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. Author manuscript; available in PMC: 2015 Jan 1.
Published in final edited form as: Can J Physiol Pharmacol. 2013 Aug 13;92(1):1–8. doi: 10.1139/cjpp-2013-0063

Non-surgically-induced disuse muscle atrophy and neuromuscular dysfunction upregulates alpha7 acetylcholine receptors

Mohammed A S Khan 1, Nita Sahani 1, Kevin A Neville 1, Michio Nagashima 1, Sangseok Lee 1, Kaneki Tomoki Sasakawa Masao 1, J A Jeevendra Martyn 1
PMCID: PMC4086193  NIHMSID: NIHMS596584  PMID: 24383867

Abstract

Previous models of disuse have invariably used surgical methods require repetitive plaster casts applications. A method of disuse atrophy that does not require repetitive application is described. A modified plastic pipette tubing was applied to one hindlimb from thigh to foot resulting in immobilization of knee in extension and ankle in plantar flexion position. This method resulted in loss of soleus muscle mass to 11, 22, 39, and 45% at 3, 7, 14 and 21 days, respectively, in association with a significant decrease of tibialis twitch (25%) and tetanic tensions (26%) at 21 days, compared to contralateral side and/or sham immobilized controls. Immunohistochemical analysis of soleus using fluorescent α-bungarotoxin revealed a significant increase in the number of synapses per unit area (818+31 vs 433+16/mm2) and increase in muscle fibers per unit area (117 vs 83/mm2) most likely related to atrophy of muscle fibers bringing synapses closer. A three fold increase in alpha7 acetylcholine receptor (α7AChR) protein expression along with increased expression of α1AChR subunit on immobilized vs contralateral side was observed. The physiology and pharmacology of the novel finding of upregulation of α7AChRs with disuse requires further study.

Keywords: α7acetylcholine receptors, external plastic casting, immobilization, muscle atrophy, neuromuscular dysfunction

Introduction

Skeletal muscle amounts to about 40 percent of the body mass in normals. In pathological states or muscle-related genetic disorders, there is loss of muscle mass and decreased muscle strength (Elizabeth and Peter 2007). Muscle wasting can also occur in the absence of systemic or genetic disease states as in spaceflights, prolonged bed rest, limb unloading or plaster casting (Brunelli and Rovere-Querini 2008; Grosset and Onambele-Pearson 2008; Larsson 2007; Kortebein et al. 2008; Ferrando et al. 2006; Zhang et al. 2007; Ferrando et al. 1996; Frick et al. 2008; Kong et al. 2009). Immobilization changes the structure and function of the peripheral neuromuscular system and alters both protein synthesis and degradation (Robinson et al. 1991).

The models used to study mechanisms of disuse-induced neuromuscular dysfunction and muscle wasting in rodents include spinal cord or motor nerve denervation, plaster casting, pinning of joints, and external splinting using staples (Damrongrungruang et al. 2004; Bajusz 1964; Frimel et al. 2005; Fink et al. 2008; Mokhtarian et al. 1999). Undoubtedly, previously enumerated methods to study muscle atrophy have been used extensively and are of great value. The aforementioned models of disuse, however, have some disadvantages. For example, pinning or stapling of the joints is associated with the trauma and foreign body-induced inflammatory process in the muscle (Fink et al. 2008; Caron et al. 2009). Tail suspension model has been successfully utilized to see atrophic response as a model for weightlessness that mimics spaceflight (Kandarian et al. 1996), but does not reflect immobilization of bed rest or casting as there is limb movement albeit lack of weight bearing in the former. Denervation or spinal cord transection results in loss of trophic factors from the nerve and does not truly reflect the bed rest or cast-induced immobilization, where there is no anatomical disruption of the motor nerves. Plaster casting often requires repetitive application due to chewing of the material by the animal (Frimel et al. 2005). In order to study molecular mechanisms of disuse (immobilization)-induced muscle atrophy, an effective non-surgical method, which does not require repetitive application and the absence of trauma- or surgery-induced inflammation in muscle confounding the signaling pathways, is necessary. The present report describes an easy and effective mouse model of disuse that produces the functional and biochemical changes of immobilization including (1) progressive loss of muscle mass in slow and fast muscles, (2) decreased single twitch and tetanic tensions in muscle and (3) increased expression of muscle alpha1 acetylcholine receptor (α1AChR) subunit. A novel finding is the upregulation of neuronal α7 acetylcholine receptors (α7AChRs) on muscle membrane, previously reported only following denervation.

Materials and methods

Immobilization by External Plastic Casting (EPC)

The protocol was approved by the Subcommittee on Research Animal Care at Massachusetts General Hospital. Male C56Bl/6J mice, 10 –12 weeks old and weighing 25– 28 g, (Jackson Laboratories) were anesthetized with pentobarbital (60–70 mg/kg i.p.) and the left hindlimb was immobilized by applying EPC for 3, 7, 14 and 21 days. Each experiment used groups of 4–5 mice. EPC was prepared from the plastic cylinder used of a pipette tube of 3 ml with a diameter of 1.2 and 0.8 cm at the proximal and distal ends, respectively. The proximal and distal ends of the pipette were cut to 3–4 cm length, which, when applied, extended from the upper thigh to the foot of the mouse. Initially, a piece of nylon (2.0) surgical suture was glued with TissueSeal® to the plantar aspect of the foot and used to pull the foot gently through the EPC. This places the foot in plantar-flexion and the knee joint in extension. The skin surfaces of the thigh and foot ends were attached with TissueSeal® to the inner surface of the EPC as an additional precaution to prevent any movement. Sponge padding was added at the proximal end to prevent pressure ulceration. The contralateral (non-immobilized) side served as the control. Sham-immobilized animals, subjected to same immobilization procedures except that the EPC was removed immediately, served as additional controls for the 21 day immobilization group.

Figure 1A demonstrates sequential pictures of the assembly of EPC, which produces non-surgical immobilization. We describe the preparation and application of EPC in five simple steps, which required only 5 to 10 min to complete the whole immobilization procedure. The method illustrated was very simple and materials used were inexpensive. Figure 1B1 provides a diagrammatic representation of EPC-immobilized hindlimb. The femoral (thigh) end and metatarsal (foot) end of the hindlimb were adhered to the inner wall of the tube with the help of TissueSeal® adhesive, which placed the knee joint in extension and foot in plantar flexion position. Figure 1B2 provides an image of the EPC-immobilized hindlimb at 14 days after EPC showing the effectiveness of the casting with knee in extension and ankle joint in plantar flexion. At the end of the immobilization period, the EPC was easily dismounted by cutting EPC under anesthesia (pentobarbital 60–70 mg/kg i.p.) with no injury to the leg. Image 1B3 is an X-ray image of the immobilized during EPC together with contralateral himdlimb. The X-ray image reveals that contralateral hindlimb was in the resting position, while immobilized hindlimb is in the stretched immobilized position. Figure 1B4 shows the immobilized limb after removal of the casting at 14 days. In all the images described in Fig. 1B (1–4), the contralateral (non-immobilized) limb was in the resting position. Naïve sham-immobilized (n=5) at 21 days and the contralateral limbs for all time points, served as controls to immobilized limb. Previous studies have indicated that the contralateral sham-immobilized limb behaves like sham-immobilized naïve animals (Fink et al. 2008).

Fig. 1. Stepwise process of hindlimb immobilization of mice by EPC.

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A. Brief description of each of 6 steps is described. B. Schematic, macroscopic and X-ray image of hindlimb immobilization of mouse. B1. Cartoon of EPC. B2. Picture of the hindlimb immobilized with EPC. B3. X-ray image of immobilized hindlimb and B4. Immobilized hindlimb after removal of EPC on day 14. The contralateral hindlimb is seen on the left.

Neuromuscular Function Test

Nerve evoked mechanomyography was performed after 21 days (n=4) of immobilization as described previously (Fink et al. 2008). The mice were anesthetized with pentobarbital (60–70 mg/kg i.p.), and tracheostomized and ventilated with air at ~140–150 breaths/minute with a tidal volume of 6–8 ml/kg (Hugo Saches Electronic-Harvard Apparatus). With the mice in dorsal recumbency, the tendon of insertion of the tibialis muscle was surgically exposed on each side and individually attached to separate grass FT03 force displacement transducers.

Stimulation electrodes on the sciatic nerve, exposed at the thigh, produced nerve-mediated stimulation of the tibialis muscles. The knee was stabilized with a clamp to prevent limb movement during nerve stimulation. Baseline tensions of approximately 10 grams, which yielded optimal evoked tensions, were applied on the immobilized and contralateral tibialis muscles. Supramaximal electrical stimuli of 0.2 msec duration were applied to the nerve at 2 Hz for 2 sec every 20 sec for 30 min or 50 Hz (tetanus) once using a Grass S88 stimulator. The evoked single twitch and tetanic tensions of the respective tibialis muscles were recorded via a Grass P122 amplifier and Grass Polyview Software. At the end of functional studies, the muscle tissues were excised, wet weight assessed and then stored at −80 ° C.

Immunohistochemistry of Whole-mount Soleus Muscle to Evaluate Expression of AChR Synapses

Soleus muscles from contralateral and immobilized sides were isolated by dissection of mouse hindlimbs. The muscles were washed in saline to remove blood and then fixed in 4% paraformaldehyde followed by whole-mount immunochemical staining with 1.5 µg/mL Alexa Flour 594-labeled BTX (Molecular Probes, Carlsbad, CA) in PBS for 1 h at 4 °C. After washing with PBS, the muscles were mounted on the slides under the cover slip by pressing with the help of magnet and viewed under the microscope for expression of AChRs. The number of postsynaptic endplates was evaluated by labeling the AChRs with Alexa Flour 594-labeled BTX on the muscle membrane. These synapses were counted by five staff members in the neighboring laboratory. Selection of staff members was based on participant interest, muscle biology background and ability to distinguish and count NMJ numbers. Nuclei in muscle fibers were localized by staining the section with 300nM DAPI for 5min.The counts given by them are indicated as mean ± standard error.

Cross Sections of Soleus Myofibers

For the cross sectional area of soleus myofibers from immobilized and contralateral sides, the muscles were harvested, washed in saline, and placed on to the L-shaped tin plate. The distal end of the muscle was coated with OCT (optimum cutting temperature) and dipped in methyl butane cooled with liquid nitrogen for 10 s. The frozen muscles were immediately transferred to microtube and stored at − 80 ° C until used. At a later time, the frozen muscles previously embedded in OCT were sectioned to 8 microns in thickness, collected on slides and dried at room temperature. The sections on slides were blocked in goat serum in phosphate buffered saline (PBS)-Tween for 30 min followed by overnight incubation at 4 °C with laminin antibody (dilution of 1:1000) to detect muscle membrane surrounding the myofiber. The laminin-antibody-bound sections were washed with PBS and incubated with Rhodamine-conjugated mouse secondary antibody in goat serum at room temperature in dark for 1 h to visualize the laminin signal. Then, they were washed and mounted with Vectashield (mounting medium containing DAPI) (Vector Laboratories, Burlingame, CA) for viewing via fluorescent microscope. Images were acquired by Nikon Eclipse E800 microscope under 5X and 40X objectives (Nikon Inc., Melville, NY). To assess the morphology of soleus muscle fibers from immobilized and contralateral sides, the frozen cross sections of muscles were stained with Hematoxylin and Eosin staining according to manufacturer protocol. ImageJ software was used to measure the cross sectional area of muscle fibers.

Immunoblot Analysis for α7AChR Protein Expression

For immunoblot analysis, frozen muscle samples (n=3) were powdered using hammer and homogenized in tissue lysis buffer (Sigma, St. Louis, MO) containing complete protease inhibitor (Roche, San Francisco, CA). All the extraction procedures were performed in cold room. The crude protein extract was centrifuged at 14000 rpm for 30 min to obtain clear supernatant. The protein samples were normalized using BSA as standard and then 40 µg of protein per lane was fractionated by Nu-PAGE, and blotted onto nitrocellulose membrane (Invitrogen, Carlsbad, CA). The membrane was blocked by 5% nonfat dry milk. Anti-α7AChR and anti- α1AChR (Abcam, Cambridge, MA) at dilution of 1:1000 served as primary antibodies. The membranes were incubated with HRP-conjugated goat anti-rabbit secondary antibodiy for 30 min (dilution of 1:5,000). Specific proteins were detected with chemiluminescence substrate, HyGLO quick spray (Denville Scientific Inc., Metuchen, NJ), and the intensity of protein bands were visualized by exposure of membranes to Kodak X-Omat films (Denville Scientific Inc., Metuchen, NJ).

Identification of Presence or Absence of Muscle Edema

Muscle edema and cellular swelling can be caused by venous compression of the applied EPC and/or immobilization. In order to rule out the muscle edema after hindlimb immobilization by the EPC, the wet and dry muscle masses of soleus, tibialis and gastrocnemius muscles were measured (n=5). To determine the wet weight, the muscles were weighed immediately after harvesting. The same muscles were left in oven at 60 °C to dehydrate overnight and then the dry weights were measured as described previously (Cartee et al. 1996).

Statistical Analyses

Results are presented as mean ± SE. Statistical significance of differences between groups was performed by Student t-tests when comparing two groups. For three or more groups, one-way ANOVA was used and subjected to Tukey's multiple comparison tests. All analyses were performed using Prism 4.0 (GraphPad, San Diego, CA).

Results

The animals in the cage showed no signs of pain, as evidenced by free movement, ate and drank ad libitum because the food pellets and transgel was provided at base of the cage within the easy reach of animals. The animals appeared healthy with no tearing, ruffled fur or withdrawal (isolation). Body weight change at 21 days of immobilization was not significant (23.6 + 0.6 g, n=4, p >0.05) compared to starting body weight (25.2 + 0.5g, n=4, p >0.05). Sham-immobilized animals did not have significant gain in body weight (28.0 + 0.6 g to 28.9 + 0.5 g, n=5, p >0.05) after 21 days. Figure 2A depicts the macroscopic views of gastrocnemius, tibialis and soleus muscles on the immobilized and contralateral sides after harvesting of the tissues after 21 days of immobilization. The immobilized side (right panel) muscles were smaller as compared to contralateral side (left panel). The soleus muscle mass appeared most atrophied after immobilization for 21 days. The entire cross-sectional areas of contralateral and immobilized soleus muscles were 2244 and 1124 mm2, respectively (Fig. 2B). Determination of soleus muscle weight showed the progressive loss of muscle mass of 11 ± 4.9% (p >0.05), 22 ± 5.6% (p <0.05), 39± 6.2% (p <0.05) and 45 +16.7% (p <0.05) at 3, 7, 14 and 21 days of immobilization, respectively, compared to contralateral side. The contralateral muscle mass of soleus when compared to sham-immobilized soleus muscle, whose contralateral side was not immobilized, did not show differences in muscle mass (Fig. 2C) when examined at 21 days. Wet and dry muscle mass changes in soleus, tibialis and gastrocnemius at day 21 are presented in Figs. 3A and B. The muscle masses in soleus, tibialis and gastrocnemius were significantly decreased approximately by 45%, 15% and 20%, respectively, compared to their respective contralateral side at 21 days. During the immobilization, no paw edema, ischemia or mortality was externally visible. The wet to dry muscle weight ratio did not differ between immobilized and contralateral side for all three muscles (Fig. 3C). The evoked single and tetanic muscle tension responses in tibialis muscle were significantly decreased at 21 days on the immobilized side compared to the contralateral side; the percentage reduction of single twitch and tetanic tensions were 25 and 26%, respectively, on the immobilized side compared to contralateral side (Fig. 3D).

Fig. 2. Hindlimb immobilization by EPC and atrophy of muscles.

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A. Macroscopic views and muscle weights from immobilized (Immob) and contralateral (Contra) sides. Macroscopically, the muscles appeared smaller after immobilization. B. Microscopic analysis of immobilized (Immob) and contralateral (Contra) soleus muscles using Hematoxylin and Eosin staining. The cross-sectional area of whole soleus muscle measured by ImageJ indicated a reduction of ~50% after immobilization compared to contralateral side. Magnification 5X. C. Determination of soleus muscle weight showed the progressive loss of muscle mass on immobilized (Immob) side, at 3 (n=4, p >0.05), 7 (n=4, p <0.05), 14 (n=5, p <0.05) and 21 (n=4, p <0.05) days as compared to contralateral (Contra) side. The soleus muscle mass in sham-immobilized mice, not immobilized on the contralateral side, was used as control for 21 day immobilization group. The percentage of contralateral (Contra) side assumed as 100%. The muscle mass of soleus of contralateral side of immobilized and non-immobilized limb were not different. The data represent the mean ± standard error. Statistical analysis was done using one-way ANOVA with Tukey's multiple comparison tests. Asterisks represent p value (*p ≤0.05), as compared to contralateral side. NS represents not significant. Immobilization induced progressive loss of skeletal muscle mass.

Fig. 3. Wet and dry weights of muscles and evoked muscle tensions.

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Wet (A) and dry (B) weights of soleus (left panel), tibialis (middle panel) and gastrocnemius (right panel) muscles. The muscle weights were normalized to body weight (mg/kg). C. Wet to dry weight ratio of soleus, tibialis and gastrocnemius muscles (n=5). The muscles did not show edematous tissues after immobilization. D. Nerve evoked single twitch and tetanic tensions were significantly reduced on the immobilized (Immob) side, shown as percentage of contralateral (Contra) side assumed as 100%. The data represent the mean ± standard error (n=5). Statistical analysis was done using Student t-test or one-way ANOVA with Tukey's multiple comparison tests. Asterisks represent p values (*p ≤ 0.05) as compared to contralateral side. EPC induced muscle weakness on immobilized side.

We next examined the synapses on muscle membrane by labeling the AChRs with fluorescent α-bungarotoxin in soleus muscle at 21 days of immobilization. The density of synapses on muscle membrane per unit area was increased in soleus muscle on immobilized side (Fig. 4A); the α-bungarotoxin labeled synapses on immobilized side were 818 + 31/mm,2 and 433+ 16/mm2 (p <0.01) on the contralateral side of the soleus muscle. The comparison of individual end-plates between immobilized and contralateral sides of soleus, tibialis and gastrocnemius did not reveal any specific alterations, except for increased convolutions of the usual pretzel shaped junctional receptors on immobilized side (Fig. 4B). Next, the cross sectional areas of myofibers on the immobilized and contralateral sides were examined. Examination of the laminin-stained cell membrane indicated the cross sectional areas of myofibers on immobilized side appeared smaller compared to contralateral side (Fig. 4C). In order to quantitate the atrophy, each muscle was numbered. It is notable that for a given area, there were more muscle fibers on the immobilized compared to contralateral side (Fig. 4D). The number of muscle fibers in atrophied (immobilized) and contralateral muscles were 117 and 83/mm2, respectively. The nuclei in the muscle fibers were localized using DAPI staining and showed the usual peripheral distribution along the cell membrane. Analyses of average cross-sectional area of each soleus muscle fibers of contralateral and immobilized side revealed that the cross-sectional area reduced from 0.88 + 0.04 to 0.57 + 0.04 mm2 in atrophied muslce as shown in Fig.4E.

Fig. 4. Muscle synapses and muscle fiber size.

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A. AChRs were stained using AlexaFlour 594 labeled-BTX in whole mount soleus muscle following immobilization for 21 days. The number of synapses per unit area on contralateral (Contra) and immobilized (Immob) sides counted at 5X magnification by five individuals indicated 433 + 16 and 818 + 31/mm2 synapses, respectively. Muscle synapses were increased due to atrophied muscle fibers on immobilized side compared to contralateral side. B. Morphological comparison of end-plates between immobilized (Immob) and contralateral (Contra) sides. The end-plates of soleus (top panel), tibialis (middle panel) and gastrocnemius (bottom panel) were stained using AlexaFlour 594 labeled-BTX. Endplates were more convoluted on immobilized side compared to contralateral side. C. Cross-sectional area of muscle fibers on immobilized (Immob) and contralateral (Contra) sides. The muscle membrane was stained using laminin specific antibody (top panel), which defined the margins of the muscle fiber, and the nuclei in the muscle fibers were localized with DAPI staining (middle panel) respectively, with merge image (bottom panel). Size of muscle fiber was reduced on immobilized side compared to contralateral side. D. Comparison of number of muscle fibers on contralateral side, Contra, (left panel) and immobilized side, Immob, (right panel). Laminin staining (upper panel). Each number represents one muscle fiber. The number of muscle fibers per unit area on contralateral (Contra) and immobilized (Immob) sides were 83 and 117/mm2, respectively. The higher number of muscle fibers per unit area indicates the atrophic muscle compared to contralateral side. H and E staining (lower panel). Arrows indicate the atrophied muscle fibers. Number of muscle fibers were decreased per unit area on immobilized side compared to contralateral side. E. Measurement of average cross-sectional area of soleus muscle fibers on contralateral side, Contra, (left panel) and immobilized side, Immob, (right panel) using ImageJ software. The average cross-sectional area of muscle fibers on contralateral (Contra) and immobilized (Immob) sides were 0.88 + 0.04 and 0.57 + 0.04 mm2, respectively. The data represent the mean ± standard error. Total 80 randomnly selected muscle fibers were counted from each side. Statistical analysis was done using Student t-test. Asterisks represent p values (*p < 0.05) as compared to contralateral side. Average muscle fiber cross-sectional area was reduced on immobilized side compared to contralateral side.

In view of the presence of α7AChR in the fetus and following denervation, we also examined its expression during disuse by immunoblot analysis of soleus muscle. The immunoblot showed increased expression of α7AChRs on the immobilized compared to contralateral side (Fig 5A); mouse brain extract and PC12 cells were used as positive controls. Densitometric analysis of α7AChRs showed about a three fold increase in its expression in muscle on the immobilized side compared to contralateral side (p <0.01) (Fig. 5B). Previous studies following immobilization have documented the upregulation of α1 subunit of AChRs (Ibebunjo and Martyn 1999; Yanez and Martyn 1996). Therefore, we examined α1AChR subunit expression by immunoblot. The α1AChR expression was also increased on immobilized side compared to contralateral side (Fig. 5C).

Fig. 5. Expression of α7AChRs in soleus muscle.

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A. Western blot analysis using α7AChR specific antibody revealed increased expression of α7AChRs in soleus muscles. Mouse brain extract (MBE) and PC12 cells were used as positive control for α7AChR. Contralateral (Contra) and immobilized (Immob) sides at 21 days of immobilization. B. Densitometric analysis of α7AChRs. The data represent the mean ± standard error (n=3). Statistical analysis was done using Student t-test. Asterisks represent p values (*p ≤ 0.05) as compared to contralateral side. C. Western blot analysis of α1AChR subunit in soleus muscles. Contralateral (Contra) and immobilized (Immob) sides at 21 days of immobilization. Immobilization increased expression of α1- and α7-AChR subunits.

Discussion

Muscle wasting associated with critical illness with and without prolonged hospitalization in bed leads to decreased ventilation, dependence on respirators, prolonged rehabilitation and increased mortality (Douglas et al. 2002; Fletcher et al. 2003; Sprague and Hopkins 2003). Any form of disuse including casting, even in the absence of systemic disease, can lead to atrophy. Therefore, it is important to understand the molecular mechanisms of disuse muscle atrophy in order to prevent its long term consequences on morbidity and mortality. Previous methods used to study disuse muscle atrophy have been useful (Damrongrungruang et al. 2004; Bajusz 1964; Frimel et al. 2005; Fink et al. 2008; Mokhtarian et al. 1999; Caron et al. 2009; Kandarian et al. 1996), but these methods do not exactly reproduce clinical events seen during simple immobilization of limb. Immobilization and loss of weight bearing occurs with heavy sedation, application of heavy bandages or casting including spica. Previous models of immobilization required repetitive re-application of the cast with time (Frimel et al. 2005) or surgical immobilization (Fink et al. 2008; Mokhtarian et al. 1999; Caron et al. 2009). The latter, in and of itself causes local muscle inflammation, which can confound the atrophic signaling pathways. The method of hindlimb immobilization described in this report mimics the disuse seen clinically with heavy sedation, or casting in the absence of systemic disease. The scenario is seen, for example, in neuropsychiatric disorders, where often no systemic diseases is present or during application of casts for varus or valgus deformities or fractures.

Our study documents progressive loss of muscle mass with time and decrease of single twitch and tetanic tensions, the latter reflecting repetitive work. There was a 45% loss in soleus muscle mass and a 25% decrease in tibialis tension by 21 days of immobilization. This method of immobilization showed atrophy in both fast (tibialis and gastrocnemius) and slow muscles..

In our recent study using EPC method of immobilization, we have shown a significant upregulation of MURF1 and atrogin mRNAs on the immobilized side compared with the contralateral side (Zhu et al. 2012). In the current study, laminin staining of the muscle cell wall and H and E staining of the muscle fibers revealed decrease in cross sectional area with atrophied muscle fibers and peripheral distribution of the nuclei. A notable novel finding is the upregulation of α7AChR on the muscle membrane. During the 21 days of immobilization, the EPC did not have re-applied as in other methods of casting.

In the normal innervated weight bearing contracting muscle, the AChRs are expressed in the junctional area only. These AChRs spread and increase throughout the muscle membrane after denervation (Yanez and Martyn 1996; Mishina et al. 1986). After complete denervation, end-plates in adult soleus muscle fibers break down into smaller clusters due to the loss of connectivity (nerve terminals) between nerve and muscle (Huze et al. 2009). In previous studies, both denervation and immobilization lead to loss of muscle mass and increase in total AChRs on the muscle membrane (Yanez and Martyn 1996; Tsuneki et al. 2003). In contrast to denervation, after immobilization the end-plates maintained the typical pretzel shape, similar to contralateral side, but appeared more convoluted and distorted most likely due to the atrophy of the muscle fibers. The significant increase (1.7 fold) in numbers of synapses (per unit area) on muscle membrane assessed by labeled α-bungarotoxin, which binds to mature, immature and α7AChRs may reflect the atrophy bringing the end-plates closer together. Our western blot analysis of the soleus muscle fibers, however, revealed that the α7AChRs was upregulated by three fold along with α1AChRs on the immobilized side as compared to contralateral side after 21 days of immobilization. Thus, this study provides evidence for the first time that α7AChRs are expressed in non-denervated adult muscle and these α7AChRs are post-synaptic. The expression of α7AChRs, however, has been noted in the newborn and denervated muscle (Fischer et al. 1999; Tsuneki et al. 2003). The presence of α7AChRs in non-neuronal tissues has received increased attention recently. Stimulation of α7AChRs leads to modulation of cytokine release (Khan et al. 2012). The presence of α7AChRs can also lead to resistance to non-polarizing relaxants as shown following denervation (Tsuneki et al. 2003). The clinical implications of the role of increased expression of α7AChRs in resistance to non-depolarizing relaxants seen clinically during immobilization (Ibebunjo and Martyn 1999; Fink et al. 2008) needs further study.

If there is pressure on venous or lymphatic systems by the EPC and/or padding, this can lead to edema of muscle tissues. Wet to dry weight ratios of soleus, tibialis and gastrocnemius muscles ruled out the possibility that immobilization with EPC caused tissue edema. The body weight loss, which is amounted to ~1–1.6 g over 14 and/or 21days, was not significant compared to starting weight. Even in other form of hindlimb immobilization by external fixation, it has been shown that there was no significant difference between the body weights of control and immobilized groups after four weeks (Zarzhevsky et al. 2001). Sham-immobilized animals gain a little body weight (~0.5 –1 g) but not significant. Thus, the immobilization procedure did not affect the ability of the animal to consume food and fluid.

For our studies at 7 and 14 days, we used the contralateral side as control, while at 21 days, in addition to contralateral side, we had separate naïve animals as additional controls. One might presume that immobilization of one limb will lead to overuse hypertrophy of the other limb. Comparison of the muscle masses of contralateral leg to immobilized naïve legs indicate no differences in muscle masses at day 21. In other words, there is no hypertrophy of the contralateral side when one side is immobilized. This observation is consistent with previous studies, which examined muscle changes at earlier times including 7 and 14 days (Ibebunjo and Martyn 1999; Ibebunjo et al. 1997; Fink et al. 2008; Zhu et al. 2012).

Conclusion

A non-surgical, non-invasive model closer to the clinical scenario seen in some forms of human immobilization in the absence of systemicl illness is described. The method is simple, quick and painless compare to existing methods. The model described can be used to study the molecular mechanisms of immobilization-induced muscle atrophy and also test therapeutic measures to correct it. We document the upregulation of α7AChRs on the muscle membrane similar to that seen in denervation. The functional and pathophysiological role of α7AChRs requires further study.

Aknowledgements

This work was supported grants from the NIH RO1-055082 and P50-25000 project IV and from Shriners Hospitals for Children® Research Philanthropy to JAJM. We thank Dr. Hui Zheng for his advise on statistical analyses.

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