Vibration training has emerged as a therapeutic strategy to improve the musculoskeletal system in health and rehabilitation. In health, it is used as an exercise modality to better athletic performance (reviewed by Wilcock et al. ). In rehabilitation, bone has been the primary focus because it has been shown to be osteogenic (32). Vibration training is also being explored as an alternative exercise modality for conditions of muscle weakness, such as in geriatrics and with bed rest. The results of these studies have been largely mixed. For example, a meta-analysis showed that whole-body vibration training was effective in improving leg muscle strength in older adults (14), whereas another analysis concluded that there is only weak support for the efficacy of whole-body vibration for muscle strength (17). Similarly, some but not all studies on young healthy adults show that vibration training improves muscle function (reviewed by Rittweger ). Thus, the broader application of vibration training to musculoskeletal health is limited because its effects on skeletal muscles are not unanimous.
One aspect that likely contributes to the various results of vibration training on skeletal muscle function is the magnitude of acceleration or the intensity of the vibration. This parameter is typically expressed as a fraction of gravity (g) and it is a primary factor that dictates the mechanical signal delivered from the oscillating platform to the subject standing on the device. Most commercially available, whole-body vibration equipment delivers accelerations >1.0g, called high-intensity vibration in this article. Vibration platforms that deliver accelerations ≤1.0g (low intensity) have also been investigated. In terms of effects on skeletal muscle, it has been shown that 2–12 months of low-intensity vibration training results in increased muscle strength (19), balance (19), grip strength (25), and muscle mass (11,24). Subjects in those studies were selected based on clinical conditions associated with poor bone health, and it is possible that muscle weakness was also a characteristic of those subjects. Thus, it is not clear if low-intensity vibration training has the potential to enhance skeletal muscle of healthy individuals without muscle weakness.
Low-intensity vibration training is used to investigate osteogenic effects in animal models including sheep, rats, and mice (e.g., [28,29,36]), but myogenic effects have been much less studied. In two notable studies, BALB/c mice were subjected to low-intensity vibration training for a duration of 6 wk. Xie et al. (37) reported that cross-sectional areas of soleus muscle and type I and II fibers within that muscle were greater in vibrated than control mice. However, Murfee et al. (20) reported low-intensity vibration reduced the number of arterioles and venules in the distal region of soleus muscle, an undesirable microvascular adaptation. In addition, others have reported that high-intensity vibration causes injury to rodent muscle as indicated by fiber swelling and centrally located nuclei (21,22). However, none of these studies evaluated the extent to which muscle function was beneficially or detrimentally altered with vibration training. Thus, a more comprehensive analysis of skeletal muscle after vibration training, particularly low-intensity vibration training, is needed to determine function adaptations. As such, the primary objective of this study was to use a low-intensity vibration platform, designed specifically for mice to test the hypothesis that contractility is improved in muscles of the hindlimb in response to vibration training. A subset of mice housed in small cages to evoke physical inactivity was also studied to determine whether vibration training was more effective under conditions promoting muscle weakness.
Traditional exercise training can elicit changes in skeletal muscle strength, oxidative capacity, and fiber-type distributions, which in turn can affect muscle’s resistance to fatigue. There are indications that vibration training may influence muscle fatigue (15,27). Therefore, histological analyses reflecting oxidative capacity, capillarity, and fiber types as well as functional analyses of muscle fatigability and recovery from fatigue were assessed to determine whether low-intensity vibration training can provide a strong enough stimulus to evoke such cellular and parallel functional adaptations in muscles of mice.
Animal and study design.
Male C57BL/6J mice age 8 wk were housed at 20°C–23°C on a 12-h light–12-h dark cycle in a facility accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International. Mice were provided food ad libitum, and intake was recorded weekly. Mice were randomized to one of four conditions, either without or with vibration treatment (sham and vibrated, respectively) and either housed in traditional sized cages to permit normal cage activity (Sham-Active, n = 8; Vibrated-Active, n = 10) or small cages that restricted physical activity (Sham-Restricted, n = 8; Vibrated-Restricted, n = 8). Sample size was determined a priori from our past experience using C57BL/6 mice and physiological muscle outcome measurements in our laboratory. Specifically, the number of mice used was based on that needed to detect a 10% difference among groups in strength assuming a minimum power of 0.7 and an α-level of 0.05. Mice in restricted groups were placed individually in small cages as reported previously (23), except that in the current study, a ceiling was also used to minimize vertical movement (length × width × height: 12 × 8.5 × 6.3 cm). This small-cage intervention reduces ambulation by ∼85% (23). Mice in traditional cages were housed three per cage. All mice received vibration or sham treatment 5 d·wk−1 for a total of 6 wk.
After 6 wk of treatment, when mice were 14 wk of age, contractile functions were analyzed for the anterior and posterior crural muscle groups of the left hindlimb and the soleus muscle of the right hindlimb. These muscles and muscle groups were selected because they are near the vibrating platform and transmissibility of the vibration signal is not damped by soft tissue in the mouse hindlimb (9). For the in vivo testing of the anterior and then posterior crural muscles, mice were anesthetized with fentanyl citrate (0.2 mg·kg−1 body mass [BM]), droperidol (10 mg·kg−1 BM) and diazepam (5 mg·kg−1 BM). Immediately after in vivo analyses, mice were injected i.p. with sodium pentobarbital (50 mg·kg−1 BM) and muscles were excised. First, the soleus muscle was dissected for ex vivo contractile analyses and then tibialis anterior (TA), extensor digitorum longus (EDL), gastrocnemius and contralateral soleus muscles were dissected, weighed, and either mounted in optimal cutting temperature compound or frozen in LN2. Epididymal fat pads were excised and weighed as well. Mice were then killed with an overdose of sodium pentobarbital (200 mg·kg−1 BM). All procedures were approved by the Institutional Animal Care and Use Committee at the University of Minnesota and adhere to the American College of Sports Medicine animal care and standards. All investigators were blinded to treatment groups while performing analyses of contractile function and dissections.
Mice in the vibration groups were exposed to low-intensity vibration via a vertically oscillating platform. The vibration system was designed after the work of Fritton et al. (10), with modifications including the use of a cylindrical platform driven by accelerometer feedback to ensure fidelity, to minimize error, and to eliminate wave-transmission modes to create a uniform exposure across the entire platform. Briefly, a circular platform was driven by a linear actuator (BEI, Kimco; Magnetics, San Marcos, CA) using custom-built LabVIEW software (National Instruments, Austin, TX). The frequency and acceleration of vibration were continually monitored and fed back into the system using an accelerometer (50g; Endevco, San Juan Capistrano, CA) mounted to the underside of the aluminum platform. During vibration, mice were temporarily housed within one of four compartments of an acrylic cage fixed to the top of the platform. To maximize the exposure of the vibratory stimulus to the mouse hindlimb, a ceiling height of 6.3 cm was used to minimize vertical activities such as rearing and jumping and to be consistent with housing of the restricted mice. The vibration stimulus was applied for 15 min, 5 d·wk−1 for 6 wk using a vibration frequency of 45 Hz and an acceleration magnitude of 1.0g. These parameters created a peak-to-peak displacement of 0.24 mm. Sham mice were placed in an identical acrylic cage for the same duration of time as Vibrated mice but did not receive any vibration stimulus.
In vivo measurements of anterior and posterior crural muscle contractility.
The anterior (TA, EDL, and extensor hallucis longus muscles) and then posterior (soleus, plantaris, and gastrocnemius muscles) crural muscle groups were tested for contractile capacities in anesthetized mice using methods previously described (4,13). Contraction of the anterior crural muscles was elicited via percutaneous stimulation by platinum-iridium needles (Model E2-12; Grass Technologies, West Warwick, RI) placed on either side of the left common peroneal nerve. Peak-isometric torque was defined as the greatest torque measured by a 300B-LR servomotor (Aurora Scientific, Aurora, Ontario, Canada) during a 200-ms stimulation using 1-ms square-wave pulses at 300 Hz and increasing voltage from 3.0 to 9.0 V (models S48 and SIU5; Grass Technologies). The fatigability of the anterior crural muscles was then assessed by subjecting the muscles to 120 submaximal isometric contractions for 2 min using 330 ms stimulations at 50 Hz. A rest period of 5 min followed, and then recovery from fatigue was tested by remeasuring peak isometric torque. Immediately after the testing of the anterior crural muscles, the left common peroneal nerve was severed, and the testing of the posterior crural muscles began. The contraction of this muscle group was elicited by stimulating the sciatic nerve (7). Peak-isometric torque was determined during a 200-ms stimulation with varying voltages (3.0–7.0 V) at 300 Hz using 1-ms square-wave pulses. Fatigability was assessed by 120 isometric contractions produced for a 2-min duration using 500-ms stimulations at 60 Hz. After 5 min of rest, recovery from fatigue was tested by remeasuring peak isometric torque of the posterior crural muscles. Isometric torque–time tracings from the peak prefatigue contractions were analyzed to determine maximal rates of contraction and relaxation for both anterior and posterior muscle groups.
Ex vivo soleus muscle contractility.
Isolated soleus muscles were mounted to a dual-mode muscle lever system (300C-LR; Aurora Scientific) in a 1.5-mL bath containing oxygenated (95% O2) Krebs–Ringer bicarbonate buffer at 25°C using a 5-0 suture. Muscle lengths from proximal to distal myotendinous junctions were measured using digital calipers after muscles were set to their anatomic resting length (Lo). Contractile characteristics were measured as described previously (18) in the order of passive stiffness, maximal isometric tetanic force (Po), and active stiffness. Tetanic force-time tracings were analyzed to determine maximal rates of contraction and relaxation. Following these measurements, fatigability was tested by employing a protocol of sixty 1-s tetanic contractions at a rate of 12 tetani per minute for 5 min and then remeasuring Po 5 and 10 min later to assess recovery (12). The soleus muscles were then removed from the bath assembly, trimmed, blotted, weighed, and frozen in liquid nitrogen. Soleus muscle mass and Lo were used to calculate physiological cross-sectional area, which was then used for calculating specific Po (5,34).
To determine whether greater rates of relaxation with vibration could be attributed to enhanced calcium handling, we quantified the expression of two calcium sequestering proteins (6). Briefly, frozen TA muscles were pulverized by mortar and pestle, solubilized in 1% sodium dodecyl sulfate with 5 mM ethyleneglycotetraacetic acid and a cocktail of protease inhibitors, and measured for total protein concentration using a NanoDrop ND-100 spectrophotometer (Thermo Fisher Scientific, Wilmington, DE). Proteins were separated on 4%–20% poly-acrylamide gels (120 V for 90 min), transferred to polyvinylidene difluoride membranes, and then probed using antibodies against sarcoplasmic reticulum calcium ATPase (SERCA) 1a (1:1000 dilution of MA3-911; Pierce, Rockford, IL), parvalbumin (1:5000 dilution of PARV-19; Sigma, St. Louis, MO), and glyceraldehyde 3-phosphate dehydrogenase (1:5000 dilution of G9545; Sigma). Secondary antibodies were diluted (1:10,000), and detected and analyzed with the Odyssey Infrared Imaging System (Li-Cor Biosciences, Lincoln, NE) using the 700- and 800-nm channels.
Ten-micron-thick transverse sections were cut from the midbelly of TA and soleus muscles on a cryostat at −20°C. Muscles used for histology were contralateral to those tested in vivo or ex vivo for contractility. Sections were stained for nicotinamide adenine dinucleotide (NADH)–tetrazolium reductase reactivity as an indicator of mitochondrial enzyme activity (1) and by periodic acid–Schiff reaction to determine the number of capillaries per fiber (2). For each stain, three to five muscles per group were evaluated, with ∼300 fibers per muscle assessed for NADH reactivity and ∼200 fibers per muscle for capillarity. Lastly, fiber-type distribution based on myosin heavy chain isoforms were determined and classified as types I, 2a, or 2b using antibodies BA-F8, SC-71, and BF-F3, respectively (as originally developed by Schiaffino and obtained from the Developmental Studies Hybridoma Bank under the auspices of the National Institute of Child Health and Human Development and maintained by Department of Biology, University of Iowa, Iowa City, IA) and immunofluorescent secondary antibodies (3). If a fiber did not react to antibodies against any of these myosin heavy chain isoforms, it was classified as a 2× fiber. Three to five muscles per group were assessed for fiber type, with all fibers of each muscle being evaluated (range, 354–661 fibers for soleus muscle and 2363–3243 fibers for TA muscle). Data were calculated as the percentage of NADH-positive fibers, the average number of capillaries per fiber, the percentage of fiber type, and the average fiber cross-sectional area per fiber type for each muscle analyzed. Those muscle means were then used for statistical analyses. Investigators were blinded to treatment groups while performing all histological analyses.
Data were analyzed using two-way ANOVAs to determine the effects of vibration (sham vs vibrated) and physical inactivity (active vs restricted). Holmes–Sidak post hoc tests were performed when significant interactions were detected. Significance was accepted at P ≤ 0.05. Statistical analyses were carried out using SigmaStat (version 3.5; Systat Software Inc., Chicago, IL).
Six weeks of low-intensity vibration was well tolerated by mice. There were no detectable changes in behavior when vibration was initiated or during the 15-min exposure. Also, Vibrated and Sham mice behaved similarly in the vibration and sham chambers, respectively. The aggressive behavior of the Vibrated-Active mice was noted for ∼5 min when the mice were first returned to their cages immediately after vibration. Food intake during the first week of the study was 13%–26% greater for Vibrated-Active and Sham-Restricted mice compared with Sham-Active mice (Table 1). There was no lasting effect of vibration on food intake, but at week 6, restricted mice consumed ∼15% more food per day than active mice (Table 1).
BM did not significantly differ among groups at the beginning or end of the study (Table 1). The mass of the epididymal fat pad was not affected by vibration or activity; however, fat pad mass relative to BM was lower in Vibrated mice (Table 1). There were trends for Restricted mice to have smaller TA and soleus muscle masses compared with Active mice, but there was no effect of vibration on any hindlimb muscle mass measured (Fig. 1). Furthermore, the length and the physiological cross-sectional area of soleus muscles and protein content of TA muscles were not significantly affected by vibration (Table 1). Collectively, there was no indication that vibration induced hypertrophy of hindlimb muscles.
There were some indications that vibration training improved muscle contractility. Maximal isometric torque generated by anterior crural muscles was ∼10% greater in Vibrated compared with Sham mice (Fig. 2A) and occurred whether mice were active or restricted (interaction P = 0.642). As expected, there was also a main effect of activity with Restricted mice having ∼10% lower maximal torque than Active mice (Fig. 2A). When normalized by mass of the TA muscle, the anterior crural muscle torque of vibrated mice was ∼8% greater than that of Sham mice (57.7 ± 1.2 vs 53.3 ± 1.2 N·mm·g−1, respectively, P = 0.021), with no effect of activity (P = 0.138). The maximal rate of contraction by the dorsiflexors was unaffected by vibration training, but the maximal rate of relaxation after isometric contraction was ∼16% faster in dorsiflexors of Vibrated compared with Sham mice (Fig. 2A). To determine whether the vibration-induced increase in muscle relaxation was due to greater levels of calcium-handling proteins, the expression of SERCA and parvalbumin was measured; neither of the proteins were different between TA muscles from Vibrated and Sham mice (P ≥ 0.361).
In contrast to the anterior crural muscles, the contractility of the posterior crural muscles was largely unaffected by vibration and activity (Fig. 2B). The exception was that Vibrated-Restricted mice had greater rates of contraction by the plantarflexors than Vibrated-Active and Sham-Restricted mice.
There was a trend for maximal isometric tetanic force (Po) by isolated soleus muscles to be greater in Vibrated than Sham mice (P = 0.057; Fig. 2C), regardless of activity (interaction P = 0.887). As expected, Restricted mice had lower soleus muscle Po than did Active mice (P = 0.042; Fig. 2C). Specific Po was not affected by vibration training or activity (P ≥ 0.244), and averaged 19.71 ± 0.79 N·cm−2 across all groups. Similarly, active stiffness was not different among groups (P ≥ 0.245; 252 ± 9.7 N·m−1 for all groups). The maximal rate of contraction tended to be faster, and the maximal rate of relaxation was ∼20% faster in soleus muscles of vibrated than sham mice (Fig. 2C). There were main effects of activity on rates of contraction and relaxation with soleus muscles from Restricted mice being slower than those from active mice. There was no effect of vibration or activity on soleus muscle passive stiffness (P ≥ 0.689; 11.59 ± 0.4 N·m−1 across all groups).
Muscle fatigability, as measured by percent loss of torque or force over the course of repeated contractions, and recovery from fatigue were not affected by vibration training in any of the three muscles or muscle groups studied. The torque of the anterior crural muscles was reduced by ∼80% at the end of the 2-min bout of fatiguing contractions with no effect of vibration or activity (P ≥ 0.431; Fig. 3). Five minutes later, torque generation was equally recovered, irrespective of vibration or activity (P ≥ 0.403; Fig. 3). Similarly, posterior crural muscles lost ∼70% of torque during the bout of fatiguing contractions with no effect of vibration or activity on torque loss (P ≥ 0.126) or recovery of torque 5 min later (P ≥ 0.388). The 60 isometric contractions performed in 5 min by isolated soleus muscles caused force to go down ∼55% regardless if mice had vibration trained or not, or if mice were active or restricted (P ≥ 0.464). By 10 min after the fatiguing protocol, soleus muscles recovered to ∼95% of prefatigue Po with no difference among groups (P ≥ 0.228).
Vibration training had no effect of TA muscle fiber NADH reactivity, capillarity, or fiber-type composition (Table 2). Mice that were restricted in their activity had TA muscles with relatively fewer NADH-positive fibers, fewer type 2x fibers, and more type 2b fibers compared with those of normally active mice. Soleus muscles from Vibrated and Sham mice were not different in NADH reactivity, fiber-type composition, or fiber cross-sectional areas (Table 2 and Fig. 4). Capillarity, as determined by the number of capillaries per fiber, was 24% greater in vibrated soleus muscles, but only within the Restricted mice (Table 2).
The results of this study partially support the hypothesis that low-intensity vibration improves muscle contractility. We show that standing on a platform that vibrates at 1.0g and 45 Hz for 15 min·d−1, five times per week for 6 wk, increased strength and rate of relaxation of some but not all of the mouse hindlimb muscles tested (Fig. 2). These vibration-induced improvements in muscle contractility were independent of mouse activity. That is, the restriction of physical activity in subsets of mice successfully induced muscle weakness of the anterior crural muscles and soleus muscle, but that was not required for vibration to be effective in improving strength or relaxation.
The mechanism by which vibration improves strength is not clear. Acute vibration may facilitate contractility by enhancement of stretch and H-reflexes (26) or increased muscle activation (8). Neuromuscular adaptations in response to weeks or months of vibration training are less clear. One way that traditional resistance exercise results in strength gains is by muscle hypertrophy, but our results do not give any indication that hypertrophy was induced by 6 wk of low-intensity vibration training in C57BL/6 mice. Above all, hindlimb muscle masses and protein contents were not greater in Vibrated compared with Sham mice (Fig. 1, Table 1). Physiological cross-sectional area and length of soleus muscles (Table 1) as well as cross-sectional areas of fibers from that muscle (Table 2), were not affected by vibration training, providing further support that hypertrophy was not induced. Two previous studies on BALB/c mice subjected to 15 min·d−1 of low-intensity vibration for 6 wk report data on soleus muscle and fiber size. Murfee et al. (20) found that the soleus muscle area and the number of fibers per area were not altered by vibration training, and Xie et al. reported that the number of type I and type II muscle fibers and the cross-sectional area of those fibers were not different between soleus muscles of sham and vibrated mice (36). However, in the latter study, total cross-sectional area of soleus muscle was calculated to be 27% greater in mice that were subjected to vibration training (36). The differing soleus muscle cross-sectional area results between that study and ours may have to do with the method by which cross-sectional area was determined (anatomical vs physiological) or is possibly due to differential responses to vibration. That is, the designs of the two studies were the same except that we used an acceleration of 1.0g and they used 0.3g. Overall, there is no convincing evidence that low-intensity vibration training causes hypertrophy in hindlimb muscles of mice.
Improvement in the intrinsic ability of muscle to generate torque or force is another mechanism by which strength gains can be realized. The maximal torque generation by the anterior crural muscles of mice that were subjected to vibration training was ∼10% greater than that of nonvibrated mice (Fig. 2A). Anterior crural muscle torque normalized by TA muscle mass was ∼8% greater in Vibrated than Sham mice, suggesting that some intrinsic ability of the TA muscle to generate torque was affected by vibration training. Specific force and active stiffness measurements of soleus muscle do not support this supposition; however, as these indicators of muscle quality were not different between Vibrated and Sham mice, Po trended toward being greater in Vibrated mice (P = 0.057; Fig. 2C). Collectively, these data leave some ambiguity as how muscle strength is improved by vibration training. Given that the anterior as opposed to the posterior crural muscles were affected by vibration, it would have been interesting to analyze the contractility of the EDL muscle and should be performed in future studies. No other studies that we are aware of have measured muscle function after vibration training in mice. We suggest that more functional types of studies need to be conducted to firmly establish the extent to which low-intensity vibration training improves muscle strength and furthermore to determine underlying mechanism of strength gains when they occur. In addition, investigation is needed into the response of different muscles and muscle groups. The anterior and posterior muscle groups analyzed in this study, while theoretically receiving similar mechanical stimuli from the vibrating platform, responded very differently in terms of contractility adaptations with those measured in the posterior muscle group being minimally affected (Fig. 2B).
The maximal rate of muscle relaxation was improved by vibration in two of three muscles or muscle groups studied. Specifically, the maximal rates of relaxation after maximal isometric contraction by anterior crural and soleus muscles were 16%–20% greater in Vibrated than Sham mice (Fig. 2A). Muscle proteins related to calcium handling were investigated in an attempt to determine possible mechanisms underlying the faster muscle relaxation, but expression levels of parvalbumin and SERCA were not different in TA muscles of Vibrated and Sham mice. Analyses of the content of these two proteins do not exclude the possibility of vibration-induced influences of calcium handling because structure or function of these or other calcium handling proteins could have occurred. To our knowledge, the influence of vibration training on calcium handling proteins and muscle relaxation has not been previously studied. An example indicating that vibration training has the potential to influence particular muscle proteins is a study showing that ryanodine receptor type 1 expression and function in soleus muscles of bed-rested humans is significantly affected by vibration training (31). Again, more investigations are required to determine the extent to which vibration training affects muscle contractility parameters such as rates of contraction and relaxation and the mechanisms underlying any such adaptations. Using mouse models and strategies that bypass activation of muscle through the nerve and in response to reflexes may help to identify neural versus muscular adaptation to vibration training.
The fatigability of hindlimb muscles and the recovery from fatigue were not affected by low-intensity vibration training (e.g., Fig. 3). Corresponding with these functional outcomes, there was no indication that vibration affected mitochondrial oxidative capacity in TA or soleus muscle as measured by the percentage of fibers stained for NADH reactivity (Table 2 and Fig. 4). Similarly, fiber-type compositions of TA and soleus muscles were not altered in response to vibration training (Table 2). Previous studies of low-intensity vibration on mouse soleus muscle also found no fiber-type transition (20,37). Finally, capillarity was investigated because blood supply to contracting muscle is an important factor in fatigability, and a previous study reported the suppression of blood vessels in mouse soleus muscle in response to low-intensity vibration (20). In the TA muscle, there was no difference between Vibrated and Sham mice in terms of the number of capillaries per muscle fiber (Table 2). In normally active mice, there was also no difference in soleus muscle capillarity; however, among the physically restricted mice, those that were subjected to vibration had 24% more capillaries per fiber than those from sham mice (Table 2). These data indicate a beneficial vascular remodeling in response to low-intensity vibration. Collectively, functional measurements of muscle fatigue and histological analyses related to muscles’ ability to resist and recover from fatigue, apart from soleus muscle capillarity, were not improved by low-intensity vibration training.
An interesting finding of the present study was that the epididymal fat pad mass, when expressed relative to BM, was ∼12% lower in Vibrated than Sham mice (Table 1), agreeing with previous reports of reduced fat masses in mice in response to vibration training (16,30). There is evidence from mouse and cell culture studies that low- and high-intensity vibration can influence adipogenesis and perhaps shift bone marrow cell fate away from adipocytes toward osteoblasts, which could contribute to vibration-induced loss of fat and gain of bone, respectively (16,30,33).
In conclusion, we show some evidence supporting the hypothesis that low-intensity vibration training improves muscle contractility, specifically strength and relaxation rates. The cellular and molecular adaptations that occurred to cause these physiological improvements were not elucidated. It is important to recognize that mechanisms underlying the remodeling of muscle in response to low-intensity vibration training as used in this study may or may not be the same as those that underlie responses to high-intensity vibration. We specifically chose to focus on low-intensity vibration because our goal is to investigate the therapeutic potential of vibration training for neuromuscular disease and low intensity is less likely to be deleterious to fragile, diseased muscle. Thus, an important point in regard to the present study is that no adverse effects of 6 wk of low-intensity vibration training were observed.
This research was supported by grants from the Muscular Dystrophy Association (DAL and DJN), the University of Minnesota Undergraduate Research Opportunity (JNM), and the National Institutes of Health; K02-AG036827 (DAL), P30-AR0507220 (University of Minnesota Muscular Dystrophy Center), and T32-AR07612 provided support to SAN, KAB, and JAC.
The authors thank Greg Cochrane and Angela Greising for technical assistance.
The authors disclose no conflict of interest.
Furthermore, publication of these results of the present study does not constitute an endorsement by the American College of Sports Medicine.
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