Duchenne muscular dystrophy (DMD) is a devastating and fatal X-linked recessive disease that affects 1 in ~3500–5000 boys (1,2). DMD is characterized by skeletal, respiratory, and cardiac muscle deterioration because of the absence of dystrophin, a cytoplasmic protein associated with the plasmalemma (3). Dystrophin links the actin cytoskeleton to the transmembrane dystrophin–glycoprotein complex and is responsible for helping to maintain membrane integrity and cellular homeostasis (4–6). Skeletal muscle weakness, a clinical hallmark of DMD, develops in childhood and progressively worsens with age. Over time, patients with DMD lose ambulation and become wheelchair bound in their early teens, ultimately dying in their late 20s from respiratory and cardiac failure (3). Because of the nature of DMD, there is a great urgency to develop therapies that mitigate disease progression and increase the life span and overall health of these patients.
The mdx, dystrophin-deficient, mouse is the most widely used animal model to study DMD (7). A key feature of the mdx mouse is dramatic reductions in muscle function due to exercise that accentuates eccentric (ECC) contractions. Indeed, strength loss immediately after isolated ECC contractions is 20%–60% greater in mdx mice compared with wild-type mice, a finding that has been demonstrated in muscle in situ and ex vivo (8–11), as well as entire muscle groups in vivo (12–15). Susceptibility to ECC contraction-induced strength loss has therefore become a standard outcome measure in preclinical studies to assess disease severity and the efficacy of potential therapies for DMD. Despite the widespread use of ECC contraction-induced injury, the mechanisms underlying the initial loss and subsequent recovery of strength, and the effect ECC exercise has on muscle function in mdx mice, remain poorly understood. For the purpose of this manuscript, we define injury as the loss of muscle function (i.e., strength loss) resulting from any structural disruption or modification that occurs because of ECC contractions.
The excessive loss of strength in mdx mice after ECC contractions provokes the idea that the primary cause(s) of the injury may be different from that of wild-type mice (12,13). In wild-type mice, ECC contraction-induced strength loss is largely due to excitation–contraction uncoupling occurring between the voltage-sensitive dihydropyridine receptors embedded in the T-tubule membrane and the sarcoplasmic reticulum (SR) Ca2+ release channel (i.e., ryanodine receptor) (16–19). Excitation–contraction uncoupling in skeletal muscle can broadly be described as any disruption between electrical events occurring at the plasmalemma and SR Ca2+ release. However, given the location and function of dystrophin in skeletal muscle, it is plausible that the site of disruption in mdx mice could be at the plasmalemma and impair the generation and/or conduction of an action potential across the plasmalemma. Electrophysiological dysfunction of the plasmalemma has historically been overlooked, likely because the muscle of mdx mice typically has normal resting membrane potentials and excitability (13,20–22). What has not been realized until very recently is that plasmalemma electrophysiological properties are drastically impaired during and immediately after ECC contractions in mdx mice (13,14,23), data that have largely gone unnoticed.
Interestingly, despite greater strength deficits observed during and immediately after ECC contractions in mdx mice, in situ and in vivo, strength recovers faster compared with that of wild-type mice (8,12). A disconnect between strength loss and recovery further supports that sites of excitation–contraction uncoupling differ between wild-type and mdx mice. Given that the site causing strength loss is thought to be at the plasmalemma in mdx mice (13,23), it seems logical that the recovery of strength would therefore be dependent on restoring plasmalemma electrophysiological function. However, a detailed EMG analysis of the time course of recovery in mdx mice has not yet been examined, as previous studies only measured one recovery time point after the injury. Thus, the aim of experiment 1 was to substantiate that plasmalemma electrophysiological function is impaired when mdx mice perform repetitive ECC contractions and determine its subsequent rate of recovery. We hypothesized that the electrophysiological function of the plasmalemma would be compromised during and immediately after a bout of ECC contractions and fully recover in the days after the injury.
Skeletal muscle possesses an intrinsic ability to adapt after ECC contraction-induced injury, in such a way that it is protected against injury from future bouts of ECC contractions (i.e., repeated-bout effect) (24). When compared with the initial injury, adaptations in normal muscle manifest as increased strength, attenuated ECC contraction-induced strength loss, and an enhanced rate of strength recovery (12,24,25). Interestingly, when mdx mice perform several bouts of ECC contractions, the muscle eventually becomes stronger but never adapts to strength deficits during or immediately after injurious ECC contractions (12). It is plausible that the plasmalemma continues to be the main site of disruption in mdx mice regardless of previous ECC contraction bouts, and that recovery of plasmalemma electrophysiological function is paramount to the return of strength. Therefore, in experiment 2, we aimed to measure plasmalemma electrophysiological function after a second bout of ECC contractions and determine the subsequent rate of recovery. We hypothesized that irrespective of whether the injury bout was novel or repeated, the electrophysiological function of the plasmalemma would be impaired and recover in parallel with strength in mdx mice.
Ethical Approval and Animal Models
Male wild-type (C57Bl/10) and mdx (C57Bl/10ScSn-DMDmdx) mice were obtained from Jackson Laboratory (Bar Harbor, ME) or bred locally from these mice and were 3–6 months old at the time of the experimental protocols. Mice were initially anesthetized in an induction chamber using isoflurane and then maintained by inhalation of 1.5% isoflurane mixed with oxygen at a flow rate of 125 mL·min−1 for all surgical procedures. This anesthetic regimen was also used when torque and EMG measurements were made. After the final contraction protocol, mice were euthanized with an overdose of sodium pentobarbital (150 mg·kg−1 body mass). All animal procedures were in accordance with the standards set by the Institutional Animal Care and Use Committees at the University of Minnesota.
To determine recovery of the electrophysiological function of the plasmalemma in injured muscle, mice (wild-type, n = 8; mdx, n = 14) were chronically implanted with stimulating electrodes on the common peroneal nerve and EMG electrodes on the tibialis anterior (TA) muscle. Using these surgical techniques, we were able to use an in vivo approach to assess strength of the anterior crural muscles (TA, extensor digitorum longus, and extensor hallucis muscles) and plasmalemmal excitability of the TA muscle in a longitudinal study design. Specifically, anterior crural muscle isometric torque and TA muscle M-wave root mean square (RMS) were measured before, immediately after and at 1, 2, 6, 9, and 14 d after a single bout of 50 maximal ECC contractions (bout 1). Because the same mice were tested at every time point, preinjury values served as the control for all measurements.
To determine whether the plasmalemma continues to be the main site of disruption across repeated bouts of ECC contractions, a subset of mice from experiment 1 (wild-type, n = 6; mdx, n = 7) were subjected to a second injury. Specifically, after the 14-d assessment, mice were injured for a second time with 50 maximal ECC contractions (bout 2) and tested immediately after and at 1, 2, and 14 d postinjury. All assessments in experiment 2 were identical with those of experiment 1. When comparing changes across ECC contraction bouts or recovery time points, each measurement was expressed relative to the initial ECC contraction or preinjury isometric value for that given bout, respectively. Although this method of data interpretation may result in a less prominent repeated-bout effect due to strength gains (12), the overall goal of the second injury in this study was to determine whether M-wave RMS was reduced to the same degree between bouts 1 and 2 in mdx mice. Data pertaining to the repeated-bout effect in the muscle of wild-type mice when expressed as millinewton-meter per kilogram of torque have been published by Ingalls and colleagues (19,25).
An incision was made through the biceps femoris muscle in the left hindlimb, and a nerve cuff made of platinum iridium wire (Medwire-Sigmund Chon 10Ir9/49 T, Mt. Vernon, NY) and silastic tubing was placed around the common peroneal nerve (13,26,27). The proximal end of the nerve cuff was run subcutaneously to the dorsal cervical region and connected to a stimulator and stimulus isolation unit (Models S48 and SIU5, respectively; Grass Technologies, West Warwick, RI). No less than 21 d after implanting the stimulating nerve cuff, TA muscle EMG electrodes to record M-waves were then implanted in the anesthetized mouse (13,26,27). Briefly, deinsulated ends of two platinum iridium wires, offset by ~2 mm, were routed underneath the superficial fascial sheath of the TA muscle. The electrode wire spacing theoretically permitted sampling of EMG activity from the full thickness of the TA muscle beneath the electrodes (27,28). The wires were secured to adjacent tissue, and the proximal ends of the wires were run subcutaneously to the dorsal cervical region and connected to an EMG amplifier (Model P55, Grass Technologies). In vivo muscle testing with simultaneous M-wave measurements were initiated no less than 14 d after the EMG wire implantation.
Torque measurements and injury protocol
The in vivo maximal isometric torque of the anterior crural muscles was assessed as previously described (13,18,27,29). The anesthetized mouse (see above) was placed on a temperature-controlled platform to maintain rectal body temperature at 37°C, and the left knee was clamped and the left foot was secured to an aluminum footplate that is attached to the shaft of the servomotor system (Model 300B-LR; Aurora Scientific, Aurora, Ontario, Canada). The contractile function of the anterior crural muscles was assessed by measuring isometric torque as a function of stimulation frequency (20–300 Hz; 150-ms train with 0.1-ms pulses). The anterior crural muscles were then injured by performing 50 electrically stimulated maximal ECC contractions. During each ECC contraction, the foot was passively moved from 0° (positioned perpendicular to tibia) to 19° of dorsiflexion where the anterior crural muscles performed a prelengthening 100-ms isometric contraction followed by an additional 20 ms of stimulation while the foot was moved from 19° of dorsiflexion to 19° of plantarflexion at 2000°·s−1. A 5-min rest after the ECC contraction protocol was given before reassessing contractile function via the aforementioned torque-frequency protocol.
Analysis of the electrically evoked myoelectric signal was done using the amplitude measure (i.e., M-wave RMS) as previously described (13,26,27). M-wave RMS was calculated for the full 150 ms of the isometric contractions during the torque-frequency protocols and across the first 100 ms (i.e., the isometric portion) of the ECC contraction protocol. Because the anterior crural muscles were maximally recruited via electrical stimulation of the common peroneal nerve, a decrease in M-wave RMS was interpreted as impairment of action potential generation and/or conduction (13,26,27). Figure 1A and B depicts representative isometric tetanic torque time and M-wave time tracings for wild-type and mdx mice.
A one-way repeated-measures ANOVA was used to assess differences within strains across time, whereas a two-way ANOVA was used to probe for differences between strains across time or injury bout for torque and M-wave RMS. When significant interactions or main effects were calculated, differences were tested with Holm–Sidak post hoc tests. An α level of 0.05 was used for all analyses. Values are presented as mean ± SE. All statistical testing was performed using SigmaPlot version 12.5 (Systat Software, San Jose, CA).
In both mouse strains, maximal ECC torque and M-wave RMS decreased over the injury protocol (P < 0.001; Fig. 2A and B); however, deficits were far greater in mdx mice (P < 0.001). At the completion of the injury protocol, maximal ECC torque and M-wave RMS decreased 68% ± 1% (P < 0.001) and 78% ± 3% (P < 0.001), respectively, in mdx mice, but only 28% ± 2% (P < 0.001) and 12% ± 3% (P = 0.004), respectively, in wild-type mice. Moreover, although maximal ECC torque and M-wave RMS both decreased over the contraction protocol in wild-type mice, the reduction observed in M-wave RMS was greater than the deficit recorded in maximal ECC torque (P = 0.003) (Fig. 2A).
As with maximal ECC torque and M-wave RMS during the injury protocol, reductions in maximal isometric torque and M-wave RMS assessed immediately postinjury were greater in mdx mice (P < 0.001). When compared with preinjury values, maximal isometric torque and M-wave RMS decreased 62% ± 3% (P < 0.001) immediately postinjury in mdx mice (Figs. 1B and 2D), whereas in wild-type mice, maximal isometric torque only decreased 35% ± 3% (P < 0.001) and M-wave RMS did not change (P = 0.390; Figs. 1A and 2C). By day 2, maximal isometric torque was not different from preinjury torque in wild-type mice (P = 0.311) (Fig. 2C). Maximal isometric torque and M-wave RMS took longer to return to preinjury values in mdx mice compared with wild-type mice (P < 0.001) (Fig. 2C and 2D"). In mdx mice, M-wave RMS and maximal isometric torque were not different from preinjury values by day 6 (P = 0.106) and day 9 (P = 0.333), respectively (Fig. 2D).
To characterize plasmalemma electrophysiological function during and after a repeated bout of ECC contractions, a subset of mice was subjected to a second bout of 50 maximal ECC contractions 14 d after the initial injury (Fig. 3A–D). In wild-type mice, maximal ECC torque at the 50th contraction was greater after bout 2 (105.94 ± 3.78 vs 123.55 ± 4.63 mN·m·kg−1, P = 0.008) and ECC torque deficits tended to be less (29% ± 1% vs 24% ± 1%, P = 0.054; Fig. 3A), whereas reductions in M-wave RMS did not differ between bouts (11% ± 4% vs 8% ± 2%, P = 0.457; Fig. 3C). Maximal ECC torque at the 50th contraction (52.55 ± 3.42 vs 55.19 ± 2.64 mN·m·kg−1, P = 0.468) and deficits in ECC torque (69% ± 2% vs 67% ± 2%, P = 0.193; Fig. 3B) and M-wave RMS (85% ± 2% vs 81% ± 3%, P = 0.423; Fig. 3D) were similar between bouts 1 and 2 in mdx mice.
Maximal isometric torque and M-wave RMS in wild-type mice did not differ between bouts immediately after or at 1, 2, or 14 d after the injuries (P ≥ 0.132; Fig. 4A and C). Maximal isometric torque in mdx mice was also similar between bouts immediately after, 1 d, and 14 d after the injuries (P ≥ 0.108; Fig. 4B). Two days postinjury after the second bout of ECC contractions, mdx mice produced 32% more maximal isometric torque compared with the first bout (P = 0.011; Fig. 4B). No differences in M-wave RMS were measured between the bouts at any time point during the maximal isometric contractions in mdx mice (P ≥ 0.266; Fig. 4D).
We had four primary results from this study, all of which support our hypotheses. First, plasmalemma electrophysiological dysfunction is a major contributor to ECC contraction-induced strength loss in mdx mice. Second, loss of plasmalemma electrophysiological function is a transient event after ECC contraction-induced skeletal muscle injury in mdx mice. Third, excitation–contraction uncoupling at the plasmalemma is present across repeated bouts of ECC contractions. Lastly, a second ECC contraction bout does not exacerbate plasmalemma electrophysiological dysfunction beyond that of the initial injury or influence its subsequent rate of recovery. These results substantiate that a major plasmalemma-based mechanism is one of the underlying causes for the loss and return of strength after ECC contractions in mdx mice, which is distinctly different from that of wild-type mice.
Loss of plasmalemma electrophysiological function because of a bout of ECC contractions was dramatic in mdx mice. M-wave RMS decreased 78% in mdx mice, but only 12% in wild-type mice corresponding to maximal ECC torque deficits of 68% and 28%, respectively (Fig. 2A and B). In fact, it took less than four ECC contractions to achieve a ~12% reduction in M-wave RMS in mdx mice (vs 50 ECC contractions in wild-type mice). As observed across the ECC contractions, maximal isometric strength and M-wave RMS in mdx mice were also drastically reduced immediately postinjury, whereas maximal isometric torque deficits in wild-type mice did not parallel the loss in M-wave RMS (Figs. 1 and 2C, D). These data corroborate two important physiological components of ECC contraction-induced strength loss previously reported by various research groups (14,23), including our own (13,27). First, functional dystrophin is required to maintain plasmalemmal excitability during maximal ECC contractions, and second, ECC contraction-induced strength loss is mechanistically different between wild-type and mdx mice.
Transmission failure at the neuromuscular junction (NMJ) or an impaired ability to conduct an action potential along the plasmalemma, two proximal events during muscle excitation, would cause decrements in M-wave RMS in injured mdx mice. Evidence that NMJ transmission failure contributes to ECC contraction-induced strength loss is supported by histological data of altered NMJ morphology in the injured muscle of mdx mice but not wild-type mice (14,15). However, not all studies support neuromuscular transmission failure as a primary mechanism contributing to loss of M-wave RMS in mdx mice (13,23). Roy et al. (23) reported that despite NMJ morphology being influenced by ECC contractions, bypassing the NMJ via direct stimulation of the injured muscle did not markedly improve strength in mdx mice. It should also be noted that the NMJ of mdx muscle is discontinuous and dispersed before ECC contraction-induced injury (14,15,23), yet in vivo maximal isometric torque is not different when compared to muscle with normal NMJ morphology (i.e., wild-type muscle) (14,15,30,31). Thus, alterations in NMJ morphology may not always translate into NMJ dysfunction. Although we cannot fully rule out NMJ transmission failure in our study, these results suggest that plasmalemmal inexcitability in the injured muscle of mdx mice does not originate at the NMJ but rather occurs at a downstream site on the plasmalemma.
Using the same in vivo injury model as described in the present study, Call and coworkers (13) demonstrated that a significant number of fibers were depolarized in mdx muscle. Immediately after a bout of ECC contractions, 50% of the mdx muscle fibers were more positive than −55 mV, compared with only 7% from that of wild-type muscle fibers (13). When skeletal muscle fibers are depolarized above −55 mV, they are rendered inexcitable and contribute minimally to force development (32–34). These data indicate that loss of M-wave RMS after the injury was due to the sustained depolarization of a large number of mdx muscle fibers, which contributed to fewer action potentials being generated and/or conducted along the plasmalemma (13,27). Theoretically, changes in transplasmalemmal K+ gradients, along with Na+, Cl−, and/or Ca2+ could explain muscle fiber depolarization and thus reductions in M-wave RMS (23,35). In support of this theory, a Cl− channel inhibitor improved force 10 min after a bout of ECC contractions in mdx mice, and authors attributed the improvement in force to an overall increase in muscle excitability (23). It is likely that multiple plasmalemma-associated proteins and channels are affected by ECC contractions (36,37) and contribute to plasmalemma electrophysiological dysfunction in mdx mice.
It is also possible that the ECC contractions additionally disrupted other sites distal to the plasmalemma, as mdx fibers that lost M-wave RMS were depolarized and inexcitable postinjury. For instance, ECC contractions have also been shown to impair force development at the myofilaments in mdx fibers. Blaauw et al. (38) demonstrated Ca2+-activated force by permeabilized mdx fibers, taken from muscle previously injured in vivo, generated 28% less force than uninjured fibers. By using this experimental approach, the researchers were able to bypass voltage-gated SR Ca2+ release and make conclusions about force development at the myofilaments (38). These data suggest that ECC contractions may impair force development at the myofilaments in mdx mice, in addition to disrupting plasmalemma electrophysiological function.
In this experiment, we also assessed the recovery of maximal isometric torque and M-wave RMS at five time points between 1 and 14 d after the ECC contractions. Plasmalemma electrophysiological function was only down 14% in mdx mice 2 d postinjury, despite a 62% deficit being recorded immediately after the ECC contractions (Fig. 2D). Maximal isometric torque also showed a remarkable rebound, although not quite as rapid as M-wave RMS. To put this rate of recovery into perspective, maximal isometric torque in wild-type muscle injured by 150 ECC contractions is still reduced by 25%–30% after 7 d, although the initial injury only causes a 45%–55% deficit (39,40). In the present study, maximal isometric torque was reduced 62% immediately after the ECC contractions, but it only remained 18% lower 6 d into recovery (Fig. 2D). These data emphasize that because the underlying mechanisms of the injury are different in mdx and wild-type mice, the recovery processes will also be different. We suggest that the full recovery of strength in mdx mice is primarily dependent on restoring plasmalemma electrophysiological function and any other damaged or modified sites downstream of the plasmalemma.
To further examine plasmalemma electrophysiological function in mdx mice, a subset of mice were injured for a second time. We show that reductions in maximal ECC torque and M-wave RMS did not differ between contraction bouts in mdx mice (Fig. 3B and D), whereas ECC torque deficits tended to be less in wild-type mice after the second injury (Fig. 3A). These results confirm previous reports that demonstrate mdx mice show no protection from ECC torque drop (12), which we contend is due to reductions in plasmalemma electrophysiological function that are present regardless of injury bout. As hypothesized, the recovery of plasmalemma electrophysiological function was also similar across injury bouts in mdx mice (Fig. 4D). These data confirm that repeated bouts of ECC contractions do not exacerbate muscle injury or recovery in mdx mice, which is partially facilitated by the loss and return of plasmalemma electrophysiological function.
Excitation–contraction uncoupling at the plasmalemma through repeated bouts of ECC contractions suggests that loss of excitability may be an adaptive mechanism in the muscle of mdx mice. Recently, ECC contraction-induced strength loss in mdx muscle was theorized to function as a circuit breaker that potentially protects the muscle from cell-lethal structural damage (10). Our data support these concepts for several reasons. First, plasmalemma electrophysiological function was drastically impaired by a limited number of ECC contractions, regardless of the contraction bout (Figs. 2B and 3D). Second, loss of plasmalemmal excitability was a transient event, recovering soon after the injury (Fig. 2D). Lastly, the muscle of mdx mice did not adapt to ECC contraction-induced loss of plasmalemma electrophysiological function (Fig. 3D) but was still able to recover from repeated injuries (Fig. 4B and D). Additional studies are needed to determine whether sites downstream of the plasmalemma are also disrupted by ECC contraction-induced injury, and thus, if loss of plasmalemmal excitability is simply the most proximal circuit breaker in mdx muscle. Elucidating the roles of other proteins associated with the dystrophin–glycoprotein complex (e.g., dystroglycans and utrophin) that might contribute to the ECC contraction-induced electrophysiological dysfunction of the plasmalemma is also important future work.
In conclusion, exercise that accentuates ECC contractions induces significant strength loss in mdx mice because of plasmalemma electrophysiological dysfunction, likely mediated by a failure to conduct action potentials in depolarized muscle fibers. Despite the rather dramatic perturbation observed in plasmalemma electrophysiological function during and immediately after the injury, mdx mice possess a remarkable ability to recover plasmalemmal excitability. Furthermore, a second injury does not appear to exacerbate the mdx phenotype, as plasmalemma electrophysiological function is lost and restored at a similar rate as the first injury. These results highlight that ECC contraction-induced strength loss is mechanistically different between wild-type and mdx mice. The data also indicate that functional dystrophin is necessary for excitation to occur at the plasmalemma during a series of ECC contractions, but not essential for the complete recovery of plasmalemma electrophysiological function or maximal isometric strength.
This work was funded by the National Institutes of Health (grant no. T32-AG029796 and T32-AR007612 to C. W. B.), a grant from the University of Minnesota Bob Allison Ataxia Research Center (to D. A. L.), and a research endowment from the American College of Sports Medicine Foundation (to C. W. B).
The authors have no conflict of interest. The authors declare that the results of the study are presented clearly, honestly, and without fabrication, falsification, or inappropriate data manipulation and that the results of the present study do not constitute endorsement by the American College of Sports Medicine.
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