and long-term effects of testosterone on diaphragm in castrated and normal male rats. J Appl Physiol 82, 134–143.
Rauch F, Bailey DA, Baxter-Jones A, Mirwald R & Faulkner R (2004). The ‘muscle-bone unit’ during the pubertal growth spurt. Bone 34, 771–775.[Medline]
Schaible TF, Malhotra A, Ciambrone G & Scheuer J (1984). The effects of gonadectomy on left ventricular function and cardiac contractile proteins in male and female rats. Circ Res 54, 38–49.
Scheuer J, Malhotra A, Schaible TF & Capasso J (1987). Effects of gonadectomy and hormonal replacement on rat hearts. Circ Res 61, 12–19.
Suzuki S & Yamamuro T (1985). Long-term effects of estrogen on rat skeletal muscle. Exp Neurol 87, 291–299.[CrossRef][Medline]
Thomas DK, Storlien LH, Bellingham WP & Gillette K (1986). Ovarian hormone effects on activity, glucoregulation and thyroid hormones in the rat. Physiol Behav 36, 567–573.[CrossRef][Medline]
Toth MJ, Poehlman ET, Matthews DE, Tchernof A & MacCoss MJ (2001). Effects of estradiol and progesterone on body composition, protein synthesis, and lipoprotein lipase in rats. Am J Physiol 280, E496–E501.
|
Acknowledgements |
|---|
This article has been cited by other articles:
|
|
 |
|
  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 | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
