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The Journal of Physiology logoLink to The Journal of Physiology
. 2018 Jul 14;596(14):2883–2900. doi: 10.1113/JP275833

Alterations in the muscle force transfer apparatus in aged rats during unloading and reloading: impact of microRNA‐31

David C Hughes 1, George R Marcotte 1, Leslie M Baehr 2,3, Daniel W D West 2,3, Andrea G Marshall 1,2, Scott M Ebert 4,5,6, Arik Davidyan 7, Christopher M Adams 4,5,6, Sue C Bodine 1,2,3, Keith Baar 1,2,3,✉
PMCID: PMC6046073  PMID: 29726007

Abstract

Key points

  • Force transfer is integral for maintaining skeletal muscle structure and function. One important component is dystrophin. There is limited understanding of how force transfer is impacted by age and loading.

  • Here, we investigate the force transfer apparatus in muscles of adult and old rats exposed to periods of disuse and reloading.

  • Our results demonstrate an increase in dystrophin protein during the reloading phase in the adult tibialis anterior muscle that is delayed in the old muscle. The consequence of this delay is an increased susceptibility towards contraction‐induced muscle injury.

  • Central to the lack of dystrophin protein is an increase in miR‐31, a microRNA that inhibits dystrophin translation. In vivo electroporation with a miR‐31 sponge led to increased dystrophin protein and decreased contraction‐induced muscle injury in old skeletal muscle.

  • Overall, our results detail the importance of the force transfer apparatus and provide new mechanisms for contraction‐induced injury in ageing skeletal muscle.

Abstract

In healthy muscle, the dystrophin‐associated glycoprotein complex (DGC), the integrin/focal adhesion complex, intermediate filaments and Z‐line proteins transmit force from the contractile proteins to the extracellular matrix. How loading and age affect these proteins is poorly understood. The experiments reported here sought to determine the effect of ageing on the force transfer apparatus following muscle unloading and reloading. Adult (9 months) and old (28 months) rats were subjected to 14 days of hindlimb unloading and 1, 3, 7 and 14 days of reloading. The DGC complex, intermediate filament and Z‐line protein and mRNA levels, as well as dystrophin‐targeting miRNAs (miR‐31, ‐146b and ‐374) were examined in the tibialis anterior (TA) and medial gastrocnemius muscles at both ages. There was a significant increase in dystrophin protein levels (2.79‐fold) upon 3 days of reloading in the adult TA muscle that did not occur in the old rats (P ≤ 0.05), and the rise in dystrophin protein occurred independent of dystrophin mRNA. The disconnect between dystrophin protein and mRNA levels can partially be explained by age‐dependent differences in miR‐31. The impaired dystrophin response in aged muscle was followed by an increase in other force transfer proteins (β‐dystroglycan, desmuslin and LIM) that was not sufficient to prevent membrane disruption and muscle injury early in the reloading period. Inserting a miR‐31 sponge increased dystrophin protein and decreased contraction‐induced injury in the TA (P ≤ 0.05). Collectively, these data suggest that increased miR‐31 with age contributes to an impaired dystrophin response and increased muscle injury after disuse.

Key points

  • Force transfer is integral for maintaining skeletal muscle structure and function. One important component is dystrophin. There is limited understanding of how force transfer is impacted by age and loading.

  • Here, we investigate the force transfer apparatus in muscles of adult and old rats exposed to periods of disuse and reloading.

  • Our results demonstrate an increase in dystrophin protein during the reloading phase in the adult tibialis anterior muscle that is delayed in the old muscle. The consequence of this delay is an increased susceptibility towards contraction‐induced muscle injury.

  • Central to the lack of dystrophin protein is an increase in miR‐31, a microRNA that inhibits dystrophin translation. In vivo electroporation with a miR‐31 sponge led to increased dystrophin protein and decreased contraction‐induced muscle injury in old skeletal muscle.

  • Overall, our results detail the importance of the force transfer apparatus and provide new mechanisms for contraction‐induced injury in ageing skeletal muscle.

Introduction

Ageing is associated with a progressive decline in muscle mass (sarcopenia) and strength (dynapenia) (Matthews et al. 2011; Clark & Manini, 2012). The loss in muscle strength with age is primarily attributed to decrements in muscle protein mass (cross‐sectional area) and/or decreased neural activation. However, the decrease in muscle quality (lower force per unit area) could also result from oxidation of the contractile proteins (Javadov et al. 2015) or impaired force transfer (Street, 1983; Ramaswamy et al. 2011; Hughes et al. 2015). There is evidence that contractile protein oxidation increases with age; however, whether this contributes to the loss of muscle quality remains equivocal (Thompson et al. 2006). The role of force transfer in this process has, in contrast to the other factors, received less attention. Skeletal muscle transmits force in both the longitudinal and lateral directions (Street, 1983). Longitudinal force transfer is the passage of stress along the length of the muscle fibre, from Z‐line to Z‐line. In contrast, lateral force transmission is necessary to transmit force from the centre of the fibre to the overlying connective tissue and extracellular matrix, linking parallel muscle fibres tightly together (Huijing, 1999; Zhang & Gao, 2014). While both lateral and longitudinal force development are critical, more than 80% of force produced within a muscle is believed to be transferred laterally (Street, 1983; Ramaswamy et al. 2011). Given that a loss of strength has been identified as a predictor of all‐cause mortality (Metter et al. 2002), understanding force transmission and the cytoskeletal proteins involved may provide new avenues for enhancing muscle quality in elderly individuals.

Two costameric protein families, the dystrophin‐associated glycoprotein complex (DGC) and the integrins, are responsible for lateral force transmission, providing a strong mechanical link between the extracellular matrix (ECM) and adjoining fibres (Ramaswamy et al. 2011; Gao & McNally, 2015). Components of the DGC complex, dystrophin, α‐sarcoglycan and sarcospan, are essential for mechanoprotection from shear stress and reduce contraction‐induced injury (Lovering & De Deyne, 2004; Ramaswamy et al. 2011; Gao & McNally, 2015). The α7β1 integrin also plays a role in protecting muscle from contraction‐induced injury and transducing the biochemical response to loading (Boppart et al. 2006, 2008). These two costameric complexes work together to provide a tight connection between the sarcomere and the ECM (Hughes et al. 2015). In fact, in animals that lack dystrophin the α7 integrin is upregulated, and when both proteins are knocked out, the result is a more severe muscular dystrophy (Rooney et al. 2006). Recent evidence has identified a loss of dystrophin protein in aged muscles that occurs independent of gene expression initially in the flexor muscles and later in the extensors (Ramaswamy et al. 2011; Hughes et al. 2017). In response to the initial drop in dystrophin, it appears that the other force transmission proteins increase in an attempt to compensate (Hughes et al. 2017); however, the compensatory increases cannot overcome the loss of dystrophin, resulting in reduced lateral force transmission and an increase in contraction‐induced injury to the aged muscle (Ramaswamy et al. 2011; Hughes et al. 2017).

Loss of dystrophin protein independent of expression in disease, age or injury has led to the identification of several microRNAs (miRs) that target dystrophin, termed dystromirs (Greco et al. 2009; Cacchiarelli et al. 2011; Fiorillo et al. 2015; Hughes et al. 2017). Several labs have independently identified at least three miRs that can target dystrophin (miR‐31, miR‐146b and miR‐374). Previous work has elegantly shown that these miRs bind to the 3' untranslated region (UTR) of the dystrophin mRNA and decrease translation efficiency (Cacchiarelli et al. 2011; Fiorillo et al. 2015). Of the three dystromirs, miR‐31 has been observed to be increased >70‐ and >20‐fold in skeletal muscle of mdx mice and muscular dystrophy (Duchenne and Becker) patients, respectively (Greco et al. 2009; Fiorillo et al. 2015). Furthermore, miR‐31 expression appears to be elevated after an episode of muscle damage/injury (Greco et al. 2009). In terms of aged skeletal muscle, we have previously demonstrated that miR‐31 is elevated in old rat muscle (28 months) and may contribute to the lower dystrophin protein observed (Hughes et al. 2017). Interestingly, dystrophin protein is maintained for longer in muscles that are loaded more frequently (e.g. gastrocnemius), suggesting that loading could alter dystrophin protein independent of dystromir level (Hughes et al. 2017).

Aged muscle is more susceptible to injury from contraction (Barash et al. 2002; Lovering & De Deyne, 2004). This is very important for musculoskeletal function since: (1) there is a reduced capacity for recovery following contraction‐induced muscle injury with age, and (2) contraction‐induced muscle injury may contribute to sarcopenia (Holloszy et al. 1995). Old muscle also shows a reduced capacity to recover from unloading and reloading, a more physiological model that is very similar to bedrest and recovery in humans (Suetta et al. 2009; Hvid et al. 2010). These data indicate that old muscle might suffer more injury following unloading and subsequent reloading. Since the force transfer apparatus and the cytoskeleton play an important role in preventing contraction‐induced injury/membrane damage, and the old tibialis anterior (TA) muscle displays a decrease in dystrophin, we hypothesized that force transfer within old TA muscle may be more affected by unloading and reloading resulting in greater muscle injury.

Even though force transfer is a key component of force development, rate of force development and injury prevention, there is limited research into how these cytoskeletal proteins are affected by periods of disuse and mechanical loading; further, the role of ageing on these proteins remains largely unexplored (Hughes et al. 2015). Therefore, the aims of this study were to: (1) investigate the impact of unloading and reloading on the force transfer apparatus in adult and aged skeletal muscle; (2) determine the role of unloading and reloading on the expression of the dystromir microRNAs; and (3) determine the impact of alterations in dystromirs and cytoskeleton proteins on the susceptibility of the muscle to contraction‐induced muscle injury. We hypothesized that: (1) cytoskeletal proteins involved in force transmission are decreased in aged muscle during reloading; (2) lower levels of force transfer proteins during reloading makes old muscle more prone to injury; and (3) increasing dystrophin protein protects old muscle from contraction‐induced muscle injury.

Methods

Animals/ethical approval

Adult (9 months) and old (28 months) male Fischer 344‐Brown Norway rats were obtained from the National Institute of Aging. The Fischer Brown Norway (FBN) rat is a well‐established rodent model for the study of ageing due to their increased lifespan and lower incidence of disease pathologies (Turturro et al. 1999). These ages were selected since muscle mass plateaus at approximately 9 months of age and signs of sarcopenia appear at age 30 months in the FBN rats (Hwee & Bodine, 2009). Therefore, the animals in this study were at their muscular peak (9 months) and pre‐sarcopenic (28 months). The rats utilized in the unloading and reloading study are from a larger study that specifically examined the recovery of hindlimb muscles from disuse (Baehr et al. 2016). Once in the facility, rats were allowed to acclimatize in their cages for at least 1 week prior to testing. All animal procedures were approved by the Institutional Animal Care and Use Committee at the University of California, Davis.

Hindlimb unloading/reloading

Unloading of the hindlimbs was achieved by tail suspension as previously described (Baehr et al. 2016). The rats were attached via a plastic bar to a swivel mounted at the top of the cage, allowing free 360° rotation. The rats were maintained in an ∼30° head‐down tilt position with their hindlimbs unloaded for a period of 14 days (n = 6–7/group). Rats had access to food and water ad libitum. Animals were released from the tail suspension on day 15 at which time the rats were individually housed and allowed unrestricted cage activity for periods ranging from 1 to 14 days (n = 6–7/group).

In vivo electroporation

The electroporation of DNA plasmids was performed as previously described (Ebert et al. 2010; MacKenzie et al. 2013). Briefly, after a 2 h pre‐treatment with 0.4 units/μl of hyaluronidase, 20 μg of plasmid DNA was injected into the TA muscle, the hindlimbs were placed between two paddle electrodes and subjected to 8 pulses (20 ms) of 160 V/cm using an ECM‐830 electroporator (BTX Harvard Apparatus, Cambridge, MA, USA). Two groups of 28 month‐old FBN rats (n = 8/group) were injected with either a miR‐31 sponge (CMV‐miR‐31 sponge was a gift from Bob Weinberg (Addgene plasmid no. 25025)) or cs2‐GFP plasmid as a control (GFP), donated by David Furlow, UC Davis, Davis, CA, USA. The use of a pre‐treatment with hyaluronidase allows for enhanced levels of transfected muscle without increased levels of muscle damage during the electroporation process (McMahon et al. 2001). Seven days post‐transfection, animals undertook a lengthening contraction protocol (described below).

Lengthening contraction protocol

To determine the propensity for muscle injury, 28 month‐old FBN rats electroporated with either the GFP or sponge plasmids underwent a bout of acute unilateral resistance exercise using a protocol described previously (Baar & Esser, 1999; Hamilton et al. 2010). Briefly, rats were chemically restrained (2.5% isoflurane) and the right sciatic nerve was stimulated (100 Hz, 3–6 V, 1 ms pulse, 9 ms delay) for 10 sets of six repetitions (repetition length = 2 s). In this model, muscle fibres in the extensor digitorum longus (EDL) and TA perform high‐force lengthening contractions. Following exercise, all animals (GFP and Sponge groups) were given an analgesic (buprenorphine, 0.1 mg/kg) and returned to their cages for 24 h. TA muscles were harvested after 24 h and immediately frozen in liquid nitrogen for histochemical and biochemical analysis.

Muscle collection

Following completion of the unloading or reloading time period, rats were anaesthetized with 2.5% inhaled isoflurane, and the following muscles were excised from both hindlimbs: medial gastrocnemius (MG), plantaris (PLN) and TA. All muscles were weighed and frozen in liquid nitrogen for biochemical analyses, except the right PLN and TA, which were pinned on cork at a length approximating resting length (L o) and frozen in liquid nitrogen‐cooled isopentane for histological analyses. We chose to freeze the MG for biochemistry to account for the large number of assays we were undertaking with this tissue. We froze the PLN for histochemistry to serve as a representative plantar flexor muscle whose recruitment and recovery from unloading is similar to the MG.

Immunoblotting

Frozen TA and MG muscles were homogenized in sucrose lysis buffer (50 mM Tris pH 7.5, 250 mM sucrose, 1 mM EDTA, 1 mM EGTA, 1% Triton X‐100, and protease inhibitors). The supernatant was collected following centrifugation at 10,000 g for 5 min and protein concentrations were determined in triplicate using the DC protein assay (Bio‐Rad, Hercules, CA, USA). Twenty micrograms of protein was subjected to SDS‐PAGE on 4–20% Criterion TGX gels (Bio‐Rad) and transferred to nitrocellulose membrane for 2 h. Membranes were blocked in 1% fish skin gelatin dissolved in Tris‐buffered saline with 0.1% Tween‐20 (TBST) for 1 h and then probed with primary antibody overnight at 4⁰C. The next day, membranes were washed and incubated with HRP‐conjugated secondary antibodies at 1:10,000 for 1 h at room temperature. Immobilon Western Chemiluminescent HRP substrate (Millipore, Hayward, CA, USA) was then applied to the membranes for protein visualization by chemiluminescence. Image acquisition and band quantification was performed using the ChemiDoc MP System and Image Lab 5.0 software (Bio‐Rad). Total protein staining (whole lane) of the membrane (via Ponceau) was used as the normalization control for all blots. The following antibodies were used in this study at a concentration of 1:1000: dystrophin (Santa Cruz, Cat. no. 365954), β‐dystroglycan (Hybridoma Bank, Cat. no. MANDAG2), α‐sarcoglycan (Hybridoma Bank, Cat. no. IVD3 A9), Dicer (Santa Cruz, Cat. no. 136979), GFP (Cell Signaling, Cat. no. 2956), phosphor‐UBF Ser637 (Santa Cruz, Cat. no. 21639), laminin‐2α (Santa Cruz, Cat. no. 20142), β1‐integrin (Santa Cruz, Cat. no. 6622), desmin (Hybridoma Bank, Cat. no. D76), desmuslin (Santa Cruz, Cat. no. 49651), α‐actinin (Cell Signaling, Cat. no. 6487) and muscle LIM protein (Santa Cruz, Cat. no. 166930). Sarcospan cross‐reacted with the antibody for dystrophin (Santa Cruz, Cat. no. 365954) and was determined by molecular weight (Ohlendieck & Campbell, 1991).

RNA isolation and real‐time quantitative polymerase chain reaction (RT‐qPCR)

Prior to RNA isolation, aliquots of frozen muscle powder from the TA and MG were weighed in order to calculate RNA per milligram of wet muscle tissue. Total RNA was extracted with Tri Reagent Solution (Ambion Inc., Austin, TX, USA; AM9738) according to the manufacturer's instructions. Following isolation, RNA concentrations were determined spectrophotometrically (Epoch Microplate Spectrophotometer, BioTek Instruments Inc., Winooski, VT, USA). cDNA was synthesized using a reverse transcription kit from 1 μg of total RNA. Gene expression was analysed by quantitative PCR (qPCR) using SYBR Green JumpStartTaq ReadyMix on an ABI 7900HT thermocycler. Each sample was run in triplicate. Primer sequences are available on request. Gene expression was normalized to tissue weight as previously described (Heinemeier et al. 2009) since all standard housekeeping genes showed significant change either with ageing or unloading/reloading.

Determination of microRNAs (dystromirs)

Following RNA isolation and concentration quantification, 1 μg of RNA was DNase treated (Invitrogen, Carlsbad, CA, USA; 18068‐015) and then reverse transcribed using the TaqMan MicroRNA Reverse Transcription Kit (Cat. no. 4366596). Following reverse transcription, cDNA was preamplified using TaqMan PreAmp Mastermix (Cat. no. 4488593). Reverse transcription, preamplification and real‐time (RT)‐qPCR were performed according to the manufacturer's instructions as listed in Custom RT and Preamplification Pools Protocol (Life Technologies 4465407). RT‐qPCR was performed using TaqMan Universal MasterMix II, no UNG (Cat. no 4440040) with 10 μl reaction volumes on a BioRad CFX384 Touch Real‐Time PCR detection system. Gene expression was calculated using the ΔΔ threshold cycle (C T) method (Livak & Schmittgen, 2001) using aTaqMan microRNA Control Assay, small nucleolar RNA U87 (Cat. no. 4427975) as a housekeeping gene. Importantly, the absolute C T for U87 was unaffected by age or loading. A 1 day reloading time point was included in this analysis due to initial observations of increased dystrophin protein at 3 days of reloading. All microRNA data is expressed relative to the adult control baseline time point, group average ± standard error of the mean (SEM).

Histology

Serial cross‐sections (10 μm) were cut from the TA and PLN using a Leica CM 3050S cryostat (Leica Microsystems, Buffalo Grove, IL, USA). A standard haematoxylin and eosin (H & E) protocol was used to visualize centralized nuclei. Nuclei were analysed from four images (taken from four distinct regions) per muscle, per animal. The total number of fibres within each image was then counted and the data were expressed as the percentage of fibres with centralized nuclei. Overall, ≥250 fibres were counted per muscle, per animal. Images for Baseline, 3 days and 7 days of reloading were selected due to alterations in centralized nuclei at these time points and the impact of early reloading on cytoskeleton and transmembrane protein alterations. With regard to IgG‐positive fibres, TA muscle sections were fixed in cold acetone for 5 min at −20°C, followed by three 5 min phosphate‐buffered saline washes. Sections were then incubated with Alexa Fluor 488‐conjugated goat anti‐rat immunoglobulin G (IgG; [H + L]; 1:100, Life Technologies, Carlsbad, CA, USA) for 1 h at room temperature (RT). After three 5 min phosphate‐buffered saline washes, slides were cover‐slipped using ProLong Gold Antifade reagent with 4',6‐diamidino‐2‐phenylindole (DAPI; Life Technologies). Slides were imaged on a Zeiss Axio Imager.M1 fluorescent microscope using the EC Plan‐Neofluar 20× objective. For comparative analysis, exposure length remained fixed for all samples. Images were analysed using FIJI software. Six animals were analysed per group (GFP and Sponge), for both control (non‐stimulated contralateral leg) and stimulated TA muscles at 24 h post‐stimulation. IgG‐positive fibres were determined using greyscale images for the analysis. This allowed the positive fibres to be identified by the presence of white to grey colouring inside the fibre, whereas IgG‐negative fibres remained completely black. Blinded analysis was performed for the number of IgG‐positive fibres (those showing IgG staining in the cytoplasm) and these data are used to indicate fibres with damaged membranes (Janssen et al. 2014; Hughes et al. 2017).

Statistical analysis

All data for unloading and reloading time points were analysed using two‐way ANOVA using GraphPad Prism software (GraphPad Software, Inc., La Jolla, CA, USA). Tukey's post hoc analysis was used to determine differences when interactions existed. Statistical significance was set at P < 0.05. Statistical significance is presented as follows on all figures: * P < 0.05 between age groups, † P < 0.05 vs. old baseline within group, # P < 0.05 vs. adult baseline within group. For the in vivo electroporation study,a two‐way ANOVA was performed to assess the effect of plasmids and stimulation. Tukey's post hoc analysis was used to determine differences when interactions existed. Statistical significance is presented as follows: § P < 0.05 vs. GFP control muscle, P < 0.05 vs. stimulated sponge muscle. For statistical analysis of percentage of IgG‐positive fibres, an unpaired Student's t test was performed. All data are presented as mean ± SEM.

Results

Effect of reloading on dystrophin protein levels in old muscle

In the MG muscle, no significant changes in dystrophin protein were observed in either adult or aged muscles over the period of unloading and reloading (Fig. 1 A and B). In the TA muscle, unloading did not significantly alter dystrophin levels at either age. However, at 3 days of reloading, dystrophin protein increased 2.79‐fold over control levels in the adult TA muscle without changing in the old muscle (interaction P ≤ 0.03; P ≤ 0.05 vs. old muscle, Fig. 1 A and D). By 7 and 14 days of reloading, dystrophin protein was similar in the adult and aged TA muscles (Fig. 1 D).

Figure 1. Disconnect between alterations in dystrophin protein levels and mRNA expression for adult and aged skeletal muscle during reloading.

Figure 1

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A, representative western blots for dystrophin protein and Ponceau‐stained membranes used for normalization from adult and aged medial gastrocnemius (MG) (B) and tibialis anterior (TA) (D) muscles over the unloading and reloading time course (TA muscle, interaction P ≤ 0.03; main effect for age and time P ≤ 0.001). Dystrophin expression was determined by RT‐qPCR over the time course in adult and aged MG (C) and TA muscles (E) (MG muscle, main effect for time and age P ≤ 0.001; TA muscle, main effect for age P ≤ 0.001). For adult and aged muscles n = 6/time point. * P < 0.05 between age groups, † P < 0.05 vs. old baseline within group, # P < 0.05 vs. adult baseline within group. Data are presented as mean ± SEM.

Effect of reloading on dystrophin mRNA levels in old muscle

Unlike dystrophin protein, dystrophin mRNA levels in the ankle extensor (MG) started higher (2.48‐fold) in the old and increased over the time course, peaking after 7 days of reloading in both the adult and old muscles (P ≤ 0.001 main effect for time and age, Fig. 1 C). As in the MG, the old TA muscle showed 1.95‐fold greater dystrophin mRNA expression at baseline; however, dystrophin mRNA was unchanged throughout the unloading–reloading time course (both age groups; P ≤ 0.001 main effect for age, Fig. 1 E). The disconnect between the dystrophin protein levels and mRNA expression over the time course provides evidence that dystrophin protein levels are controlled post‐transcriptionally in a load‐ and age‐dependent manner.

Dystromirs contribute to the load‐ and age‐dependent regulation of dystrophin protein

In both the MG and TA muscles, miR‐31 was 7‐fold higher in old muscle compared to adult counterparts (MG muscle P ≤ 0.001 main effects for age; TA muscle P ≤ 0.03 for interaction and P ≤ 0.001 main effects for age, Fig. 2 A and D). Even though miR‐31 started higher in the old muscles, there was no significant change in miR‐31 in either the adult or old MG muscles throughout unloading and reloading. In the adult TA muscle, the first day of reloading tended to decrease miR‐31 expression, whereas in the old muscle 1 day of reloading increased miR‐31 expression 64.2% (P ≤ 0.01, Fig. 2 D inset). These changes in miR‐31 levels were transient, returning to baseline levels by 3 days of reloading and remaining unchanged from day 3 to day 14. For miR‐146b, the MG and TA muscles showed small transient fluctuations in expression throughout the time course (Fig. 2 B and E). After 1 day of reloading miR‐146b expression was significantly increased in the adult MG muscle (interaction P ≤ 0.03; P ≤ 0.05 between ages at time point, Fig. 2 B), and after 3 days of reloading, miR‐146b expression was significantly elevated in old TA muscle (interaction P ≤ 0.03; P ≤ 0.01 for age effect, Fig. 2 E). miR‐374 expression in the MG was not significantly different between adult and old muscle, although there was a main effect for time (P = 0.005, Fig. 2 C). In the TA muscle, miR‐374 expression was significantly higher in the adult muscle after 7 days of reloading (interaction P ≤ 0.001; effect for age P ≤ 0.01, Fig. 2 F).

Figure 2. Effect of unloading and reloading on miRNAs (dystromirs) in adult and aged medial gastrocnemius (MG, top row) and tibialis anterior (TA, bottom row) muscles.

Figure 2

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Expression of miR‐31 (A and D), miR‐146b (B and E) and miR‐374 (C and F) determined by RT‐qPCR and quantified using the ΔΔC T method with U87 used as a housekeeping gene. The inset in D reflects the percentage change in miR‐31 levels between day 1 of reloading (REL) and day 14 of unloading (UL). For adult and aged muscles n = 5–6/time point. In A and D there is a significant difference between age groups at all times P < 0.05. In B and E there is an interaction P ≤ 0.03 and main effect for age P ≤ 0.01. In F there is an interaction P ≤ 0.001 and main effect for age P ≤ 0.01. * P < 0.05 between age groups, † P < 0.05 vs. old baseline within group, # P < 0.05 vs. adult baseline within group. Data are presented as mean ± SEM.

Evidence for compensatory changes in other cytoskeleton proteins due to a lack of dystrophin in old tibialis anterior muscle

β‐Dystroglycan showed no effect of age in the MG muscle (main effect for time P ≤ 0.01, Fig. 3 A); however, in the TA muscle β‐dystroglycan was 4‐fold higher at 7 days of reloading (interaction P ≤ 0.02; main effect for age and time P ≤ 0.001, Fig. 3 D). The large intermediate filament protein, desmuslin, displayed the greatest alterations between unloading and reloading. In the MG muscle, desmuslin levels significantly decreased in adult rats after 14 days of unloading (interaction P ≤ 0.04; main effect for age and time P ≤ 0.05, Fig. 3 B); however, the changes in the MG were minor compared with the TA muscle. In the TA muscle, desmuslin levels increased following 7 days (6‐fold) and 14 days (9‐fold) of reloading in the old animals, whereas no significant change in desmuslin was observed in the adults (interaction P ≤ 0.001; main effect for age and time P ≤ 0.001, Fig. 3 E). The Z‐disc muscle LIM protein (MLP) showed a similar pattern, increasing significantly in old TA muscle compared to adult counterparts after 7 and 14 days of reloading (interaction P = 0.06; main effect for age and time P ≤ 0.05, Fig. 3 F). In contrast, no difference in MLP was observed in the MG muscle (Fig. 3 C). The fact that these proteins changed only in the old TA at 7 and 14 days suggest a potential compensatory response within the DGC, intermediate filaments and myofibrillar Z‐line for the lack of upregulation of dystrophin protein in old TA muscle during the early period of reloading.

Figure 3. Compensatory increases in cytoskeleton proteins that link the extracellular matrix to the Z‐line and thick filament in adult and aged medial gastrocnemius (MG, top row) and tibialis anterior (TA, bottom row) muscles during unloading/reloading.

Figure 3

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Levels of β‐dystroglycan (A and D), desmuslin (B and E) and muscle LIM protein (C and F) over the time course of unloading/reloading. For adult and aged muscles n = 6/time point. In A there is a main effect for time P ≤ 0.01. In D there is an interaction P ≤ 0.02 and main effect for age and time P ≤ 0.001. In B there is an interaction P ≤ 0.04 and main effect for age and time P ≤ 0.05. In E there is an interaction P ≤ 0.001 and main effect for age and time P ≤ 0.001. In F there is an interaction P = 0.06; main effect for age and time P ≤ 0.05. * P < 0.05 between age groups, † P < 0.05 vs. old baseline within group, # P < 0.05 vs. adult baseline within group. Data are presented as mean ± SEM.

Effect of unloading and reloading on other members of the DGC, integrin family and Z‐line proteins

Examination of extracellular components of the DGC revealed that laminin 2‐α was significantly elevated after 7 days of reloading in both the old MG (interaction P ≤ 0.03; main effect for age and time P ≤ 0.001, Fig. 4 A) and TA (main effect for age and time P ≤ 0.001, Fig. 4 D) muscles compared to adult counterparts. Two membranous components of the DGC, α‐sarcoglycan and sarcospan, were significantly elevated in old MG muscle compared to adult counterparts after 3, 7 and 14 days of reloading (Fig. 4 B and C). Similar to the MG muscle, α‐sarcoglycan was significantly elevated after 3 and 14 days of reloading in the old TA compared to adult counterparts (main effect for age P ≤ 0.01, Fig. 4 E), whereas sarcospan was significantly elevated at all reloading time points in the old TA muscle compared to adult rats and was also significantly increased above old baseline at 3 and 14 days of reloading (interaction P ≤ 0.001; main effect for age and time P ≤ 0.001, Fig. 4 F).

Figure 4. Dystrophin‐associated glycoprotein complex (DGC) protein levels in adult and aged medial gastrocnemius (MG, top row) and tibialis anterior (TA, bottom row) muscles during unloading/reloading.

Figure 4

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Western blots were performed to assess for alterations in laminin‐2α (A and D), α‐sarcoglycan (B and E) and sarcospan (C and F) in the MG and TA muscles over the time course of unloading/reloading. In A there is an interaction P ≤ 0.03 and main effect for age and time P ≤ 0.001. In D there is a main effect for age and time P ≤ 0.001. In E there is a main effect for age P ≤ 0.01. In F there is an interaction P ≤ 0.001 and main effect for age and time P ≤ 0.001. For adult and aged muscles n = 6/time point. * P < 0.05 between age groups, † P < 0.05 vs. old baseline within group. Data are presented as mean ± SEM.

Since the family of integrin proteins forms a parallel pathway for force to be transmitted laterally and have been observed to play a role in protection from contraction‐induced muscle injury (Boppart et al. 2006; Rooney et al. 2006; Liu et al. 2012; Marshall et al. 2012), we sought to address the impact of unloading and reloading on β1‐integrin expression in old muscle. In the MG muscle, there was a main effect for both time (P ≤ 0.01) and age (P ≤ 0.02), but no interaction between factors was observed (Fig. 5 A). On the other hand, in the TA muscle, a main effect was only observed for time (P ≤ 0.01; Fig. 5 D). The pattern of β1‐integrin protein content closely matched that of dystrophin in the adult TA muscle, with a peak at 3 days of reloading. However, unlike dystrophin, the old animals showed a similar increase in β1‐integrin at this early reloading time point. Desmin, which connects the integrins with the contractile machinery, was significantly higher in the old MG (interaction P ≤ 0.001; main effect for age P ≤ 0.001) and TA (interaction P ≤ 0.001; main effect for age and time P ≤ 0.05) muscle across all reloading time points compared to adult rats (Fig. 5 B and E). The response of α‐actinin in the TA muscle mimicked what was observed for β‐dystroglycan and desmuslin, showing a significant increase in old muscle following 7 and 14 days of reloading (main effect for age and time P ≤ 0.001; Fig. 5 F).

Figure 5. Integrin, intermediate filament and Z‐line protein levels in adult and aged medial gastrocnemius (MG, top row) and tibialis anterior (TA, bottom row) muscles during unloading/reloading.

Figure 5

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Western blots were performed to assess for alterations in β1‐integrin (A and D), desmin (B and E) and α‐actinin (C and F) in the MG and TA muscles over the time course of unloading/reloading. In A there is a main effect for age and time P ≤ 0.02. In B there is an interaction P ≤ 0.001 and main effect for age P ≤ 0.001. In E there is an interaction P ≤ 0.001 and main effect for age and time P ≤ 0.05. In F there is a main effect for age and time P ≤ 0.001. For adult and aged muscles, n = 6/time point. * P < 0.05 between age groups, and † P < 0.05 vs. old baseline within group. Data are presented as mean ± SEM.

Effect of unloading and reloading on muscle injury and markers of membrane repair/regeneration

To determine whether the delayed dystrophin protein response in the old TA muscle made the muscle more prone to membrane damage and subsequent regeneration, we determined the expression of genetic indicators of muscle regeneration (Cyclin D2 and Cyclin A2) and membrane repair (dysferlin) in the MG and TA, as well as the gold standard measure of regenerating muscle fibres – centralized nuclei (PLN and TA) – over the time course (Fig. 6). In the MG, dysferlin mRNA was only significantly elevated in old muscle at baseline and after 14 days of reloading (main effect for age and time P ≤ 0.004, Fig. 6 B) compared to adult muscle (Fig. 6 B). The old TA muscle showed alterations in dysferlin mRNA with early reloading resulting in significant elevations, whereas no change was evident in adult muscle over the time course (interaction P ≤ 0.001; main effect for age and time P ≤ 0.001, Fig. 6 C). There was no difference in Cyclin A2 mRNA expression between the adult and old MG muscle during unloading or reloading (Fig. 6 D). Cyclin A2 expression was significantly increased in the old TA throughout the reloading time period and was significantly higher compared to adult rats at 7 and 14 days of reloading (interaction P ≤ 0.001; main effect for age and time P ≤ 0.001, Fig. 6 E). For Cyclin D2 mRNA expression, there was no significant difference between adult and old MG muscle over the time course (data not shown). As with Cyclin A2, the old TA muscle displayed significantly higher Cyclin D2 at 7 and 14 days of reloading (data not shown). In the PLN muscle, old animals had more centralized nuclei than adult; however, there was no effect of either unloading or reloading in either age group (main effect for age P ≤ 0.001, Fig. 6 F). In contrast, 7 and 14 days of reloading in the old TA muscle resulted in significant (∼2‐fold increases, P ≤ 0.01 vs. Adult) increases in the proportion of centralized nuclei (interaction P ≤ 0.02; main effect for age and time P ≤ 0.01, Fig. 6 G).

Figure 6. Markers of membrane damage/injury and regeneration in adult and aged medial gastrocnemius or plantaris (MG or PLN, top row) and tibialis anterior (TA, bottom row) muscles during unloading/reloading.

Figure 6

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Representative images for the TA muscle H&E used to perform centralized nuclei quantification, with baseline, 3 and 7 days of reloading presented (A). Nuclei were analysed from 4 images per muscle (tibialis anterior (TA) and plantaris (PLN)), per animal, per time point (scale bar = 100 μm). Data are reported as a percentage of fibres with centralized nuclei (F and G). B and C, dysferlin mRNA expression for both muscles and age groups during unloading and reloading. D and E, Cyclin A2 expression over the time course. For adult and aged muscles, n = 6/time point. In B, there is a main effect for age and time P ≤ 0.004. In C there is an interaction P ≤ 0.001 and main effect for age and time P ≤ 0.001. In E there is an interaction P ≤ 0.001 and main effect for age and time P ≤ 0.001. In F there is a significant difference between age groups at all times P ≤ 0.05. In G there is an interaction P ≤ 0.02 and main effect for age and time P ≤ 0.01. * P < 0.05 between age groups, and † P < 0.05 vs. old baseline within group. Data are presented as mean ± SEM.

miR‐31 inhibition increases dystrophin and partially protects old skeletal muscle from contraction‐induced muscle injury

As a proof of concept experiment and to further explore the potential interaction between miR‐31, dystrophin and contraction‐induced injury in ageing skeletal muscle, we electroporated a miR‐31 sponge into the TA muscle of old rats 1 week prior to a single bout of lengthening contractions. Incorporation of the empty control vector (GFP) and miR‐31 sponge plasmids into skeletal muscle fibres was confirmed by histochemical and biochemical analyses (Fig. 7 A). The sponge‐treated muscles displayed an increase in dystrophin protein compared to GFP‐treated animals (main effect for plasmid P = 0.007 and stimulus P ≤ 0.01; Fig. 8 G) that varied with electroporation efficiency. There was no significant difference in dystrophin mRNA or miR‐31 expression between GFP and sponge‐treated TA muscles (Fig. 8 F and I). In addition, Dicer protein (another 3′UTR target for miR‐31) was also elevated in the sponge‐treated animals (main effect for plasmid P < 0.001; Fig. 8 H), confirming the effectiveness of the miR‐31 sponge.

Figure 7. Confirmation of plasmid incorporation into skeletal muscle fibres and similar contractile activity between groups with the stimulation protocol.

Figure 7

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Electroporation efficiency was confirmed histochemically (magnification 20×; GFP control TA muscle presented) and biochemically through western blot for the presence of GFP protein (n = 8/group; A). GFP protein has a molecular mass of 27 kDa. Muscle mass was determined 24 h post‐stimulation protocol (B). In B there is a main effect for stimulus P ≤ 0.02. Representative western blot and quantification of phosphorylation of upstream binding factor (UBF) at serine 637 site (n = 6–8/group; C). [Color figure can be viewed at http://wileyonlinelibrary.com]

Figure 8. miR‐31 inhibition partially protects aged skeletal muscle from contraction‐induced muscle injury via upregulation of dystrophin protein.

Figure 8

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Representative images for the control and stimulated TA muscles from GFP‐ and sponge‐treated animals stained for immunoglobulin (IgG). Visible membrane damage (IgG‐positive fibres) 24 h after contraction‐induced muscle injury. White arrows indicate IgG‐positive muscle fibres; (A). Level of muscle membrane damage was quantified as increases in percentage of IgG‐positive fibres; n = 6/group (B). C–E, mRNA expression for key markers of the membrane repair process (dysferlin, MG‐153 and annexin A2; for all markers an interaction P ≤ 0.02; main effect for stimulus P ≤ 0.01 was observed). F, dystrophin mRNA expression in GFP‐ and sponge‐treated TA muscles. Quantification of 3′UTR targets for miR‐31, dystrophin (G) and dicer (H) protein content in GFP‐ and sponge‐treated TA muscles. For dystrophin protein, main effect for stimulus P ≤ 0.01 and plasmid P ≤ 0.001 was observed. A main effect for plasmid P ≤ 0.001 was observed in dicer protein content. I, miR‐31 expression in GFP‐ and sponge‐treated TA muscles. Representative western blots for dystrophin (DYS) and dicer protein (J). § P < 0.05 vs. GFP control muscle, and  P < 0.05 vs. stimulated sponge muscle. n = 6–8/group. Data are presented as mean ± SEM. [Color figure can be viewed at http://wileyonlinelibrary.com]

Seven days following electroporation, the right leg underwent a 20 min bout of lengthening contractions to induce injury. A similar magnitude of muscle contraction was confirmed via phosphorylation of upstream binding factor (UBF), as we have previously observed elevations in this marker 24 h after this lengthening contraction protocol (West et al. 2016). UBF phosphorylation increased 3‐fold in both the GFP‐ and sponge‐treated animals (P = n.s.; Fig. 7 C), suggesting a similar contractile stimulus. There was no significant difference in muscle mass between control and stimulated muscles in sponge‐treated animals after 24 h (Fig. 7 B). There was, however, a significant increase in muscle mass in the stimulated muscles of GFP‐treated animals (main effect for stimulus P ≤ 0.02; Fig. 7 B), suggesting oedema. Assessment of IgG‐positive muscle fibres, an indicator of muscle membrane damage, indicated that the sponge‐treated TA muscles displayed significantly less contraction‐induced membrane injury than GFP controls (Fig. 8 A and B). To confirm the lower membrane disruption, we measured expression of the membrane repair genes dysferlin, MG‐153 and annexin A2. In GFP‐stimulated muscles, dysferlin, MG‐153 and annexin A2 expression were all significantly elevated compared with GFP control (interaction P ≤ 0.02; main effect for stimulus P ≤ 0.01; Fig. 8 C–E). Sponge‐treated muscles did not increase dysferlin, MG‐153 and annexin A2 mRNA expression in response to contraction (P = n.s; Fig. 8 C–E).

Discussion

Ageing muscle displays an inability to fully recover muscle size and strength following a prolonged period of disuse (Suetta et al. 2009, 2013; Baehr et al. 2016). Here we provide evidence that the loss of lateral force transmission makes old muscles, especially flexor muscles, more susceptible to contraction‐induced injury during the early reloading phase following a period of disuse. Specifically, we demonstrate that there is lower dystrophin protein in old TA muscles and a delay in the response of dystrophin to reloading that is unique to the flexor muscles of old animals. We also identify alterations in microRNAs, known to target dystrophin, over the time course of unloading and reloading, which may contribute to the post‐transcriptional differences in dystrophin protein levels in aged and adult muscle. Our proof of concept experiment using in vivo electroporation to decrease miR‐31 activity highlights the impact of this microRNA on dystrophin content and the critical role that dystrophin plays in protecting muscle fibres from contraction‐induced muscle injury. The lack of dystrophin protein in old TA muscles, both at baseline and in response to reloading, is accompanied by increases in β‐dystroglycan, α‐sarcoglycan, sarcospan, desmuslin and muscle LIM protein (all linked to mechanoprotection), as well as increased markers of membrane repair and a greater number of fibres with centralized nuclei, suggest an attempt to compensate for the loss of dystrophin protein that is insufficient to prevent membrane damage and regeneration upon reloading in aged rats.

We have previously observed that the dystrophin content of muscle differs as a function of loading history and age (Hughes et al. 2017). In healthy adult flexor muscles, dystrophin protein levels can be 65% lower than an equivalent extensor muscle. Further, the amount of dystrophin in the flexors drops another ∼60% between 9 and 28 months. In the current study, dystrophin protein levels were unaffected by loading in the gastrocnemius muscle, which started with high dystrophin levels in both the adult and old. In the TA muscle, however, during the early reloading phase (first 3 days) the adult rats showed a 2.9‐fold increase in dystrophin protein that did not occur in the aged muscle. It is our assertion that the prolonged increase in dystrophin protein in the adult muscles is the result of a transient drop in miR‐31 levels 1 day into reloading. A transient decrease in miR‐31 can have a prolonged effect on dystrophin protein levels due to the long half‐life of dystrophin protein. In mice, following knockdown of dystrophin mRNA, dystrophin protein remains detectable 12 weeks (Ghahramani Seno et al. 2008) or 26 weeks (Ahmad et al. 2000) later, indicating that acute increases in translation can have the persistent effects on dystrophin protein that we see following reloading. Across all of the other cytoskeletal proteins that were measured during either unloading or reloading, the increase in dystrophin was the only change that was greater in the adult muscles; that is, in general, force transmission‐related proteins increased to a greater degree in old muscle, especially during reloading. There was a similar reloading‐induced increase in β1‐integrin at day 3 in adult and old, indicating that some cytoskeletal proteins respond normally to loading. Dystrophin and β1‐integrin are suggested to play a role in protecting the sarcolemma from damage, maintaining membrane integrity and preventing contraction‐induced muscle injury (Marshall et al. 2012; Gao & McNally, 2015). Therefore, the delayed rise in dystrophin protein in the old muscle may make the TA more susceptible to contraction‐induced injury and membrane damage immediately upon reloading following an unloading period. Four pieces of data from the current study support this hypothesis, and show the progression of early injury during reloading. First, there was a 2.6‐fold increase in dysferlin mRNA at day 3 of reloading only in the old TA muscle, indicating an acute increase in membrane damage. Second, there was a 40% drop in desmin protein in the old TA between days 3 and 7 of reloading. Desmin protein decreases following load‐induced muscle damage (Barash et al. 2002), suggesting more load‐induced injury in the old TA. Third, the TA muscle from old rats showed a 2.9‐fold (3 days) and 5.1‐fold (7 days) increase in Cyclin A2 gene expression, a marker that is associated with the early phase of muscle injury (Yan et al. 2003). Fourth, the TA muscles from the old animals demonstrated a 2‐ and 1.6‐fold increase in fibres with centralized nuclei at days 7 and 14, respectively. Centralized nuclei are the gold standard indicator of regenerating muscle fibres (Folker & Baylies, 2013). Together, these data indicate that immediately upon reloading, aged muscle displayed greater membrane disruption and muscle damage followed by remodelling and repair. The old muscles were able to increase dystrophin protein by 7 days of reloading, suggesting that the dystrophin response is delayed but not lost in pre‐sarcopenic muscle. However, whether the impaired dystrophin protein response to loading is more pronounced in sarcopenic animals remains to be determined.

The observed 2.6‐fold increase in dysferlin mRNA at day 3 of reloading only in the TA muscle from old rats is striking. Dysferlin is a key member of a muscle‐specific repair complex involved in the response to membrane damage and is pivotal in the patch repair of membranes disrupted by mechanical stress (Han & Campbell, 2007). Although the loss of force is a gold standard measure of contraction‐induced muscle injury, a secondary marker is the loss of sarcolemma integrity, leading to lesions on injured skeletal muscle that are frequently focal (Tidball, 2011). The elevation in dysferlin expression is indicative of membrane disruption/damage and subsequent repair (Han & Campbell, 2007). It is important to note that the changes in dysferlin expression occurred in response to both a physiological stimulus (unloading/reloading) and the more extreme eccentric contraction‐induced model. Indeed, using a lengthening contraction protocol, we observed dysferlin mRNA expression to be elevated (2‐fold) after 24 h. A similar 2‐fold increase in dysferlin mRNA was observed at 3, 7 and 14 days of reloading. The similar degree of dysferlin upregulation may indicate that high force is not required for contraction‐induced muscle injury in old animals. The subsequent remodelling and regeneration observed here in response to the return to normal loading might be more clinically relevant than other experimental models (i.e. cardiotoxin, or freeze injury) more commonly used to study muscle injury and repair. Further, it should be noted that the greater reloading‐induced injury in the old animals occurred even though these animals demonstrated reduced cage activity compared to their adult counterparts (Baehr et al. 2016). Together, these data indicate that old muscles that are low in dystrophin are more prone to contraction‐induced muscle injury during reloading.

In both the MG and TA there was approximately 2‐fold more dystrophin mRNA in the aged muscle compared with the adult; however, dystrophin protein levels did not reflect these differences in expression. Reloading resulted in a progressive increase in dystrophin mRNA in the MG that peaked at 7 days (4.4‐fold in the adult and 2.7‐fold in the old) before returning towards baseline at 14 days (Fig. 1). In spite of this significant rise in dystrophin mRNA in the MG, there was no change in dystrophin protein levels during the reloading time course. In contrast, there was a rapid increase in dystrophin protein in the TA muscle that peaked at 3 days of reloading that was not paralleled by an increase in dystrophin mRNA. These observations suggest that dystrophin protein levels are determined through a post‐transcriptional mechanism that is modified by age and loading. One post‐transcriptional change that may explain the disconnect between dystrophin mRNA and protein is the expression of specific microRNAs (dystromirs). In the aged TA muscle there was a significant spike (∼70%) at day 1 of reloading, whereas in the adult muscle there was a decline (∼30%) in miR‐31 expression. Together with the respective changes in protein at the subsequent time point, no change in the old and a 2.9‐fold increase in the adult, these data suggest that dystrophin levels in muscle are dynamically controlled post‐transcriptionally and that this process is impaired with ageing. Even though changes in miR‐31 appear to be predictive of dystrophin protein, other dystromirs that were not measured in the present study may also influence dystrophin protein levels. MicroRNAs, such as miR‐222, miR‐223, miR‐34c and miR‐449, that are elevated in both mdx mice and human muscular dystrophies (Greco et al. 2009; Fiorillo et al. 2015), may also contribute to alterations in dystrophin in aged skeletal muscle. Future studies are needed to determine which microRNAs are altered by age and loading and the mechanism underlying the increase in miR‐31 expression with ageing.

Previous work by Caccahiarelli and colleagues (2011) observed a reduction in dystrophin coinciding with elevated levels of miR‐31 in both mdx and Duchenne muscular dystrophy muscle samples. In addition, the authors showed, using in vitro studies, the elevation of dystrophin protein with the addition of a miR‐31 sponge plasmid. Through in vivo electroporation, we were able to test the hypothesis that miR‐31 alters dystrophin protein content in aged skeletal muscle. The miR‐31 sponge sequesters endogenous miR‐31 away from its normal mRNA targets allowing those targets to be translated. Since miRNA sponges work by sequestering and not reducing mRNA content (Ebert & Sharp, 2010), it is not surprising that we see an increase in dystrophin protein without a change in miR‐31 or dystrophin mRNA levels. Supporting the role of miR‐31 in controlling dystrophin protein, the miR‐31 sponge increased dystrophin protein in old muscles. This increase in dystrophin partially protected aged skeletal muscle from contraction‐induced muscle injury, as evidenced by the reduced number of IgG‐positive fibres and no significant change in markers of membrane repair 24 h post‐stimulation. It is interesting to note that following stimulation there was a similar reduction in dystrophin protein in both GFP‐ and sponge‐stimulated muscles. The acute reduction of dystrophin protein may reflect dystromirs other than miR‐31 (Fiorillo et al. 2015) or the acute injury (Neri et al. 2007; van Putten et al. 2012, 2013). Our finding of improved dystrophin levels in muscle following treatment with a miR‐31 sponge is in contrast to work from Hildyard and Wells (2016) who found that, in mdx mice, a miR‐31 sponge in combination with exon skipping did not increase dystrophin protein more than exon skipping alone. The work of Hildyard and Wells (2016) contrasts not only with the current study but with the work of others (Cacchiarelli et al. 2011; Fiorillo et al. 2015). It is possible that the skipped message produced by Hildyard and Wells (2016) does not contain the same 3'UTR needed for miR‐31 to exert control over the rate of dystrophin translation. Overall, our data add to the growing literature providing insight into the interaction between miR‐31 and dystrophin and the impact of decreased dystrophin protein on skeletal muscle function with age.

In response to the delayed upregulation of dystrophin protein in the old muscles, a number of other cytoskeletal proteins (β‐dystroglycan, sarcospan, desmuslin, α‐actinin and muscle LIM protein) are upregulated during the reloading phase. Interestingly, the proteins that are upregulated form a chain linking the cell membrane to the Z‐line and thick filament. β‐Dystroglycan, through its partner α‐dystroglycan, connects dystrophin to the extracellular matrix. Further, β‐dystroglycan has recently been observed to play a key role in stabilizing dystrophin and limiting the amount of contraction‐induced muscle injury (Rader et al. 2016). Desmuslin is an intermediate filament that links the dystrophin binding protein α‐dystrobrevin to desmin. Desmuslin is an important mechanical linker between dystrophin and the Z‐line (Mizuno et al. 2001). As would be expected for a lateral force transmission protein, desmuslin knockout mice are more prone to injury following lengthening contractions and show a loss of force production and mean fibre diameter at baseline (Garcia‐Pelagio et al. 2015). α‐Actinin, the protein that mechanically links the thick and thin filaments within the Z‐line (Sjöblom et al. 2008), is also upregulated more in the old TA muscle at 7 days of reloading. Finally, muscle LIM protein, the structural connection between α‐actinin and the thick filament (Pomies et al. 1997), is increased only in the aged TA muscle. Muscle LIM protein also has the capacity to act as a mechanical sensor and a transcriptional factor by shuttling between the cytoplasm and nucleus (Arber et al. 1994). For example, 6 h after acute resistance exercise muscle LIM protein increases approximately 4.3‐fold over resting levels (Chen et al. 2002). However, given that the increase in muscle LIM protein occurs only in the aged TA muscle and that the increase parallels changes in β‐dystroglycan, desmuslin and α‐actinin, these data suggest that the increase in muscle LIM protein is due to its role in force transfer and not mechanical signalling. Overall, the increase in cytoskeletal proteins linking dystrophin to both the extracellular matrix and the thick and thin filaments in the aged TA muscle may represent an attempt to compensate for the delayed dystrophin response. On the other hand, the 7 and 14 day reloading response of desmin, α‐actinin, desmuslin and muscle LIM could be indicative of skeletal muscle remodelling during the regeneration and repair of the aged muscle. Regardless, given the greater membrane damage and accentuated injury in the old TA, these compensatory responses are not sufficient to protect the muscle from the loss of dystrophin.

Summary

The force transmission apparatus of pre‐sarcopenic animals shows altered responses following a period of unloading and reloading. Specifically, adult ankle flexor muscles can rapidly increase dystrophin protein in response to reloading and this rapid response is lost in old animals. We demonstrate that the amount of dystrophin protein in a muscle is negatively regulated through a post‐transcriptional mechanism that is exacerbated with ageing. The core of this response is mediated by alterations in the dystrophin‐targeting microRNA miR‐31, which is increased ∼7‐fold with ageing. The failure of old flexor muscles to rapidly increase dystrophin protein in response to reloading is associated with a nearly 2‐fold increase in miR‐31 upon reloading and is followed by an increase in proteins that link the extracellular matrix to the Z‐line and thick filament (β‐dystroglycan, sarcospan, desmuslin, α‐actinin and muscle LIM protein). This potential compensation was evident at baseline (Hughes et al. 2017); however, following reloading the compensatory effects are increased 3‐ to 4‐fold. Overall, this response is not sufficient to prevent membrane damage early in the reloading process. However, when miR‐31 activity is decreased in aged muscle using a miR‐31 sponge, dystrophin protein increased and contraction‐induced skeletal muscle injury decreased. While the regulation of dystrophin in the extensor muscles needs to be investigated further, these data suggest that miR‐31 is a novel target for the protection of old flexor muscles from contraction‐induced muscle injury.

Additional information

Competing interests

None declared.

Author contributions

Conception and design of the experiments: D.C.H., D.W.D.W., L.M.B., S.C.B. and K.B. Collection, analysis and interpretation of data: D.C.H., G.R.M., L.M.B., D.W.D.W., A.G.M., S.M.E., A.D., C.M.A., S.C.B. and K.B. Drafting the article or revising it critically for important intellectual content: D.C.H., G.R.M., L.M.B., D.W.D.W., A.G.M., S.M.E., A.D., C.M.A., S.C.B. and K.B. All authors have approved the final version of the manuscript and agree to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for authorship are listed.

Funding

D.W.D.W. was supported by a postdoctoral fellowship from the Natural Sciences and Engineering Research Council of Canada. This work was supported by a project grant to K.B. from the National Institute on Aging of the National Institutes of Health under award number R01AG045375 and a Merit Award from the United States Department of Veterans Affairs to S.C.B. under award number VARR&D E7766‐R.

Biography

Through previous mentorships from Dr Adam Sharples, Prof. Keith Baar and currently with Prof. Sue Bodine at the University of Iowa, David C. Hughes’s work has utilized in vitro and in vivo models to target novel areas of muscle physiology relating to ageing and disease. Recent work has focused on the impact of the cytoskeleton protein network in ageing, atrophy and injury. David hopes to obtain a tenure‐track faculty position to mentor potential scientists of tomorrow and develop an independent research programme in skeletal muscle physiology.

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Edited by: Scott Powers & Anne McArdle

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