Department of Molecular and Cellular Physiology, University of Cincinnati, Cincinnati, Ohio 45267-0576
Submitted 19 November 2003 ; accepted in final form 30 July 2004
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
dihydropyridine receptor mRNA; decreased use; passive tension; denervation; tenotomy; hindlimb immobilization
In contrast to the established role of regulated gene expression for the contractile and metabolic proteins, much less is known about the regulation by use of genes that specify membrane proteins involved in muscle activation, although they also contribute to contractile performance. Muscle activation is controlled by voltage-driven movements of the dihydropyridine receptor (DHPR), an L-type calcium channel that associates with the ryanodine receptor/calcium release channel at specialized internal junctions with the sarcoplasmic reticulum (10). The DHPR is absolutely required for excitation-contraction coupling (37), and its content in phenotypically different skeletal muscles is a marker of contractile speed, being more abundant in fast- compared with slow-twitch muscles (18).
Previous studies of the regulation of DHPR gene expression by muscle use produced mixed results. Those studies focused primarily on the slow-twitch soleus (Sol) muscle subjected to disuse produced by removing mechanical load (18) or nerve input (28). DHPR 1s mRNA in the Sol muscle increases to levels normally seen in the extensor digitorum longus (EDL) muscle after 28 days of unloading by hindlimb suspension (18) or 50-day denervation (28). A few studies using phenotypically fast-twitch muscles report weak or no regulation of DHPR mRNA during denervation-induced disuse and led to the proposal that DHPR gene expression is not highly regulated in fast-twitch muscles. DHPR mRNA increases moderately in the mouse Sol muscle and less in the flexor digitorum longus after 15-day denervation (36) and does not change in the rat EDL muscle during long-term (50 day) denervation (28) or mechanical unloading (18).
A few studies have examined the effect of increased use on DHPR gene expression. DHPR 1s mRNA and protein both decrease significantly when a fast-twitch muscle is stimulated chronically in a pattern typical for slow-twitch muscle (27), suggesting that regulated DHPR gene expression can occur in association with known phenotype transitions. Additionally, DHPR protein content does not change in the rat plantaris muscle subjected to increased mechanical load (19) and increases in fast- and slow-twitch rat muscles after 12 wk of moderate exercise training (33), but the contribution of regulated gene expression to these adaptations is not known.
Collectively, these studies establish that DHPR gene expression is regulated during altered muscle activity, especially by decreased activity or load. However, they do not reveal a clear relationship between the change in use and the change in DHPR mRNA, suggesting that important physiological signals that control its regulation remain to be identified.
To address this question, we examined further the regulation of DHPR gene expression in phenotypically different skeletal muscles during physiological adaptations to disuse in vivo. We focused on disuse because it produces a measurable end point, atrophy, that has broad clinical relevance. We subjected the Sol, tibialis anterior (TA), gastrocnemius (Gastr), and EDL muscles of the mouse to denervation, tenotomy, or limb immobilization and measured changes in DHPR 1s mRNA with quantitative Northern blot analysis. DHPR
1s protein in immobilized Gastr and TA muscles was measured by Western blot. Each of these models produces disuse by a different mechanism, thereby allowing us to dissociate the contributions of nerve input and passive or active tension to DHPR gene regulation. Denervation eliminates nerve input, both electrical and trophic, thereby rendering the muscle inactive while preserving tendon attachments and resting tension. Tenotomy preserves nerve input at near-normal levels (39) but unloads and shortens the muscle, preventing it from generating either passive (resting) or active tension. Immobilization preserves both nerve input and tendon attachments, while allowing resting tension to be manipulated by fixing the muscle in a lengthened or shortened position (16). Resting tension decreases in shortened but immobile muscles and increases significantly in lengthened but immobile muscles (16). Tenotomy and limb immobilization have not been used previously to study DHPR gene expression. A preliminary report of this work has appeared (29, 30).
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Selection of control muscles for denervation measurements.
It is widely accepted that the contralateral muscles of animals subjected to unilateral denervation can hypertrophy because of extra weight bearing by the untreated limb, with associated changes in metabolism or gene expression. However, hypertrophy of the contralateral muscles does not always occur (40). Consequently, we examined which muscles would best serve as control under the conditions of this study. We found that DHPR 1s mRNA content of the TA muscle of untreated mice is comparable to that in the contralateral TA muscle of mice subjected to 14-day unilateral denervation of the opposite leg (Fig. 1 and Table 1). Overall, the DHPR mRNA of contralateral muscles tended to stay the same (TA, Gastr) or to decrease by 1115% (Sol, EDL) compared with control, untreated muscles. The wet weight of contralateral muscles tended to decrease, but the changes did not reach statistical significance. Thus, under our conditions, the contralateral muscles of mice denervated for 14 days do not undergo any significant hypertrophy. Additionally, the small decrease in DHPR
1s content in the contralateral Sol and EDL muscles would tend to overestimate any potential increase in DHPR mRNA caused by denervation. For these reasons, the muscles of untreated mice were used as controls for analysis of changes in the DHPR mRNA content of 14-day treated mice. Additionally, these measurements establish that our assay can detect statistically significant changes in DHPR
1s mRNA content as small as 1115% in mouse skeletal muscle samples. DHPR mRNA is more abundant in the mouse EDL than Sol muscle, with a ratio of
1s in Sol to
1s in EDL of 0.66, similar to that in rat (ratio = 0.65; Ref. 27).
|
|
Organ culture. Freshly dissected Sol and EDL muscles were fixed with pins to the bottom of a 4-ml culture dish, four muscles/dish, and incubated for 1924 h in a medium optimized for adult skeletal muscle culture [M199 (GIBCO-BRL), as modified in Ref. 41, at 37°C in an atmosphere of 95% air-5% CO2]. The muscles were fixed either at resting length or stretched to 1.11.25 times resting length. Resting lengths of the EDL and Sol muscles were 12.7 ± 0.1 (n = 10) and 9.3 ± 0.1 (n = 9) mm, respectively.
The average sarcomere length of stretched and unstretched muscles was measured at the end of the culture period by measuring the average of 10 sarcomere spans under a light microscope at x400 magnification from different regions of the muscle. This was done to verify that the muscles retained the imposed length change throughout the culture period and as a control for the possibility that length changes in nonmuscle elements such as a tendon could have occurred (e.g., if a tendon lengthened by tearing on the pins used to fix the muscle while the muscle shortened). Sarcomere lengths at 24 h were 13% greater in stretched EDL than EDL muscles cultured without stretch and 17% greater in stretched Sol than Sol muscles cultured without stretch (Table 2), confirming that the stretched muscles retained the lengthened position throughout the culture period. Before harvesting, each muscle was stimulated electrically to confirm that it retained the ability to generate evoked contractions.
|
Statistical analysis. Values are given as means ± SE. Differences in mean values were evaluated with an unpaired Students t-test, with significance accepted at P < 0.05.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
DHPR 1s mRNA in TA and EDL muscles after peroneal nerve cut.
The finding that denervation can preferentially upregulate DHPR mRNA in the Sol but not EDL muscle is consistent with previous reports that show a greater effect of denervation on slow-twitch compared with fast-twitch muscles (28). However, the finding that DHPR mRNA is strongly upregulated in the fast-twitch TA muscle contrasts with previous interpretations that the regulation by denervation occurs mainly in slow-twitch muscles. To investigate the basis for this difference, we removed nerve input to a more limited subset of muscles by performing a 14-day section of the peroneal nerve. In contrast to sciatic nerve section, which denervates all four muscles, peroneal section selectively denervates the TA and EDL muscles. Figure 3 shows that DHPR
1s gene expression is remarkably upregulated in both TA (1.8-fold) and EDL (1.6-fold) muscles after denervation compared with the corresponding innervated controls. DHPR mRNA tended to increase in the TA muscle after sciatic compared with peroneal nerve cut (1.8-fold compared with 1.6-fold). The TA muscle underwent comparable paralysis and atrophy in both treatments. Muscle mass decreased by 57% (n = 18 muscles) after peroneal section and by 55% after sciatic section (n = 16 muscles); the differences were not significant. The DHPR mRNA content of the EDL muscle increased significantly after peroneal nerve section (1.6-fold) compared with no change after sciatic section (1.05-fold; significantly different at P < 0.05). These results suggest that an important factor other than nerve input or denervation-related disuse per se contributes to DHPR gene regulation under these conditions; for example, DHPR mRNA is upregulated in the TA muscle after sciatic nerve section and in the TA and EDL muscles after peroneal nerve section but unchanged in the Gastr muscle, whether denervated (sciatic cut) or innervated (peroneal cut) (data not shown). We hypothesized that the effect of denervation on DHPR gene expression may depend on the passive tension on the denervated muscle, which can be influenced by the surrounding muscles. The mice continue to ambulate after both sciatic and peroneal nerve section but do so in different ways. All lower leg muscles are inactive after sciatic cut, and the animal uses them in variable and unpredictable ways for support and balance, whereas after peroneal cut the dorsal muscles including Gastr and Sol muscles remain innervated and contract during ambulation. In this case, active shortening of the Gastr and Sol muscles is expected to lengthen the TA and EDL muscles in a consistent manner.
|
As expected, unilateral immobilization of one limb for 4 days did not produce atrophy or hypertrophy in either the immobilized or contralateral muscles, and the contralateral muscle was used as reference. Figure 4 compares changes in DHPR 1s mRNA content in the Sol, TA, EDL, and Gastr muscles after unilateral limb immobilization in a maximally dorsiflexed position, which maintains the Sol and Gastr muscles in a lengthened position (increased passive tension) and the TA and EDL muscles at a shorter than normal resting length (decreased passive tension). DHPR
1s mRNA changed dramatically in all muscles, being upregulated in the Sol and Gastr muscles and downregulated in the TA and EDL muscles. Sorting these results by the change in passive tension (Fig. 4B) reveals that the change in DHPR mRNA correlates directly with the passive tension on the muscle. DHPR
1s mRNA increases in the lengthened Sol and Gastr muscles (to 229 ± 23% and 151 ± 21% of control, respectively) and decreases in the shortened TA and EDL muscles (to 73 ± 6% and 75 ± 4% of control, respectively). This result demonstrates that resting tension per se is an important physiological regulator of DHPR gene expression.
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
DHPR gene expression and control of muscle size during disuse.
Sustained disuse of skeletal muscles is associated with a loss in mass, or atrophy, which can vary in severity in phenotypically different muscles and models of disuse (9, 17, 20, 32). Muscle size is controlled by both translational and transcriptional mechanisms, with translational processes playing a greater role (13). Our results show a dissociation between DHPR gene expression and control of cell size in three models of disuse. For example, 2 wk of denervation or tenotomy induces significant and comparable atrophy in both slow- and fast-twitch mouse muscles, whereas DHPR gene expression can be up- or downregulated or unchanged. In the TA muscle after peroneal nerve cut, DHPR mRNA and cell size change in opposite directions. In muscles immobilized for 4 days, DHPR mRNA changes in opposite directions in different muscles without change in muscle size. Other models of disuse also suggest a dissociation between DHPR gene expression and cell size. DHPR 1s mRNA is upregulated +1.8-fold in the unloaded rat Sol muscle after 24 h, before atrophy develops (18), and it is upregulated to dramatically different degrees in rat Sol muscle subjected to 14-day TTX treatment or 14-day denervation, two models that produce comparable atrophy (31). This consistent dissociation in different models of disuse suggests that DHPR gene expression is regulated independently of the pathways that control muscle size, although the two processes can coincide.
DHPR gene expression and muscle phenotype during disuse.
Phenotypic changes occur in association with several models of disuse, mediated in large part by remodeling of MHC isoform gene expression (13). The relationship between DHPR gene expression and muscle phenotype during adaptations to disuse is not known. Our finding that DHPR 1s mRNA in the denervated Sol muscle increases to levels normally found in the fast EDL muscle is similar to results in the denervated rat Sol muscle (28) and in the unloaded rat Sol muscle (18), in which MHC isoform mRNAs shift toward a phenotypically faster profile (17). This could occur if common signals control DHPR gene expression and fiber phenotype. Our finding that DHPR mRNA can increase dramatically in phenotypically fast-twitch muscles after denervation was unexpected, because denervation of fast-twitch muscles shifts MHC mRNAs toward a slower profile (17), a change that predicts less DHPR mRNA. Other studies of DHPR mRNA in slow- and fast-twitch rat muscles report little or no change after denervation (31). The present study reexamined this question with both global and selective denervation and included three muscles that are composed predominantly of fast fiber types (6). Our results demonstrate clearly that DHPR gene expression can be regulated in fast-twitch muscles during denervation disuse but show no consistent relationship between regulation of DHPR gene expression and muscle phenotype or nerve input per se. It increases in immobilized, lengthened muscles, decreases in immobilized, shortened muscles, and declines in all tenotomized muscles, in every case independent of phenotype. Thus DHPR
1s gene expression is regulated in both fast- and slow-twitch muscles during disuse, but the regulation is not linked to the change in muscle phenotype that can accompany these models.
Passive tension regulates DHPR gene expression.
Results obtained with cast immobilization reveal that DHPR 1s mRNA is highly sensitive to the passive tension on the muscle, a parameter that is not controlled in the other models. Short-term cast immobilization allowed us to subject phenotypically different muscles to sustained disuse in either a shorter or a lengthened position while preserving innervation and tendon attachments, in each case before detectable changes in muscle size. In vitro measurements confirm the effect of stretch and further suggest that some degree of resting tension is required to maintain DHPR mRNA expression. Cultured muscles, which lack all tendon attachments and resting tension, downregulate DHPR mRNA dramatically within 24 h; however, much of this loss can be prevented by applying even a small amount of stretch. Collectively, these results demonstrate that passive tension is an important physiological signal that controls DHPR gene expression in all muscle types. Increased stretch to a muscle upregulates DHPR gene expression, whereas decreased tension downregulates it.
DHPR gene expression in the denervated and tenotomized muscles can be understood in this context. After sciatic nerve section, the sampled muscles do not generate active contraction or undergo the length changes that accompany normal ambulation. In this case, the change in DHPR mRNA is variable and does not correlate with muscle phenotype or size. However, when the TA and EDL muscles are selectively denervated by peroneal nerve cut, the Gastr and Sol muscles remain innervated and generate active contractions during body movements, which in turn can passively stretch the inactive TA and EDL muscles. The finding that DHPR mRNA is dramatically upregulated in inactive TA and EDL muscles suggests that passive stretch, more than nerve input or inactivity per se, is the major stimulus for DHPR gene expression under these conditions. In the case of tenotomy, all affected muscles shorten and cannot undergo passive stretching during body movement (1) and DHPR mRNA decreases dramatically in all muscles. Thus shortening a muscle or decreasing resting tension by any of these manipulations downregulates DHPR gene expression.
This conclusion differs from a previous study in which it was proposed that a decrease in chronic tension stimulates increased DHPR gene expression (18). DHPR mRNA increases significantly in the rat Sol muscle unloaded by hindlimb suspension, before atrophy develops. Because passive tension is not fixed, it is possible that the chronic tension on the Sol muscle may be different than assumed or that another signal may act synergistically with passive tension in this model.
Regulation of DHPR gene expression by stretch compared with other muscle genes. Previous studies have highlighted the importance of passive stretch as a signal for muscle growth, phenotypic differentiation, and size (11, 14, 15). Maintained stretch is required for the growth and survival of muscles in culture (26, 38). Stretch alone, independent of nerve input or other mechanical signals, can serve as a stimulus for both phenotypic remodeling and muscle size. Increased stretch leads to hypertrophy even if a muscle is inactive, and, conversely, a muscle that is stimulated chronically can atrophy if stretch is not imposed (12, 16). Stretch can oppose the atrophy that occurs in immobilized and shortened muscles (32), and this model has been used to identify early and downstream genes involved in stretch-induced hypertrophy (21, 22, 34), which may use independent but interacting pathways (34). Identified stretch-responsive skeletal muscle genes include the MHC isoform genes Ankrd2 (21), Smpx (22), and Serhl (34), IGF-1, mechanogrowth factor (42), and muscle regulatory factors (5, 23, 24). Of these, the MHC genes are the best characterized. Stretch makes a greater contribution than activity alone to MHC gene expression (13) and can influence MHC gene expression in denervated muscles in a muscle-specific manner (25). Stretch is required to maintain expression of the slow and neonatal MHC mRNAs that predominate in slower or less differentiated fiber types and to repress expression of the fast MHC mRNAs (13, 14, 24). In this respect, our finding that increased passive stretch promotes increased DHPR mRNA, a direction toward a faster muscle phenotype, is a direction opposite to the regulation of MHC mRNAs by stretch.
Relationship between DHP mRNA and protein in response to stretch. The finding that DHPR mRNA and protein do not follow closely at 4 days suggests that DHPR expression is controlled at multiple levelsincluding both transcription and protein translation/turnover. This result does not mean that changes in DHPR protein do not occur but rather that, if they do occur, their turnover rate is not tightly synchronized with DHPR mRNA. In general, multiple levels of regulation are expected for a protein as essential to muscle function as DHPR. Studies of DHPR mRNA and protein in other models of disuse are not consistent on this point. DHPR mRNA is increased in 25- to 50-day denervated rat muscles (28), but DHPR mRNA and protein are not changed detectably at 10 days after denervation (31). In denervated chick skeletal muscles, DHPR protein increases within 3 days of denervation, peaks at 15 days, then declines as atrophy becomes evident (35). More measurements of the stability and turnover of both DHPR mRNA and protein, in isolation from changes in cell size or phenotype, are required to evaluate this question.
In conclusion, this study adds DHPR to the growing list of muscle genes whose expression is regulated by stretch. A direction for future research is to identify the molecular stretch sensors and signaling pathways that control DHPR gene expression and their relationship to the pathways that control muscle phenotype and size.
![]() |
FOOTNOTES |
---|
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 |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
2. Baldwin KM and Haddad F. Effects of different activity and inactivity paradigms on myosin heavy chain gene expression in striated muscle. J Appl Physiol 90: 345357, 2001.
3. Baldwin KM and Haddad F. Skeletal muscle plasticity: cellular and molecular responses to altered physical activity paradigms. Am J Phys Med Rehabil 81: S40S51, 2002.[CrossRef][ISI][Medline]
4. Booth FW and Criswell DS. Molecular events underlying skeletal muscle atrophy and the development of effective countermeasures. Int J Sports Med 18, Suppl 4: S265S269, 1997.[ISI][Medline]
5. Booth FW and Thomason DB. Molecular and cellular adaptation of muscle in response to exercise: perspectives of various models. Physiol Rev 71: 541585, 1991.
6. Burkholder TJ, Fingado B, Baron S, and Lieber RL. Relationship between muscle fiber types and sizes and muscle architectural properties in the mouse hindlimb. J Morphol 221: 177190, 1994.[ISI][Medline]
7. Chomczynski P and Sacchi N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162: 156159, 1987.[CrossRef][ISI][Medline]
8. Cougnon MH, Moseley AE, Radzyukevich TL, Lingrel JB, and Heiny JA. Na,K-ATPase - and
-isoform expression in developing skeletal muscles:
2 correlates with t-tubule formation. Pflügers Arch 445: 123131, 2002.[CrossRef][ISI][Medline]
9. Edgerton VR, Roy RR, Allen DL, and Monti RJ. Adaptations in skeletal muscle disuse or decreased-use atrophy. Am J Phys Med Rehabil 81: S127S147, 2002.[CrossRef][ISI][Medline]
10. Franzini-Armstrong C, Protasi F, and Ramesh V. Comparative ultrastructure of Ca2+ release units in skeletal and cardiac muscle. Ann NY Acad Sci 853: 2030, 1998.
11. Goldspink DF, Cox VM, Smith SK, Eaves LA, Osbaldeston NJ, Lee DM, and Mantle D. Muscle growth in response to mechanical stimuli. Am J Physiol Endocrinol Metab 268: E288E297, 1995.
12. Goldspink G. Selective gene expression during adaptation of muscle in response to different physiological demands. Comp Biochem Physiol B Biochem Mol Biol 120: 515, 1998.[CrossRef][ISI][Medline]
13. Goldspink G. Changes in muscle mass and phenotype and the expression of autocrine and systemic growth factors by muscle in response to stretch and overload. J Anat 194: 323334, 1999.[CrossRef][ISI][Medline]
14. Goldspink G, Scutt A, Loughna PT, Wells DJ, Jaenicke T, and Gerlach GF. Gene expression in skeletal muscle in response to stretch and force generation. Am J Physiol Regul Integr Comp Physiol 262: R356R363, 1992.
15. Goldspink G, Williams P, and Simpson H. Gene expression in response to muscle stretch. Clin Orthop S146S152, 2002.
16. Hnik P, Vejsada R, Goldspink DF, Kasicki S, and Krekule I. Quantitative evaluation of electromyogram activity in rat extensor and flexor muscles immobilized at different lengths. Exp Neurol 88: 515528, 1985.[CrossRef][ISI][Medline]
17. Huey KA and Bodine SC. Changes in myosin mRNA and protein expression in denervated rat soleus and tibialis anterior. Eur J Biochem 256: 4550, 1998.[Abstract]
18. Kandarian S, OBrien S, Thomas K, Schulte L, and Navarro J. Regulation of skeletal muscle dihydropyridine receptor gene expression by biomechanical unloading. J Appl Physiol 72: 25102514, 1992.
19. Kandarian SC, Peters DG, Favero TG, Ward CW, and Williams JH. Adaptation of the skeletal muscle calcium-release mechanism to weight-bearing condition. Am J Physiol Cell Physiol 270: C1588C1594, 1996.
20. Kandarian SC and Stevenson EJ. Molecular events in skeletal muscle during disuse atrophy. Exerc Sport Sci Rev 30: 111116, 2002.[ISI][Medline]
21. Kemp TJ, Sadusky TJ, Saltisi F, Carey N, Moss J, Yang SY, Sassoon DA, Goldspink G, and Coulton GR. Identification of Ankrd2, a novel skeletal muscle gene coding for a stretch-responsive ankyrin-repeat protein. Genomics 66: 229241, 2000.[CrossRef][ISI][Medline]
22. Kemp TJ, Sadusky TJ, Simon M, Brown R, Eastwood M, Sassoon DA, and Coulton GR. Identification of a novel stretch-responsive skeletal muscle gene (Smpx). Genomics 72: 260271, 2001.[CrossRef][ISI][Medline]
23. Loughna PT and Brownson C. Two myogenic regulatory factor transcripts exhibit muscle-specific responses to disuse and passive stretch in adult rats. FEBS Lett 390: 304306, 1996.[CrossRef][ISI][Medline]
24. Loughna PT, Izumo S, Goldspink G, and Nadal-Ginard B. Disuse and passive stretch cause rapid alterations in expression of developmental and adult contractile protein genes in skeletal muscle. Development 109: 217223, 1990.[Abstract]
25. Loughna PT and Morgan MJ. Passive stretch modulates denervation induced alterations in skeletal muscle myosin heavy chain mRNA levels. Pflügers Arch 439: 5255, 1999.[CrossRef][ISI][Medline]
26. Nakai J. Skeletal muscle in organ culture. Exp Cell Res 40: 307315, 1965.[ISI][Medline]
27. Pereon Y, Navarro J, Hamilton M, Booth FW, and Palade P. Chronic stimulation differentially modulates expression of mRNA for dihydropyridine receptor isoforms in rat fast twitch skeletal muscle. Biochem Biophys Res Commun 235: 217222, 1997.[CrossRef][ISI][Medline]
28. Pereon Y, Sorrentino V, Dettbarn C, Noireaud J, and Palade P. Dihydropyridine receptor and ryanodine receptor gene expression in long-term denervated rat muscles. Biochem Biophys Res Commun 240: 612617, 1997.[CrossRef][ISI][Medline]
29. Radzyukevich T and Heiny JA. Calcium channel gene expression during skeletal muscle atrophy (Abstract). Biophys J 78: 2554a, 2000.
30. Radzyukevich T and Heiny JA. Skeletal muscle DHPR gene expression is regulated by stretch and calcium (Abstract). Biophys J 80: 69a, 2001.[ISI]
31. Ray A, Kyselovic J, Leddy JJ, Wigle JT, Jasmin BJ, and Tuana BS. Regulation of dihydropyridine and ryanodine receptor gene expression in skeletal muscle. Role of nerve, protein kinase C, and cAMP pathways. J Biol Chem 270: 2583725844, 1995.
32. Roy RR, Baldwin KM, and Edgerton VR. The plasticity of skeletal muscle: effects of neuromuscular activity. Exerc Sport Sci Rev 19: 269312, 1991.[Medline]
33. Saborido A, Molano F, Moro G, and Megias A. Regulation of dihydropyridine receptor levels in skeletal and cardiac muscle by exercise training. Pflügers Arch 429: 364369, 1995.[ISI][Medline]
34. Sadusky TJ, Kemp TJ, Simon M, Carey N, and Coulton GR. Identification of Serhl, a new member of the serine hydrolase family induced by passive stretch of skeletal muscle in vivo. Genomics 73: 3849, 2001.[CrossRef][ISI][Medline]
35. Schmid A, Renaud JF, Fosset M, Meaux JP, and Lazdunski M. The nitrendipine-sensitive Ca2+ channel in chick muscle cells and its appearance during myogenesis in vitro and in vivo. J Biol Chem 259: 1136611372, 1984.
36. Shih HT, Wathen MS, Marshall HB, Caffrey JM, and Schneider MD. Dihydropyridine receptor gene expression is regulated by inhibitors of myogenesis and is relatively insensitive to denervation. J Clin Invest 85: 781789, 1990.[ISI][Medline]
37. Tanabe T, Beam KG, Powell J, and Numa S. Restoration of excitation-contraction coupling and slow calcium current in dysgenic muscle by dihydropyridine receptor complementary DNA. Nature 336: 134139, 1988.[CrossRef][ISI][Medline]
38. Vandenburgh H and Kaufman S. In vitro model for stretch-induced hypertrophy of skeletal muscle. Science 203: 265268, 1979.[ISI][Medline]
39. Vrbova G. Changes in the motor reflexes produced by tenotomy. J Physiol 166: 241250, 1963.[ISI]
40. Wallis MG, Appleby GJ, Youd JM, Clark MG, and Penschow JD. Reduced glycogen phosphorylase activity in denervated hindlimb muscles of rat is related to muscle atrophy and fibre type. Life Sci 64: 221228, 1999.[ISI][Medline]
41. Wetzel DM and Salpeter MM. Fibrillation and accelerated AChR degradation in long-term muscle organ culture. Muscle Nerve 14: 10031012, 1991.[ISI][Medline]
42. Yang S, Alnaqeeb M, Simpson H, and Goldspink G. Cloning and characterization of an IGF-1 isoform expressed in skeletal muscle subjected to stretch. J Muscle Res Cell Motil 17: 487495, 1996.[ISI][Medline]