Effects of glutathione depletion and age on skeletal muscle performance and morphology following chronic stretch-shortening contraction exposure
- First Online:
- Cite this article as:
- Baker, B.A., Hollander, M.S., Kashon, M.L. et al. Eur J Appl Physiol (2010) 108: 619. doi:10.1007/s00421-009-1258-4
- 106 Views
The involvement of glutathione in the response of skeletal muscle following repetitive, high-intensity mechanical loading is not known. We examined the influence of a glutathione antagonist [l-Buthionine Sulfoximine (BSO)] had on the adaptability of skeletal muscle during chronic mechanical loading via stretch-shortening contractions (SSCs) in young and old rats. Left dorsiflexor muscles of young (12 weeks, N = 16) and old (30 months, N = 16), vehicle- and BSO-treated rats were exposed three times per week for 4.5-weeks to a protocol of 80 maximal SSCs per exposure in vivo. Skeletal muscle response to the SSC exposure was characterized by muscle performance, as well as muscle wet-weight and quantitative morphological analyses following the exposure period. Results reveal that generally, muscle performance increased in the young rats only following chronic SSC exposure. BSO treatment had no effect on muscle performance or morphology following the chronic SSC exposure in old rats. Muscle wet-weight was increased following exposure compared with the contra-lateral control limb, irrespective of age (p < 0.05). Muscle cross-sectional area increased approximately 20% with SSC loading in the young, vehicle rats, while increasing approximately 10% with SSC loading in old, vehicle rats compared with control rat muscle. No degenerative myofibers were noted in either age group, but edema were increased as a result of aging (p < 0.05). We conclude that our results indicate that glutathione depletion does not adversely affect muscle performance or morphology in old rats. Nevertheless, we continue to show that aging negatively influences performance and morphology following chronic SSC exposure.
KeywordsAdaptationAgingGlutathioneInflammationMuscle hypertrophyStretch-shortening contractions
Increased oxidative stress in skeletal muscle following various modes of physical activity promotes cellular damage and loss of function (Zerba et al. 1990). We have previously demonstrated that the chronic exposure of young and old rats to stretch-shortening contractions (SSCs) does not necessarily produce skeletal muscle degeneration/necrosis, but results in adaptation in young rats and a maladaptive response in old rats (Cutlip et al. 2006). Further, aging may influence the adaptive/maladaptive response (defined by changes in muscle performance and muscle morphology) in old rats via unresolved, low-grade inflammation (Cutlip et al. 2006). Since oxidative stress production following high-intensity mechanical loading occurs along with changes in cellular responses (McBride et al. 1998) in aged skeletal muscle, it is possible that increased oxidative stress reduces the ability of muscles to adapt to SSC loading as well, especially in aged populations. Moreover, increased oxidative stress may also exacerbate the maladaptive response, so to contribute to an injurious response manifested in skeletal muscle following mechanical loading/resistance-type training (Uchiyama et al. 2006). However, given that oxidative stress may also be an important stimulus for skeletal muscle adaptation (Urso and Clarkson 2003) in the absence of myofiber degeneration, modulating the oxidative stress profile and quantifying the resulting response may benefit our understanding with respect to optimizing mechanical-loading induced adaptations.
Glutathione, an important antioxidant and non-protein thiol source, is one mechanism maintaining cells in a reduced environment (Meister 1991), and serves various functions such as a reducing agent, a substrate for glutathione peroxidase, and recycling radicals generated from other antioxidant molecules. Because glutathione plays a major role in oxidant status, we modulated glutathione concentrations to determine if the adaptive capacity of skeletal muscle is under oxidant regulation. Glutathione can be depleted chronically using BSO, which is an irreversible inhibitor of γ-glutamylcysteine synthase (GCS) (Meister 1991; Leeuwenburgh and Ji 1995).
To assess skeletal muscle adaptation/maladaptation in young and old rats, we monitored changes in isometric and dynamic performance during a chronic 4.5-week exposure to SSCs. Following completion of the exposure, we also assessed morphological changes in skeletal muscle to determine if changes in performance were associated with changes in muscle morphology. Animals that have undergone glutathione depletion would be expected to be more vulnerable to mechanical loading-induced oxidative stress (Martensson and Meister 1989; Leeuwenburgh and Ji 1995). Our general hypothesis was that the glutathione acts to protect skeletal muscle from maladaptation (and subsequent muscle injury) and supports muscle adaptation and remodeling during chronic SSC loading. Thus, depleting glutathione via BSO will result in a failure of skeletal muscle to adapt to chronic SSC exposure in young rats. Further, because we previously demonstrated that old rats do not adapt to the SSC protocol administered in this study (Cutlip et al. 2006), we predicted that BSO treatment would exacerbate the effects of SSC exposure and increase indices of maladaptation or result in overt muscle injury in old rats.
We obtained male Fischer Brown Norway Hybrid rats (F344 × BN F1, n = 32) from the National Institutes on Aging colony, and housed young adult (n = 16; 3 months) and old (n = 16; 30 months) rats in an AAALAC accredited animal quarters. After 1 week of acclimatization, we randomly assigned young and old rats to BSO (n = 6), regular drinking water [vehicle, VEH (n = 6)], or cage-control groups [CON (n = 4)]. All animals, except those from the CON group, underwent exposure to a standardized experimental protocol approved by the NIOSH Animal Care and Use Committee. Temperature and light/dark cycle (dark cycle from 7.00 a.m. to 7.00 p.m.) remained constant for all animals; food and water were provided ad libitum for all groups, and water consumption was monitored for the experimental groups.
Rats randomized to the BSO group received water with 10 mM BSO, beginning 3 days prior to the first exposure (Watanabe et al. 2003). All other groups received unsupplemented water. We noted no differences in water consumption during and after the exposure period with respect to age or treatment group. Preliminary data demonstrated that 10 mM BSO depleted total glutathione levels in tibialis anterior (TA) muscle homogenate to approximately 65% of control levels in old rats.
We tested dorsiflexor muscles on a custom-built rodent dynamometer (Cutlip et al. 1997), which has been described in detail previously (Cutlip et al. 2004). Briefly, the dynamometer precisely controlled muscle length and muscle force output parameters. The software acquired and stored position, force, and velocity data in real-time as described elsewhere (Cutlip et al. 2004). We anesthetized the rats with 2% isoflurane gas using a small animal anesthetic system (Surgivet Anesco Inc., Waukesha, WI, USA) because it has no effect on in vivo force production (Ingalls et al. 1996). After anesthesia, we placed the rats supine on the heated x-y positioning table of the dynamometer with an anesthetic mask placed over the nose and mouth. We secured the knee in flexion (at 90°) with a knee holder, and the left foot in the load cell fixture using a custom-built foot holder with the ankle axis (assumed to be between the medial and lateral malleoli) aligned with the axis of rotation of the load cell fixture. We monitored each rat during the entire protocol to ensure proper anesthetic depth and body temperature.
With the experimental set-up, the functional testing has been described in detail previously (Cutlip et al. 2004). Briefly, we defined the joint position of the animal by the angle between the tibia and the plantar surface of the foot. The angular position of the load cell fixture corresponded with the angular position of the ankle. We measured the force produced by the dorsiflexor muscles at the interface of the aluminum sleeve and the dorsum of the foot. We placed platinum stimulating electrodes (Grass Medical Instruments, Quincy, MA, USA) subcutaneously to span the peroneal nerve for activation via the electrical stimulator, which resulted in muscle contraction of the dorsiflexor muscle group. To reduce the effect of excitation–contraction fatigue, all electrical stimulation times were kept to a minimum with 2 min of recovery time between stimulations (Ingalls et al. 1998).
We previously described the SSC protocol implemented in the current study in detail (Cutlip et al. 2006). Briefly, we exposed the dorsiflexor muscles of young and old experimental groups to 8 sets of 10 repetitions of SSCs with 2-min intervals between each set. Within each set, the muscles rested for 2 s between each stretch-shortening contraction. For each repetition, an electrical stimulator fully activated the dorsiflexor muscles for 100 ms, and the servomotor initiated the lengthening contraction phase with a 60 deg/s movement velocity of the load cell fixture over the prescribed range of motion of 90–140 deg ankle angle. The load cell fixture immediately returned in the concentric phase at 60 deg/s to the starting position of 90 deg ankle angle. The dorsiflexor muscles were deactivated 300 ms later with total stimulation time per repetition being 2.06 s. We administered the SSC protocol and performance tests three times per week for a total of 14 exposures over a 4.5 week period.
Isometric force test
We performed a pre-test isometric contraction (pre-test isometric force) and a post-test isometric contraction (post-test isometric force; executed immediately following the SSC protocol) on the dorsiflexor muscle group at an ankle angle of 90 deg using a 300 ms stimulation duration (Davis et al. 2003; Willems and Stauber 2001).
Dynamic force test
We measured a single SSC on the dorsiflexor muscle group 2 min preceding and following treatment with the SSC protocol as previously described (Cutlip et al. 2004). This test evaluated the muscle’s ability to generate dynamic forces and to perform work during dynamic stretch-shortening. Work was calculated in the same fashion as previous work by Cutlip et al. (2004); Geronilla et al. (2003); Ettema (1996); Stevens and Faulkner (2000); and Stevens (Stevens 1996). We carried out the single SSC by activating the dorsiflexor muscles for 300 ms then moving the load cell fixture from 70 to 140 deg at an angular velocity of 500 deg/s. The load cell fixture immediately returned to 70 deg, at 500 deg/s, and activation continued for 300 ms after cessation of the movement.
Histology and immunohistochemistry (IHC)
Twenty-four hours after the final exposure, we weighed, anesthetized (sodium pentobarbital ip, 10 mg/100 g body weight) and exsanguinated the rats. We dissected and weighed the left (exposed: LTA) and right (contra-lateral control: RTA) tibialis anterior muscles. We then divided the muscle into five equally-sized “zones” (“Zone 1” most proximal–“Zone 5” most distal), mounted them onto cork board with OTC (VWR, West Chester, Pennsylvania), froze the sections in liquid nitrogen-cooled isopentane, and stored all mounted sections at −80°C. Samples rested for 3 min at room temperature prior to freeze fixation in isopentane cooled in liquid nitrogen. We selected “Zone 3” to obtain the maximum tissue sample corresponding to the TA mid-belly, and cut transverse sections at 12-μm thickness, mounted on pre-coated microscope slides, air dried, and stained with hematoxylin & eosin (H&E) using Harris’ procedure. We assessed muscle morphology using standard stereological methods as used previously and described below. In addition, we used a glutathione primary antibody (Abcam Inc., Cambridge, MA, USA) and a fluorescently tagged secondary antibody (Cy3) and normal donkey serum for blocking (Jackson Immuno-Research Laboratories Inc., West Grove, PA) to localize and quantify total glutathione in TA muscle sections. Frozen sections used for glutathione immunolabeling that were stored at −80°C were fixed in ice cold acetone for 5 min and rinsed in PBS + 0.1 M Glycine for 5 min. We incubated slides in blocking buffer (5% normal donkey serum diluted in PBS + Triton X-100) at room temperature (RT) for 2 h. Next, we incubated slides in rabbit anti-glutathione antibody diluted in blocking buffer (1:100) for 1 h at 4°C, and following incubation, we rinsed slides in PBS and incubated for 1 h at RT in Cy3 labeled donkey anti-rabbit IgG (1:100). Finally, we rinsed slides, incubated in DAPI to stain nuclei (1:1,000, Sigma, St. Louis, MO, USA), coverslipped using Prolong Gold antifade reagent (Molecular Probes), and allowed to dry in a cool, dark area. We processed numerous slides in the absence of the primary antibody as controls for non-specific binding. Next, we captured photomicrographs using an Olympus Photomicroscope and Simple PCI Image Analysis Software. We quantified positively immunolabeled glutathione sections using one section per slide from the contra-lateral control RTA and exposed LTA muscles from each animal per group. Then we obtained 20 non-overlapping digital images at 40× magnification using the mid-point of the section as a reference and stored these as raw images. Finally, we transformed the digital color images into grey-scale images and obtained the mean area grey values (mArGV) using Optimas Image Analysis Software.
Fiber cross-sectional area
For muscle fiber cross-sectional area (CSA) analysis, we obtained ten non-overlapping digital images from H&E-stained muscle sections at a 40× magnification and analyzed fiber CSA (μm2) with Optimas Image Analysis Software. We traced approximately 200 fibers with a handheld mouse and calibrated the number of pixels inside the outlined region to a defined area in square micrometers (Cutlip et al. 2006).
We used quantitative morphometric methods to measure the volume fraction, surface densities, and average thickness of normal myofibers, degenerative myofibers, and the interstitial space (Baker et al. 2006). The interstitium was divided into endomysial and perimysial spaces, which included capillaries. We used a standardized stereological technique (Baker et al. 2006) to quantify the degree of myofiber degeneration and inflammation, which was characterized as either non-cellular interstitium (NCI), indicative of edema, or cellular interstitium (CI), indicative of cellular infiltrates. We measured fiber volume and surface density using standard morphometric analyses (Weibel 1972, 1974, 1975; Underwood 1970). Briefly, we took one of the H&E stained sections from each animal to identify the mid-point of the section on a stage micrometer. Next, we recorded point and intercept counts using a 121-point/11-line overlay graticule (12.5 mm square with 100 divisions) at 40× magnification at five equally spaced points across the section, and repeated this process 2 mm on both sides of the mid-point of the section for a total of 1,210 points or 110 intercept lines per section. We computed volume density or percent volume from the percentage of points over the tissue section to points over normal myofibers, degenerative myofibers, cellular interstitium and non-cellular interstitium plus capillaries (Weibel 1972, 1974, 1975). We counted intercepts over the line overlay for the perimeter of normal myofibers, degenerative myofibers, and interstitium to myofiber transitions. Points and intercepts over blood vessels greater than 25 μm in diameter were excluded. We evaluated one section per animal per group. Further, we used stereology to quantify the degree of myofiber degeneration, and the accompanying changes in the TA muscle from each group. We defined myofibers by the following criteria. Normal myofibers demonstrated: (1) complete contact with adjacent myofibers, (2) a smooth outer membrane, and (3) no presence of internal inflammatory cells. Degenerative myofibers displayed: (1) a loss of contact with adjacent myofibers, (2) presence of internal inflammatory cells, and (3) an outer membrane interdigitated with inflammatory cells.
Analysis of glutathione IHC densitometry, stereological measures and muscle wet weight, normalized to tibia length, were analyzed using a three-way mixed model (treatment × age × limb) analyses of variance with the animal as the random factor to account for measures in both limbs. Muscle quality and isometric and dynamic force test measures including isometric force, Fpeak (peak eccentric force), Fmin (isometric pre-stretch force), negative work, and positive work, and the dynamic force parameters Fpeak and Fmin were calculated as previously described (Cutlip et al. 2006). These measures from exposure 1 and 2 as well as data from exposure 13 and 14 were averaged and these dependent variables were analyzed using a two-way analyses of variance with repeated measures (treatment × age). All data were analyzed using JMP v. 5.1 (SAS Institute Inc., Cary, NC, USA), and post-hoc comparisons were analyzed using Fishers LSD.
Peak force (Fpeak)
Minimum force (Fmin)
Following 4.5 weeks of SSC exposures, muscle from young rats’ dorsiflexors produced greater Fmin than dorsiflexors from old rats (p < 0.05, Fig. 3c), regardless of treatment.
Unlike the observation for negative work, the ability to produce work differed with age prior to the first day of SSC exposure irrespective of treatment. Young rats’ dorsiflexors produced significantly greater work than old rats (p < 0.05, Fig. 4c). After the 4.5 weeks of SSC exposures, the young rats’ dorsiflexors produced 34.7% greater work than old rats.
Normalized muscle wet-weights
Stereological analyses of normal and degenerative myofibers
Stereological analyses of inflammation
Aging increased the volume density of NCI, indicative of edema, compared to young counterparts, regardless of treatment or limb (p < 0.001, Fig. 7c). Further, BSO-treated rats displayed increased NCI when compared to cage-control rats, as age and limb had no effect (p < 0.05, Fig. 7d). Aging did not influence the response of cellular interstitium, indicative of cellular infiltrates.
Fiber cross-sectional area
Cross-sectional area (CSA) data from young CON rats showed that approximately 46% of RTA and 48% of LTA muscle fibers were ≥2,000 µm2. CSA data from the old CON rats revealed similar results: approximately 50% of RTA and 46% of LTA muscle fibers were ≥2,000 um2. Thus, we established all myofibers ≥2,000 µm2 as being our qualitative hypertrophic threshold.
In the present investigation we, once again, report increased performance and hypertrophy in rats following SSC exposure and demonstrate that glutathione depletion does not negatively influence this outcome; however this response is blunted in old rodents. Surprisingly, an increase in cellular infiltrates was not present [this is in contrast to what we have previously reported in old rats (Cutlip et al. 2006)], although a trend did exist in the old rats for increased cellular infiltrates. It was hypothesized that depleting the host environment of glutathione would increase oxidative stress in young rats, and aging would intensify this response due to the damaging and cumulative effects of ROS that evolve over time (Bejma and Ji 1999). However, we have shown that old rats’ do not display a greater decrease in performance or dramatic change in morphology when exposed to chronic SSCs in the presence of glutathione depletion compared with exposure alone. We acknowledge that a limitation to the interpretation of the current data is our inability to show treatment efficacy for the young BSO-treated rodents using the immunohistochemical technique in the current study. In spite of this, we successfully showed that old BSO-treated rats did have decreased levels of total glutathione in skeletal muscle following treatment, yet this did not impact the functional or morphological response we observed for this group compared with the old VEH-treated group. Thus, one would expect that if glutathione depletion was detrimental to performance and morphology it would be observed with increased age, yet our old rats did not exhibit any unfavorable response following BSO treatment. Furthermore, to our knowledge, this is the first time that total glutathione has been localized and quantified in skeletal muscle using this immunohistochemical technique. Because performance and morphological changes may be achieved by specifically and selectively conserving glutathione in metabolically active tissue and allowing for maintenance of increased oxidative stress (Leeuwenburgh and Ji 1995) or by compensatory mechanisms (increases in other antioxidant systems) following glutathione depletion, this may be an explanation for not observing a decrease in glutathione levels in young rats. Thus, we suggest that glutathione depletion in old rats exposed to high-intensity mechanical loading via SSC exposure does not directly induce a maladaptive state and/or that glutathione is not critical for SSC-loading adaptation.
The differences observed with respect to isometric and dynamic performance measure prior to and following 1 and 14 SSC exposures in the current investigation compared with those published recently by our lab (Cutlip et al. 2006), may be the result of the variation we observed in the size of the TA muscle fibers. Previously, our morphometric data showed that young and old Fischer Hybrid rodent TA muscle fibers (of the same age in the current study) were smaller in size. This may account for the disparity we observed between both the initial force generating capacity of the rats in this study, as well as the adaptive/maladaptive response to exposure. The exposure paradigm used in the current study resulted in an increase of 4.2% for the average isometric force above the pre-test force in young, VEH-treated rats, while producing a 12.9% deficit in old, VEH-treated rats. The same general response was observed in the young and old rats receiving BSO treatment with respect to isometric performance. Whether a chronic increase in non-cellular interstitium promotes an environment that aggravates long-term cellular signaling leading to functional deficits is not known. Here, we show that estimates of edema were increased in old rats following SSC loading. Thus, unresolved permeability changes may have contributed to the decreased functional performance at this time. It is plausible that chronic alterations in cellular permeability with increased age may contribute to long-term maladaptation by modifying the local internal environment and ultimately affect muscle remodeling. Previous investigations support this hypothesis, because alterations in the host environment’s systemic factors with aging have been shown to adversely affect local tissue repair and regeneration (Conboy et al. 2005), which ultimately may affect function. Alternatively, age-related excitation–contraction (EC) coupling concurrent with calcium signaling and handling dysregulation has been indicated to affect specific force in single muscle fibers (Gonzalez et al. 2000). However, these events would manifest as early events (24–48 h) following the mechanical exposure, which we did not observe. Thus, although the role of calcium cannot be dismissed, a more appropriate means in which to consider EC coupling’s impact on performance with aging is that mechanical perturbation causes alterations and fragility of the transverse tubules (t-tubules) and associated receptor complexes (Payne and Delbono 2004) that may lead to decreased transmission efficiency. Furthermore, alterations in actin-myosin contractile proteins involved in cross-bridge cycling have been shown to contribute to an age-related decline in specific force (Lowe et al. 2002). Thus, these events may also influence the age-related decline in isometric and dynamic performance we observed in the current study. Furthermore, peak eccentric force increased in both the young, VEH-treated (12.1%) and young, BSO-treated (17.9%) rats but did not increase in the old, VEH- or BSO-treated rats. The data reported here for our VEH rats are in general agreement with Brooks et al. (2001) who reported a significant increase in peak force by week 6 of exposure for both adult and old animals over baseline values in mice. Disparities between our current performance data and that which has been reported previously may be related to differences between and within species and/or strains, as well as to differences we have noted with respect to muscle fiber size heterogeneity within this strain.
Changes in degenerative myofibers and cellular infiltrates were not significant following the terminal session in the current study, but changes in non-cellular interstitium, indicative of edema, in the muscles of old rats were significant. Further, degeneration and cellular interstitium were not influenced by glutathione depletion, even though estimates of edema were increased in BSO-treated rats. Collectively, the precise composition and magnitude of the general exposure response (degeneration, inflammation, and swelling) may be an important factor in signaling skeletal muscle adaptation (Tidball 2005). Thus, our data continues to indicate that the modifications made to the interstitial space are critical for adaptation, and the inability of old rats to fully adapt may be influenced by the internal host environment as hypothesized by others (Conboy et al. 2005). Furthermore, the modest morphological changes observed in old rats may have resulted from attenuation of the cellular interstitial response that we have recently reported (Cutlip et al. 2006). By decreasing the percent volume fraction of cellular interstitium, a more favorable host environment may have resulted in terms of permitting skeletal muscle adaptation. Nonetheless, we are again reporting the absence of degenerative myofibers following high-intensity mechanical loading, which supports a recent investigation from our lab (Cutlip et al. 2006), where the exposed muscles of rats did not exhibit myofiber degeneration. This may validate our hypothesis that not all high-intensity mechanical loading leads to overt skeletal muscle degeneration.
The data in the present study clearly show that SSC loading increases muscle mass in young rats, and that this response was diminished in old rats. The increase in muscle wet-weight could have resulted from and be attributed to chronic edema, but based on our findings, we do not believe this is the case. Results from our muscle cross-sectional area data support the observation that there was myofiber hypertrophy in the young, exposed rats as evidenced by a shift to larger fibers. Despite previous reports that indicate skeletal muscles in old animals are more susceptible to injury (Brooks and Faulkner 1996; Zerba et al. 1990), and recover more slowly (McBride et al. 1995; Brooks and Faulkner, 1990) from a single exposure to injurious contractions, there is evidence that old animals can be conditioned for protection from contraction-induced myofiber injury (Brooks et al. 2001). Muscle hypertrophy and improvements in force production occur in response to constant or chronic loading in aged animals, although the extent of muscle enlargement is attenuated relative to young animals (Alway 1995; Alway et al. 2002; Carson et al. 1995; Klitgaard et al. 1989a, b, Lowe et al. 1998). Since myofiber cross sectional area and stereology data from the contra-lateral limbs of both groups were not different, this suggests that there was not a different systemic response. The lack of myofiber degeneration in the exposed limbs of the old rats suggests that the decreased functional capacity observed in the TA muscle is not due to fiber degeneration, but necessitates future research into the causal factors influencing muscle structure–function relationships within aging populations.
In addition to age, several other factors may explain the difference in performance, physiological, and morphological measures between the two groups during the current chronic exposure and differences observed in a recent study from our lab (Cutlip et al. 2006). (1) The mode of exposure may significantly contribute to changes observed following repetitive exposure (anaerobic versus aerobic exposures). (2) The older animals may not have tolerated the repeated exposure to isoflurane as well as their younger counterparts, which could have affected contractile performance. (3) However, performance data clearly show that old rats have a limited capacity to adapt and that an alternate explanation to the differences observed may be influenced by inherent variability within or between cohorts of an animal strain or between species examined. As mentioned above, we have recently demonstrated that old rat’s do not adapt to the identical exposure paradigm functionally, physiologically, and morphologically. In the current study obvious maladaptation, which was observed previously (Cutlip et al. 2006), was not as striking.
Our findings suggest that glutathione depletion in old rats exposed to chronic SSCs does not induce an overt maladaptive state; however we continue to show that aging negatively influences performance and morphology following chronic SSC exposure when compared to young counterparts. Yet, we are unable to rule out completely the involvement of other oxidative pathways that may be influential in producing adaptation/maladaptation following chronic SSC exposures. Isolating elements that impact cellular signaling such as increased oxidative stress, inflammation, and/or edema in skeletal muscle under repetitive high-intensity mechanical loading that may influence the response of the oxidant/antioxidant profile continues to be of major importance when designing preventative strategies that attenuate muscular maladaptation in workplace and recreational settings.
The authors would like to thank Drs. Paul Nicolaysen, Kristine Krajnak, and Pius Joseph of the National Institute for Occupational Safety and Health (NIOSH) for their critical review and comments regarding the preparation manuscript.