This study utilized a rat synergist ablation model to integrate changes in single muscle fiber contractility, extracellular matrix composition, activation of important signaling pathways in muscle adaption, and corresponding changes in the muscle transcriptome to provide novel insight into the basic biological mechanisms of muscle fiber hypertrophy.
Keywords: permeabilized muscle fibers, contractility, atrophy, hypertrophy, extracellular matrix
Abstract
Skeletal muscle can adapt to increased mechanical loads by undergoing hypertrophy. Transient reductions in whole muscle force production have been reported during the onset of hypertrophy, but contractile changes in individual muscle fibers have not been previously studied. Additionally, the extracellular matrix (ECM) stores and transmits forces from muscle fibers to tendons and bones, and determining how the ECM changes during hypertrophy is important in understanding the adaptation of muscle tissue to mechanical loading. Using the synergist ablation model, we sought to measure changes in muscle fiber contractility, collagen content, and cross-linking, and in the expression of several genes and activation of signaling proteins that regulate critical components of myogenesis and ECM synthesis and remodeling during muscle hypertrophy. Tissues were harvested 3, 7, and 28 days after induction of hypertrophy, and nonoverloaded rats served as controls. Muscle fiber specific force (sFo), which is the maximum isometric force normalized to cross-sectional area, was reduced 3 and 7 days after the onset of mechanical overload, but returned to control levels by 28 days. Collagen abundance displayed a similar pattern of change. Nearly a quarter of the transcriptome changed over the course of overload, as well as the activation of signaling pathways related to hypertrophy and atrophy. Overall, this study provides insight into fundamental mechanisms of muscle and ECM growth, and indicates that although muscle fibers appear to have completed remodeling and regeneration 1 mo after synergist ablation, the ECM continues to be actively remodeling at this time point.
NEW & NOTEWORTHY This study utilized a rat synergist ablation model to integrate changes in single muscle fiber contractility, extracellular matrix composition, activation of important signaling pathways in muscle adaption, and corresponding changes in the muscle transcriptome to provide novel insight into the basic biological mechanisms of muscle fiber hypertrophy.
skeletal muscle displays a profound ability to adapt to increased mechanical loads transmitted across joints by undergoing hypertrophy. This adaptation involves a coordinated response of both the muscle fibers that generate force, and the extracellular matrix (ECM), which transmits this force to tendons and bones (14, 22). Forces are transmitted longitudinally at myotendinous junctions through fibrillar collagens, as well as laterally between muscle fibers through basement membrane collagens before reaching the tendon (18, 32). There are both injury-independent and injury-dependent mechanisms of muscle fiber and ECM adaptation to different forms of exercise. For injury-independent mechanisms, repetitive mechanical loading, typically in the form of endurance exercise or concentric muscle contractions, can stimulate ECM production and remodeling, with little to no changes in muscle fiber force production (5). In resistance exercise, the loads placed upon muscle during eccentric contractions induces small amounts of damage to muscle fibers and the ECM. This triggers an adaptive response in which the injured components of the muscle cells and matrix are broken down and replaced with new, and typically greater, amounts of contractile and matrix proteins, resulting in skeletal muscle hypertrophy (5, 14). When muscle is more severely injured by eccentric injuries, or in cases of immobilization or disease, muscle fibers will often undergo atrophy, which can result in a marked decline in the force-generating capacity of muscle tissue. The ECM may also wither, but more frequently, the matrix will continue to increase its volume slowly over time, a process that frequently leads to fibrosis (14, 23). Numerous biophysical, biochemical, and molecular factors participate in the process of skeletal muscle hypertrophy, and gaining a greater understanding of how these factors interact, especially in regards to the contractile and matrix elements of muscle, could help to improve treatments for severe muscle injuries and diseases.
Resistance exercise training often results in increases in whole muscle force production through muscle fiber hypertrophy, as well as an increase in the abundance and elastic energy storage capacity of the ECM (14). Myofibrils, which are composed of repeating segments of sarcomeres, are the fundamental structures within muscle fibers that generate force. Increasing the number of parallel-aligned myofibrils increases the force-generating capacity of a muscle fiber. The ratio of maximum isometric force production (Fo) of a fiber to its cross-sectional area (CSA) is referred to as specific force (sFo). Although exercise training can increase the size and force-generating capacity of muscle fibers, and healthy aging can have the opposite effect, in both cases, sFo of muscle fibers displays little to no change (4, 41, 42). When a muscle is injured, or in degenerative muscle diseases, sFo can be markedly reduced (12, 25, 30, 47). Decreases in sFo also correlate well with clinically measured parameters that predict the regenerative capacity of muscle and overall patient health (28). Although changes in sFo at the single muscle fiber level have been previously studied in long-term exercise training studies and rodent injury models, to our knowledge this process has not been studied acutely during the onset of muscle hypertrophy. The ECM of muscle also undergoes important changes during hypertrophy to optimize the transmission and storage of greater amounts of energy from larger muscle fibers. This is typically characterized by an increase in the collagen content and stiffness of the matrix, among other biochemical and mechanical changes (14). Understanding how the ECM changes is therefore critical for an integrated understanding of functional changes to muscle during the onset of muscle hypertrophy.
The rodent synergist ablation model has frequently been used in the study of skeletal muscle hypertrophy (2, 3, 9, 15, 21). This model involves the partial or full surgical removal of the Achilles tendon, resulting in a marked reduction in the contribution of the gastrocnemius and soleus muscles to ankle joint plantarflexion. The plantaris muscle, which also acts as an ankle plantarflexor, then undergoes hypertrophy to compensate for the reduction in force transmission from the gastrocnemius and soleus. The primary objectives of the current study were to use the synergist ablation model to study early and late changes in muscle fiber contractility, and markers of ECM abundance and stiffness in adult rats. Based on our existing understanding of satellite cell activity and muscle fiber regeneration, we hypothesized that fibers from the plantaris muscle would experience a reduction in sFo 3 and 7 days after synergist ablation, and that sFo would be restored to control levels by 28 days after overload. We further hypothesized that there would be a slow, progressive accumulation in the content of collagen and the abundance of collagen cross-links over the course of hypertrophy. In support of our primary objectives, our secondary objectives were to measure activation of signaling proteins that regulate fiber hypertrophy and ECM production, and analyze changes in the transcriptome of muscles subjected to synergist ablation.
METHODS
Animals and surgeries.
This study was approved by the University of Michigan Institutional Animal Care and Use Committee and followed the U.S. Public Health Service Policy on Humane Care and Use of Laboratory Animals. Six-month-old male Sprague-Dawley rats were purchased from Charles River (Wilmington, MA) and housed in specific pathogen-free conditions. Rats were randomized to serve either as controls (n = 4) or to the 3-, 7-, or 28-day group (n = 8 per group); a total of 28 rats were used. Based on previous studies in which the measured variability of control muscles was low (10), we assigned only four animals to the control group. We performed a bilateral synergist ablation procedure as previously described (2, 13, 37). In this procedure, the Achilles tendon is removed, which eliminates the ability of the gastrocnemius and soleus muscles to transmit forces longitudinally to the talocrural joint, causing hypertrophy of the synergist plantaris muscle. Removal of the Achilles tendon prevents subsequent scarring and stress shielding of the plantaris muscle, but does not totally remove the ability of the gastrocnemius and soleus muscles to transmit forces laterally to the plantaris muscle, however. Rats were anesthetized with 2% isoflurane, and the skin overlying the posterior lower limb was shaved and scrubbed with chlorhexidine and isopropyl alcohol. A midline incision was created in the skin and the paratenon was split to achieve visualization of the Achilles tendon, and a full-thickness tenectomy was performed in the midsubstance of the tendon. The entire midsubstance of the Achilles was removed to prevent spontaneous healing. The paratenon was loosely reapproximated, a splash block of a small amount of 0.5% bupivacaine was applied to the tendon stumps, and the skin was closed using 4-0 Vicryl (Ethicon, Somerville, NJ) and GLUture (Abbott, Abbott Park, IL). Ampicillin (20 mg/kg), buprenorphine (0.03 mg/kg), and carprofen (5 mg/kg) were administered to prevent infection and for analgesia during postoperative recovery. Weight bearing and cage activity were allowed ad libitum, and rats were closely monitored for signs of pain or distress.
Rats were allowed to recover for 3, 7, or 28 days after the synergist ablation procedure. At the time of harvest, rats were anesthetized with sodium pentobarbital (50 mg/kg), and plantaris muscles were harvested and blotted dry before the mass was measured. The left muscles were finely minced, divided into equal portions for biochemical and gene expression assays, and snap-frozen at −80°C. The right proximal quarter was fixed in 10% neutral-buffered formalin and embedded in paraffin. The next distal quarter was snap-frozen in Tissue-Tek (Sakura, Torrance, CA) using isopentane cooled in liquid nitrogen, and stored at −80°C until use. The distal half of the right muscle was used for permeabilized fiber contractility analysis. After tissue was removed, animals were humanely killed by anesthetic overdose and induction of bilateral pneumothorax.
Permeabilized muscle fiber contractility.
The contractile properties of permeabilized muscle fibers was performed as previously described (10, 27, 30). Briefly, bundles of fibers were dissected from plantaris muscles, placed in skinning solution for 30 min and then in storage solution for 16 h at 4°C, followed by storage at −80°C. For contractility testing, fibers were isolated from bundles that were freshly thawed on ice, placed in a chamber filled with relaxing solution, and secured to a servomotor (Model 322C; Aurora Scientific, Aurora, ON, Canada) and force transducer (403A; Aurora Scientific) using two ties of 10-0 monofilament nylon suture at each fiber end. Fiber length (Lf) was adjusted to obtain a sarcomere length of 2.5 µm using a laser diffraction measurement system. The average fiber CSA was calculated assuming an elliptical cross section, with diameters obtained at five positions along the fiber from high-magnification images at two different views (top and side). Maximum Fo was elicited by immersing the fiber in a high-[Ca2+] solution, and sFo was derived by dividing Fo by CSA. Ten to 20 type II fibers were tested from each muscle.
Histological analysis.
Muscles embedded in paraffin were sectioned at a thickness of 7 µm with a microtome and stained with hematoxylin and eosin to perform qualitative tissue assessment. For quantitative fiber size measurements, frozen muscles were sectioned at a thickness of 10 µm in a cryostat, and incubated with wheat germ agglutin (WGA) lectin conjugated to Alexa Fluor 488 (WGA-AF488; Life Technologies) to identify extracellular matrix, and with DAPI (Life Technologies) to identify nuclei. Slides were imaged using a BX-51 microscope (Olympus America, Melville, NY) outfitted with a DP-70 high-resolution digital camera. ImageJ software (National Institutes of Health, Bethesda, MD) was used to perform quantitative measurements.
Multiplex protein analysis.
A Luminex-based system was used to measure protein abundance as described (27, 38). Briefly, muscles were finely minced and placed in cold Tissue Protein Extraction Reagent (Thermo Scientific, Rockford, IL) supplemented with a protease and phosphatase inhibitor cocktail (Thermo Scientific), homogenized with a TissueRuptor (Qiagen), and vortexed for 10 min at 4°C. Tissue lysate was then spun at 12,000 g for 10 min, and the supernatant was collected and stored at −80°C. Total protein content was determined using a bicinchoninic acid protein assay (Thermo Scientific), and 50 µg of total protein was analyzed using Milliplex-magnetic bead assays (EMD Millipore, Billerica, MA). These specific total protein analytes included Smad4 and for phosphorylated proteins: p-AktS473, p-ERK1/2T185/187, p-mTORS2448, p-p38 MAPKY180/182, p-p70S6KT389/412, p-Smad2S465/467, p-Smad3S423/425, and STAT3S727. Median fluorescence intensity (MFI) values of analytes were measured in a MAGPIX system (Luminex, Austin, TX). The MFI of a particular analyte was normalized to control muscles.
Microarray and gene expression analysis.
RNA isolation and gene expression were performed as previously described (27, 37). The muscle was finely homogenized in QIAzol (Qiagen, Valencia, CA), and RNA was isolated using a miRNeasy Kit (Qiagen), and treated with DNase I (Qiagen).
Microarray measurements were performed by the University of Michigan DNA Sequencing Core as previously described (17). Four samples from each group were used for analysis. Biotinylated cDNA was prepared using a GeneChip Plus WT kit (Affymetrix, Santa Clara, CA) from 400 ng of total RNA. Following fragmentation, cDNA was hybridized on Rat Gene 2.1 ST array plates (Affymetrix) and read using the GeneTitan system (Affymetrix). ArrayStar version 13 (DNASTAR, Madison, WI) was used calculate fold changes in gene expression data. The microarray data set is available through the National Institutes of Health GEO database (accession number GSE62388). Ingenuity Pathway Analysis (IPA, Qiagen) was used to perform gene enrichment analysis.
For quantitative PCR (qPCR), RNA was reverse transcribed into cDNA using the iScipt system (BioRad, Hercules, CA). cDNA was amplified in a CFX96 real-time thermal cycler (BioRad) using iTaq SYBR green reagents (BioRad). Target gene expression was normalized to the stable housekeeping gene eukaryotic translation initiation factor 2B subunit 2 (Eif2b2), and further normalized to the relative expression values from plantaris tendons that were not subjected to synergist ablation using the 2−ΔΔCt technique. Eif2b2 was selected as a housekeeping gene from microarray data and validated with qPCR. Using this approach, any expression value greater than 1 indicates an upregulation compared with nonoverloaded controls, and any value below 1 indicates a downregulation compared with controls. Primer sequence information is provided in Supplementary Table 1.
Pyridinoline and hydroxyproline assays.
Muscle samples (50 mg wet weight) were dried overnight at 100°C, weighed immediately, and then hydrolyzed in 500 μl of 12 M HCl for 6 h at 110°C. Samples were then evaporated in a SpeedVac (Thermo Scientific) and resuspended in water. Pyridinoline levels were measured using an ELISA (Quidel, San Diego, CA) following the recommendations of the manufacturer. Hydroxyproline levels were determined using a colorimetric assay as previously described (6, 27). Samples were assayed in duplicate and absorbance values were measured using a SpectraMax M5 microplate reader (Molecular Devices, Sunnyvale, CA). Pyridinoline and hydroxyproline levels were normalized to dry muscle mass.
Statistical analysis.
Data are presented as means ± SD. Differences between groups were tested using a one-way ANOVA (α = 0.05) followed by Fisher's least significant differences post hoc sorting using Prism 7.0 software (GraphPad, San Diego, CA). Microarray data were tested using a one-way ANOVA, and a Benjamini Hochberg false discovery rate correction was used to adjust for multiple observations.
RESULTS
Compared with control rats, synergist ablation led to a 14–30% increase in muscle mass (Fig. 1A), which was accompanied by a 37 to 63% increase in muscle fiber CSA (Fig. 1B). Fibers appeared swollen at 3 days, and by 7 days, gross signs of muscle regeneration, including centrally located nuclei, were present (Fig. 1C). However, at 28 days, the muscle appeared normal with hypertrophied, less compact fibers compared with the control group (Fig. 1C). The contractile function of permeabilized muscle fibers was then measured (Table 1). Fiber CSA measurements differed somewhat compared with values obtained through histological analysis, which likely occurred due to fiber welling. Muscle CSA was reduced by 8% at the 7-day time point and increased by 13% at the 28-day time point compared with that of controls. Maximum isometric force, Fo, was 20–26% lower than controls at the 7- and 28-day time points. Specific force, sFo, was reduced by 21–25% 3 and 7 days after overload, but returned to control levels by 28 days.
Fig. 1.
Muscle morphology and histology. Muscle mass (A), muscle fiber cross-sectional area (CSA) (B), and histology identifying important features (C) from control (Ctrl) muscles and muscles taken 3, 7, or 28 days after synergist ablation. Values are means ± SD, n = 4 muscles per group in Ctrl and n = 8 muscles per group for the 3-, 7-, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher's least significant difference post hoc sorting. #Significantly different (P < 0.05) from Ctrl muscles, †significantly different (P < 0.05) from 3-day synergist ablation muscles, ‡significantly different (P < 0.05) from 7-day synergist ablation muscles.
Table 1.
Permeabilized fiber contractility
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| Fiber CSA, µm2 | 8,170 ± 1,532 | 9,222 ± 1,287 | 7,499 ± 1,141† | 9,235 ± 685.4‡ |
| Fo, mN | 0.765 ± 0.118 | 0.671 ± 0.158 | 0.565 ± 0.084* | 0.917 ± 0.049*†‡ |
| sFo, kPa | 101 ± 8.62 | 75.6 ± 13.9* | 79.9 ± 5.99* | 95.7 ± 10.1†‡ |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles,
significantly different (P < 0.05) from 7-day synergist ablation muscles.
CSA, cross-section area; Fo, maximum force production; sFo, specific force.
We next measured changes in ECM composition (Table 2). Pyridinoline was elevated by 27% 7 days after synergist ablation, and by 40% 28 days after overload, whereas hydroxyproline levels were reduced by 29–39% after overload. When pyridinoline was normalized to hydroxyproline levels, this ratio was elevated by 91–113% in overloaded muscles compared with controls.
Table 2.
Muscle extracellular matrix markers
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| Pyridinoline, ng/mg | 35.1 ± 7.5 | 41.0 ± 4.6 | 44.6 ± 8.0* | 49.2 ± 8.5*† |
| Hydroxyproline, µg/mg | 3.2 ± 1.2 | 2.0 ± 0.4* | 2.3 ± 0.4* | 2.6 ± 0.6† |
| Pyridinoline/Hydroxyproline, ng/µg | 10.2 ± 2.7 | 21.6 ± 4.5* | 19.8 ± 2.9* | 19.4 ± 4.4* |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles.
Following measurement of ECM markers, we analyzed levels of signaling proteins involved in muscle and ECM growth and remodeling (Table 3). For proteins involved in muscle hypertrophy, p-Akt and p-mTOR levels were not elevated until 7 days after synergist ablation, whereas p-p70S6K was elevated as soon as 3 days. p-Smad2 and p-Smad3, which are involved in muscle atrophy and ECM production, were also elevated 3 days after overload, and declined thereafter. Smad4 levels did not change throughout the study. p-ERK1/2, p-p38 MAPK, and p-STAT3 demonstrated generally similar responses as observed for p-Smad2 and p-Smad3.
Table 3.
Signaling protein abundance
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| p-Akt, RU | 1.00 ± 0.17 | 0.61 ± 0.21* | 1.71 ± 0.10*† | 0.79 ± 0.24‡ |
| p-mTOR, RU | 1.00 ± 0.11 | 1.12 ± 0.35 | 1.57 ± 0.11*† | 1.07 ± 0.17‡ |
| p-p70S6K, RU | 1.00 ± 0.35 | 1.90 ± 0.38* | 1.70 ± 0.39* | 0.91 ± 0.26†‡ |
| p-Smad2, RU | 1.00 ± 0.16 | 1.97 ± 0.18* | 1.40 ± 0.20*† | 1.26 ± 0.16*† |
| p-Smad3, RU | 1.00 ± 0.10 | 1.71 ± 0.12* | 1.72 ± 0.15* | 1.10 ± 0.19†‡ |
| Smad4, RU | 1.00 ± 0.12 | 1.01 ± 0.17 | 1.04 ± 0.07 | 1.07 ± 0.09 |
| p-ERK1/2 | 1.00 ± 0.10 | 1.58 ± 0.23* | 1.19 ± 0.35† | 1.04 ± 0.23† |
| p-p38 MAPK, RU | 1.00 ± 0.06 | 1.79 ± 0.24* | 0.79 ± 0.19† | 0.75 ± 0.15*† |
| p-STAT3, RU | 1.00 ± 0.07 | 1.71 ± 0.20* | 1.82 ± 0.30* | 1.19 ± 0.38†‡ |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles,
significantly different (P < 0.05) from 7-day synergist ablation muscles.
Finally, we measured changes in the transcriptome that occurred after synergist ablation. In overloaded muscles, a total of 8,343 genes, or ~23% of the measured transcriptome, were either 1.5-fold upregulated or downregulated and were also significantly different from controls. We then used qPCR to measure expression of individual genes that have a known role in muscle and ECM growth and remodeling, or had substantial changes in fold expression between groups in the microarray data set.
For growth factors and cytokines (Table 4), no difference in activin A expression was observed. Activin B, BMP7, and GDF11 demonstrated a progressive increase in expression throughout the overload period. Expressions of IGF1-Eb, TGF-β, and TWEAK peaked at 3 days, whereas CTGF, IGF1-Ea, and irisin peaked at 7 days. Myostatin was downregulated throughout the study.
Table 4.
Expression of growth factors and cytokines measured by quantitative PCR
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| Activin A | 1.00 ± 0.50 | 0.79 ± 0.77 | 1.13 ± 1.08 | 1.91 ± 1.43 |
| Activin B | 1.00 ± 0.72 | 5.53 ± 1.03* | 6.79 ± 1.63* | 38.7 ± 2.07*†‡ |
| BMP7 | 1.00 ± 0.32 | 4.31 ± 2.74 | 7.81 ± 3.87* | 25.2 ± 4.80*†‡ |
| CTGF | 1.00 ± 0.51 | 1.14 ± 0.50 | 2.03 ± 1.19*† | 0.63 ± 0.53‡ |
| GDF11 | 1.00 ± 0.05 | 1.12 ± 0.15 | 1.65 ± 0.21*† | 3.64 ± 0.30*†‡ |
| IGF1Ea | 1.00 ± 0.24 | 2.10 ± 0.58* | 4.84 ± 0.96*† | 2.35 ± 0.63*‡ |
| IGF1Eb | 1.00 ± 0.43 | 7.67 ± 1.49* | 3.53 ± 0.96*† | 2.19 ± 1.15†‡ |
| Irisin | 1.00 ± 0.15 | 1.90 ± 0.74 | 2.59 ± 0.87* | 1.40 ± 0.81‡ |
| Myostatin | 1.00 ± 0.11 | 0.08 ± 0.06* | 0.17 ± 0.10* | 0.21 ± 0.06* |
| TGF-β | 1.00 ± 0.88 | 22.6 ± 3.82* | 12.8 ± 1.92*† | 4.15 ± 0.33*†‡ |
| TWEAK | 1.00 ± 0.28 | 5.80 ± 2.73* | 0.53 ± 0.20† | 1.22 ± 0.87† |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles,
significantly different (P < 0.05) from 7-day synergist ablation muscles.
In genes involved with muscle atrophy and protein degradation (Table 5), atrogin-1, MuRF1, MuSA1, beclin-1, µ-calpain, m-calpain, and calpain-5 were elevated 3 days and 7 days after overload, but returned to control levels by the 28-day time point. Calpain-6 expression peaked at 7 days before returning to baseline levels at 28 days. The myogenic regulatory factors (Table 5) MyoD, myogenin, and MRF4 had highest expression at 7 days, as did myomaker and members of the Hippo pathway, MST1 and YAP. Expression of the contractile genes (Table 5) embryonic myosin heavy chain, perinatal myosin heavy chain, and type I myosin heavy chain also peaked at 7 days, whereas types IIA, IIB, and IIX myosin heavy chains were elevated at the 28-day time point.
Table 5.
Atrophy, myogenesis, and sarcomerogenesis genes measured by quantitative PCR
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| Atrogin1 | 1.00 ± 0.12 | 4.72 ± 0.36* | 9.21 ± 2.54*† | 0.61 ± 0.37†‡ |
| Beclin1 | 1.00 ± 0.07 | 2.78 ± 0.87* | 5.75 ± 1.84*† | 0.41 ± 0.27†‡ |
| Calpain5 | 1.00 ± 0.23 | 7.16 ± 3.05* | 10.2 ± 4.38* | 2.87 ± 0.64†‡ |
| Calpain6 | 1.00 ± 0.25 | 2.38 ± 1.31 | 9.72 ± 1.55*† | 1.31 ± 0.21‡ |
| eMyHC | 1.00 ± 0.31 | 0.27 ± 0.24* | 15.6 ± 2.00*† | 0.57 ± 0.39‡ |
| m-Calpain | 1.00 ± 0.08 | 6.86 ± 2.49* | 7.07 ± 2.67* | 0.55 ± 0.49†‡ |
| MRF4 | 1.00 ± 0.25 | 0.91 ± 0.76 | 1.71 ± 0.69† | 0.69 ± 0.50‡ |
| MST1 | 1.00 ± 0.50 | 6.58 ± 1.39* | 14.2 ± 2.35*† | 2.99 ± 0.63*†‡ |
| MURF1 | 1.00 ± 0.08 | 6.83 ± 3.93* | 5.65 ± 2.22* | 0.57 ± 0.35†‡ |
| MUSA1 | 1.00 ± 0.13 | 3.20 ± 0.98* | 3.93 ± 1.65* | 0.68 ± 0.59†‡ |
| MyHC1 | 1.00 ± 0.31 | 0.11 ± 0.08 | 4.68 ± 2.11*† | 1.18 ± 1.09‡ |
| MyHC2A | 1.00 ± 0.15 | 0.30 ± 0.10 | 3.23 ± 2.76*† | 4.53 ± 1.52*† |
| MyHC2B | 1.00 ± 0.33 | 0.66 ± 0.42 | 2.63 ± 1.63*† | 2.91 ± 0.87*† |
| MyHC2X | 1.00 ± 0.18 | 0.12 ± 0.10 | 0.34 ± 0.31 | 11.5 ± 2.97*†‡ |
| MyoD | 1.00 ± 0.16 | 0.48 ± 0.25* | 1.89 ± 0.21*† | 1.10 ± 0.51†‡ |
| Myogenin | 1.00 ± 0.37 | 2.37 ± 0.59 | 14.5 ± 1.37*† | 2.25 ± 1.93‡ |
| Myomaker | 1.00 ± 0.93 | 3.54 ± 0.46 | 68.7 ± 4.27*† | 3.96 ± 0.82‡ |
| pMyHC | 1.00 ± 0.31 | 1.18 ± 0.95 | 198 ± 15.8*† | 8.19 ± 2.55*†‡ |
| Yap | 1.00 ± 0.17 | 0.41 ± 0.22 | 5.80 ± 1.94*† | 0.92 ± 0.82‡ |
| µ-Calpain | 1.00 ± 0.13 | 5.67 ± 2.10* | 7.56 ± 1.09*† | 1.64 ± 0.87†‡ |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles,
significantly different (P < 0.05) from 7-day synergist ablation muscles.
For genes that encode ECM structural and remodeling proteins (Table 6), expression of the fibrillar type I and III collagens were elevated by 7 days after overload, and continued to be elevated at 28 days. The basement membrane collagens IV, V, VI, and VIII, along with the fibril associated collagens XII and XIV, were elevated as soon as 3 days after overload. Most of the basement membrane collagens returned to baseline levels by 28 days, whereas collagen XIV remained elevated at 28 days. The proteoglycans asporin, biglycan, lumican, and versican were upregulated 3 days after overload, and asporin and versican remained elevated at 28 days. The hyaluronic acid matrix-producing enzymes HAS1 and HAS2 were also induced 3 days and remained so throughout the study, whereas Hyal1, which breaks down hyaluronic acid, was downregulated at 3 days and induced thereafter. Elastin and the cross-linking enzyme lysyl oxidase were both elevated at 3 days, and returned to baseline by 28 days. The stromelysin MMP3, the gelatinase MMP9, the collagenase MMP13, and the membrane-tethered MMP14 were elevated by 3 days after synergist ablation, whereas the gelatinase MMP2 was elevated at 7 days, and the collagenase MMP8 by 28 days. TIMP1 and TIMP2 were also elevated 3 days after overload.
Table 6.
Extracellular matrix, MMP, and TIMP genes measured by quantitative PCR
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| Asporin | 1.00 ± 0.31 | 5.68 ± 1.47* | 7.28 ± 0.74* | 4.96 ± 0.73* |
| Biglycan | 1.00 ± 0.23 | 7.51 ± 2.37* | 12.6 ± 1.45* | 2.43 ± 1.06*† |
| Col12a1 | 1.00 ± 0.12 | 4.53 ± 4.83* | 5.62 ± 3.89* | 1.63 ± 1.04†‡ |
| Col14a1 | 1.00 ± 0.14 | 4.32 ± 1.00* | 5.83 ± 2.06* | 4.31 ± 2.44* |
| Col1a1 | 1.00 ± 0.12 | 5.65 ± 4.33* | 32.2 ± 3.23*† | 6.89 ± 3.44*‡ |
| Col3a1 | 1.00 ± 0.26 | 3.43 ± 2.13 | 15.2 ± 1.10*† | 17.3 ± 2.46*† |
| Col4a1 | 1.00 ± 0.07 | 6.93 ± 2.69* | 5.18 ± 3.88* | 2.32 ± 1.86†‡ |
| Col5a1 | 1.00 ± 0.05 | 13.4 ± 10.0* | 14.9 ± 6.66* | 3.95 ± 1.58†‡ |
| Col6a1 | 1.00 ± 0.16 | 12.8 ± 3.59* | 9.45 ± 2.52*† | 1.43 ± 0.30†‡ |
| Col8a1 | 1.00 ± 0.34 | 5.96 ± 1.21* | 5.48 ± 1.29* | 1.80 ± 1.28†‡ |
| Elastin | 1.00 ± 0.28 | 8.22 ± 1.43* | 2.61 ± 2.14*† | 1.27 ± 0.56†‡ |
| HAS1 | 1.00 ± 0.24 | 63.2 ± 9.20* | 15.8 ± 7.19*† | 16.6 ± 3.29*† |
| HAS2 | 1.00 ± 0.49 | 8.22 ± 1.07* | 9.04 ± 2.67* | 4.37 ± 1.52* |
| Hyal1 | 1.00 ± 0.16 | 0.33 ± 0.35* | 2.60 ± 0.27*† | 1.72 ± 0.54*†‡ |
| LOX | 1.00 ± 0.42 | 20.9 ± 1.18* | 10.6 ± 1.16*† | 1.74 ± 1.39†‡ |
| Lumican | 1.00 ± 0.10 | 9.02 ± 3.87* | 6.12 ± 4.21* | 1.69 ± 0.43†‡ |
| MMP13 | 1.00 ± 0.22 | 82.4 ± 39.9* | 60.7 ± 29.3* | 8.06 ± 7.28†‡ |
| MMP14 | 1.00 ± 0.95 | 28.5 ± 4.29* | 36.5 ± 8.14*† | 7.53 ± 0.32*†‡ |
| MMP2 | 1.00 ± 0.16 | 2.11 ± 0.45 | 6.19 ± 2.50*† | 2.29 ± 0.59‡ |
| MMP3 | 1.00 ± 0.82 | 55.5 ± 7.88* | 29.6 ± 16.4*† | 15.0 ± 3.53*†‡ |
| MMP8 | 1.00 ± 0.62 | 1.59 ± 1.29 | 1.54 ± 1.16 | 5.03 ± 1.81*†‡ |
| MMP9 | 1.00 ± 0.58 | 98.9 ± 11.6* | 109 ± 27.6* | 9.11 ± 2.92†‡ |
| TIMP1 | 1.00 ± 0.31 | 77.6 ± 12.1* | 10.9 ± 1.29*† | 5.00 ± 1.40† |
| TIMP2 | 1.00 ± 0.33 | 3.05 ± 1.33* | 2.39 ± 0.70 | 1.99 ± 1.53 |
| Versican | 1.00 ± 0.96 | 45.2 ± 8.34* | 14.3 ± 1.43*† | 7.60 ± 0.27*†‡ |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles,
significantly different (P < 0.05) from 7-day synergist ablation muscles.
There was a pronounced induction of several genes involved with the regulation of fibroblast and immune cell activity (Table 7), including FAP, FSP1, osteoactivin, and osteopontin, which generally tapered quickly thereafter. PDGFRa was elevated 3 and 28 days after overload. The neutrophil marker Ly6C, and the panmacrophage marker F4/80, were upregulated 3 days after overload. The M1 macrophage markers CD11b and CD68 were elevated 3 days after overload, and CD11b returned to baseline expression by 28 days. For M2 macrophage markers, CD163 was induced by 3 days, whereas CD168 was not elevated until 7 days, and both remained elevated through 28 days after synergist ablation.
Table 7.
Fibroblast and immune cell-related genes measured by quantitative PCR
| Control | 3 Days | 7 Days | 28 Days | |
|---|---|---|---|---|
| CD11b | 1.00 ± 0.48 | 50.2 ± 9.24* | 23.2 ± 7.21*† | 3.31 ± 1.15†‡ |
| CD163 | 1.00 ± 0.31 | 18.5 ± 2.05* | 3.96 ± 1.72† | 7.54 ± 4.37*†‡ |
| CD168 | 1.00 ± 0.17 | 2.09 ± 1.49 | 11.8 ± 2.54*† | 4.04 ± 0.99*†‡ |
| CD68 | 1.00 ± 0.28 | 46.2 ± 7.16* | 8.34 ± 0.86*† | 7.23 ± 3.24*† |
| F4/80 | 1.00 ± 0.20 | 14.9 ± 1.53* | 7.66 ± 1.66*† | 6.38 ± 5.95*† |
| FAP | 1.00 ± 0.18 | 23.0 ± 2.02* | 4.16 ± 2.14*† | 2.72 ± 2.53*† |
| FSP1 | 1.00 ± 0.48 | 31.4 ± 0.98* | 21.9 ± 2.04*† | 2.01 ± 0.98†‡ |
| Ly6c | 1.00 ± 0.10 | 9.03 ± 3.87* | 6.12 ± 4.21* | 1.69 ± 0.43†‡ |
| Osteoactivin | 1.00 ± 0.40 | 58.3 ± 12.2* | 8.66 ± 1.07† | 2.18 ± 1.12† |
| Osteopontin | 1.00 ± 0.09 | 331 ± 51.1* | 6.38 ± 3.96† | 1.61 ± 0.91† |
| PDGFRa | 1.00 ± 0.18 | 4.18 ± 2.41* | 2.90 ± 1.56 | 4.92 ± 2.02*‡ |
Values are means ± SD, n = 4 muscles per group in control and n = 8 muscles per group for the 3-day, 7-day, and 28-day synergist ablation groups. Differences were tested using a one-way ANOVA (α = 0.05) followed by Fisher’s least significant difference post hoc sorting.
Significantly different (P < 0.05) from control muscles,
significantly different (P < 0.05) from 3-day synergist ablation muscles,
significantly different (P < 0.05) from 7-day synergist ablation muscles.
DISCUSSION
Skeletal muscle hypertrophy involves an organized response of both muscle fibers and the ECM to adapt to increased mechanical loads placed upon the tissue. In the current study, we used a rat synergist ablation model to study early and later changes in fiber contractility and ECM composition. To support the primary objectives, we also measured activation of various signal transduction pathways, and corresponding changes in the muscle transcriptome at these time points. We found that muscle fiber sFo, which is a useful biomarker of muscle function, was reduced 3 and 7 days after onset of mechanical overload, but had returned to control levels by 28 days. Levels of collagen were also acutely reduced at the 3- and 7-day time points, but returned to normal levels 28 days after synergist ablation. Nearly a quarter of the transcripts measured using microarrays were differentially regulated throughout the study. The differences in activation of various signaling proteins and changes in numerous transcripts involved in muscle and ECM growth and remodeling generally corresponded well to the observed structural and functional changes in muscle tissue. Overall, the findings of the study support our initial hypotheses that fibers from the plantaris muscle would experience a reduction in sFo 3 and 7 days after synergist ablation, which would be restored to control levels by 28 days after overload, and that there would be a slow, progressive accumulation in collagen content and cross links over the course of hypertrophy.
The permeabilized fiber measurement technique is useful for the study of muscle function at the single cell level (34). We and numerous other groups have used this approach to study long-term changes in muscle fiber function in response to injury or resistance exercise. Chronic injury or immobilization can result in an up to 40% reduction in Fo and up to 30% reduction in sFo in rodent models, and declines of up to 30% in Fo and sFo in human muscles (10, 12, 28, 43, 47). For studies of exercise training in humans, Fo can increase by up to 55%, although the CSA of fibers increases proportionally such that sFo typically does not change or it increases only slightly (4, 42, 45).
In the current study, following synergist ablation, there was a rapid decline in sFo that occurred at the 3- and 7-day time points, along with a reduction in Fo at 7 days. We sought to not only assess functional alterations in muscle contractility, but also to correlate these with biochemical and molecular factors that regulate muscle fiber hypertrophy. Bioinformatics analysis through IPA suggested that, for genes associated with myogenesis and muscle development, most changes occur at the 3- and 7-day time points, with only a small number of genes differentially regulated at 28 days after overload. The changes in contractility at the 3- and 7-day time points correspond to marked increases in expression of signaling pathways that induce muscle atrophy and protein degradation, such as activin B, TGF-β, and TWEAK (14). Myostatin plays an important role in inducing muscle atrophy after injury, and was downregulated throughout the study, which is consistent with previous findings (7). MyoD and myogenin, which are transcriptional regulators of satellite cell proliferation and differentiation (16), and myomaker, which allows the fusion of myoblasts into myotubes (29), were upregulated to the greatest extent at 7 days. This corresponded to the appearance of small myotubes in histological sections that were observed at this same time point. The R-Smads, Smad2 and Smad3, along with ERK1/2, p38 MAPK, and STAT3, which together are downstream effectors of the activin and TGF-β receptors (35), demonstrated increased activation at the 3- and 7-day time points. The calpains, which are critical in the disassembly of mechanically damaged sarcomeres (19), and the E3 ubiquitin ligases atrogin-1, MuRF-1, and MuSA-1, which target damaged proteins to the 26S proteasome for proteolytic breakdown (35), were upregulated at the 3- and 7-day time points. MuRF-1 also plays an important role in regulating M-line structure and assembly in cardiomyocytes, and an induction of MuRF-1 may be independent of protein turnover (26). Beclin-1, which is an important regulator of autophagy, was also upregulated to a similar extent (8).
By 28 days, overloaded muscles demonstrated muscle fiber hypertrophy, an increase in Fo and a return of sFo to levels similar to those of control muscles. BMP7, which is a recently described potent activator of muscle hypertrophy (36), was markedly elevated at the 28-day time point. However, other signaling molecules that induce hypertrophy such as IGF1-Ea and IGF1-Eb (14), although elevated at 28 days, were more pronounced at 3 and 7 days. Their downstream effectors in muscle hypertrophy, Akt, mTOR, and p70S6K (14), were also elevated at 3 and 7 days, but returned to control levels by 28 days. Early markers of sarcomerogenesis, eMyHC and pMyHC, were elevated at the 7-day time point, whereas the mature myosin heavy chains MyHC2A and MyHC2X were most significantly upregulated at the 28-day time point. The combined results from the muscle fiber contractility experiments indicate that in the plantaris overload model there is acute damage to muscle fibers that reduces muscle Fo and sFo within 1 wk, and that by 1 mo the muscle has hypertrophied and generates more total force production, whereas normalized force production is similar to that of control muscle. Although definitive connections cannot be made, the alterations in muscle contractile properties generally correspond to changes in the activation of various signaling pathways and genes that regulate muscle growth and remodeling.
Muscle hypertrophy is often accompanied by increased collagen deposition (14). Following synergist ablation, hydroxyproline, which is an amino acid that serves as a biomarker for the major muscle collagens, type I and III, was decreased at the 3- and 7-day time points, but returned to control levels at 28 days. The reduction in collagen content may be due to an increase in expression of the collagenase MMP13, which was highly induced at these time points. By 7 days, type I and III collagens were induced and remained elevated at 28 days, along with the expression of type XII and XIV collagens, which assist in the formation of mature fibrillar collagen fibrils. This is consistent with a previous study that reported an increase in type I collagen expression in mice 21 days after synergist ablation (44). The basement membrane collagens, which directly surround muscle fibers and assist in the transfer of lateral and longitudinal transmission of force in muscle (14), were upregulated at the two early time points, and returned to control expression by 28 days. Changes in the basement membrane collagens were accompanied by generally similar changes in the gelatinases MMP2 and MMP9, which degrade this class of collagens. Resistance exercise also frequently increases ECM stiffness, which comes about in part due to increases in cross-links between adjacent collagen molecules (14). Pyridinoline is formed via the covalent bonding of three hydroxylysine residues between collagen molecules through a process catalyzed by the enzyme lysyl oxidase (24). In the current study, although absolute levels of pyridinoline did not increase until 7 and 28 days after overload, when normalized to hydroxyproline to reflect overall collagen abundance, relative pyridinoline levels were elevated throughout the overload period. Lysyl oxidase was also markedly upregulated at the 3-day time point, and gradually decreased over time.
There were marked changes in the minor components of muscle ECM, similar to those of the major ECM molecules. Hyaluronic acid can form an immature matrix that serves as a template for mature, collagen-rich matrix formation (2), and the two major muscle hyaluronic acid synthesis enzymes, HAS1 and HAS2, were markedly induced 3 days after overload. The proteoglycans asporin, biglycan, lumican, and versican, which play important roles in organizing the basement membrane and regulating heat dissipation during muscle contraction (46), were upregulated at the 3-day time point, and generally reduced over time, similar to that of the basement membrane collagens. Elastin, which stores elastic energy in the ECM, was also rapidly induced 3 days after overload and gradually declined by 28 days.
We also measured expression of genes that regulate activity of fibroblasts, which are the major cell type in muscle that remodel the ECM. FAP and FSP1 serve as markers of muscle fibroblasts (11, 33), and PDGFRa, which is a marker of fibroblast progenitor cells (20), were upregulated by 3 days, and expression of FAP and FSP1 was downregulated over time. MMP14 is a membrane-tethered matrix metalloproteinase that is expressed on fibroblasts to allow them to migrate through the ECM (5), and the expression pattern of MMP14 overlapped well with the patterns of FAP and FSP1. Osteopontin functions to recruit immune cells to injured muscle after eccentric exercise, and stood out as one of the most differentially regulated transcripts over time in microarray data (1). Levels of osteopontin were upregulated by more than 300-fold at the 3-day time point, but returned to control levels thereafter. For markers of immune cells, which help to work along with fibroblasts in regeneration and remodeling of muscle tissue, the neutrophil marker Ly6C (39) was elevated at 3 and 7 days. The panmacrophage marker F4/80 was elevated consistently at all time points, whereas the M1 macrophage markers CD11b and CD68 (39) peaked at 3 days, and the M2 marker CD168 (39) was highest at 7 days. The combined results from the ECM, fibroblast, and immune cell data suggest that there could be an acute breakdown of the mature fibrillar collagenous matrix 3 days after overload, along with the likely formation of an early immature hyaluronic matrix that serves as a template for new collagen synthesis at 7 days. Markers of fibroblast abundance suggest a likely rapid expansion of the fibroblast population. Unlike muscle fibers themselves, the ECM seems to still be undergoing active remodeling at 28 days, which is supported by the expression of individual genes and bioinformatics analysis. Based on studies of muscle injury (40) we expected markers of macrophages to be reduced by 28 days, especially the M1 macrophage population, but the persistence of macrophage markers might be due to continued ECM remodeling, because muscle fiber regeneration appears to be complete by this time point.
Although we provided important insight to the changes in muscle fiber and ECM adaptation during muscle hypertrophy, there are several limitations to this study. The synergist ablation technique places a constant increased load on the plantaris muscle to induce hypertrophy, and the mechanical load experienced by the plantaris is greater than what would typically be expected in at the onset of a human resistance training exercise regime. We did not measure the levels of specific cell types using immunohistochemistry, and relied on gene expression analysis to measure markers of these cells throughout the muscle. The permeabilized fiber technique allows us to measure active tension developed by myofibrils within a cell, but we did not measure calcium handling or ECM mechanics in this model. We were not able to distinguish between changes in signaling proteins between muscle and other cells in the tissue, although because of the relative size of muscle fibers to other cell types, we presume that the observed differences in protein phosphorylation largely occur in proteins contained in muscle fibers. Although we measured expression of numerous genes, it is possible that observed changes in transcripts did not reflect changes at the protein level. Finally, although we analyzed differences in muscle structure and function through 28 days after overload, it is possible that additional remodeling and adaption occur beyond this time point.
Previous studies have investigated long-term changes in muscle fiber function after undergoing several weeks of resistance training (4, 42, 45). The findings from these studies have provided important insight into the long-term changes in muscle fiber function and biochemistry in response to exercise. However, many subjects experience an acute drop in whole muscle force production when beginning a resistance-training program (31), and less is known about acute changes in muscle fiber contractility at the onset of muscle hypertrophy. The plantaris overload model has been used to study skeletal muscle hypertrophy in numerous previous papers. To our knowledge, this work was the first to explore the concurrent changes in muscle fiber contractility and ECM accumulation during hypertrophy, along with changes in signaling protein activation and gene expression in both acute and long-term time points. We think the findings from the study provide important insight into basic biological mechanisms of muscle and ECM growth, and indicate that although muscle fibers appear to mostly have completed remodeling and regeneration by 28 days after synergist ablation, the ECM continues to be actively remodeling at this time point.
GRANTS
Support for this study was provided by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grants R01-AR-063649, F31-AR-065931, and F32-AR-067086.
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the authors.
AUTHOR CONTRIBUTIONS
C.L.M., A.J.S., J.P.G., and K.B.S. conceived and designed research; C.L.M., A.J.S., J.A.G., and K.B.S. performed experiments; C.L.M., A.J.S., J.A.G., J.P.G., and K.B.S. analyzed data; C.L.M., A.J.S., J.A.G., J.P.G., and K.B.S. interpreted results of experiments; C.L.M. and A.J.S. prepared figures; C.L.M. drafted manuscript; C.L.M., A.J.S., J.A.G., J.P.G., and K.B.S. edited and revised manuscript; C.L.M., A.J.S., J.A.G., J.P.G., and K.B.S. approved final version of manuscript.
Supplementary Material
ACKNOWLEDGMENTS
We thank Stuart M. Roche and Danielle Rittman for technical assistance.
REFERENCES
- 1.Barfield WL, Uaesoontrachoon K, Wu C-S, Lin S, Chen Y, Wang PC, Kanaan Y, Bond V, Hoffman EP. Eccentric muscle challenge shows osteopontin polymorphism modulation of muscle damage. Hum Mol Genet 23: 4043–4050, 2014. doi: 10.1093/hmg/ddu118. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2.Calve S, Isaac J, Gumucio JP, Mendias CL. Hyaluronic acid, HAS1, and HAS2 are significantly upregulated during muscle hypertrophy. Am J Physiol Cell Physiol 303: C577–C588, 2012. doi: 10.1152/ajpcell.00057.2012. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 3.Chaillou T, Jackson JR, England JH, Kirby TJ, Richards-White J, Esser KA, Dupont-Versteegden EE, McCarthy JJ. Identification of a conserved set of upregulated genes in mouse skeletal muscle hypertrophy and regrowth. J Appl Physiol 118: 86–97, 2015. doi: 10.1152/japplphysiol.00351.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4.Claflin DR, Larkin LM, Cederna PS, Horowitz JF, Alexander NB, Cole NM, Galecki AT, Chen S, Nyquist LV, Carlson BM, Faulkner JA, Ashton-Miller JA. Effects of high- and low-velocity resistance training on the contractile properties of skeletal muscle fibers from young and older humans. J Appl Physiol 111: 1021–1030, 2011. doi: 10.1152/japplphysiol.01119.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 5.Davis ME, Gumucio JP, Sugg KB, Bedi A, Mendias CL. MMP inhibition as a potential method to augment the healing of skeletal muscle and tendon extracellular matrix. J Appl Physiol 115: 884–891, 2013. doi: 10.1152/japplphysiol.00137.2013. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6.Davis ME, Korn MA, Gumucio JP, Harning JA, Saripalli AL, Bedi A, Mendias CL. Simvastatin reduces fibrosis and protects against muscle weakness after massive rotator cuff tear. J Shoulder Elbow Surg 24: 280–287, 2015. doi: 10.1016/j.jse.2014.06.048. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7.Fortes MA, Pinheiro CH, Guimarães-Ferreira L, Vitzel KF, Vasconcelos DA, Curi R. Overload-induced skeletal muscle hypertrophy is not impaired in STZ-diabetic rats. Physiol Rep 3: e12457, 2015. doi: 10.14814/phy2.12457. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8.Funderburk SF, Wang QJ, Yue Z. The Beclin 1-VPS34 complex—at the crossroads of autophagy and beyond. Trends Cell Biol 20: 355–362, 2010. doi: 10.1016/j.tcb.2010.03.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9.Goodman CA, Dietz JM, Jacobs BL, McNally RM, You J-S, Hornberger TA. Yes-Associated Protein is up-regulated by mechanical overload and is sufficient to induce skeletal muscle hypertrophy. FEBS Lett 589: 1491–1497, 2015. doi: 10.1016/j.febslet.2015.04.047. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 10.Gumucio JP, Davis ME, Bradley JR, Stafford PL, Schiffman CJ, Lynch EB, Claflin DR, Bedi A, Mendias CL. Rotator cuff tear reduces muscle fiber specific force production and induces macrophage accumulation and autophagy. J Orthop Res 30: 1963–1970, 2012. doi: 10.1002/jor.22168. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 11.Gumucio JP, Flood MD, Phan AC, Brooks SV, Mendias CL. Targeted inhibition of TGF-β results in an initial improvement but long-term deficit in force production after contraction-induced skeletal muscle injury. J Appl Physiol 115: 539–545, 2013. doi: 10.1152/japplphysiol.00374.2013. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 12.Gumucio JP, Korn MA, Saripalli AL, Flood MD, Phan AC, Roche SM, Lynch EB, Claflin DR, Bedi A, Mendias CL. Aging-associated exacerbation in fatty degeneration and infiltration after rotator cuff tear. J Shoulder Elbow Surg 23: 99–108, 2014. doi: 10.1016/j.jse.2013.04.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 13.Gumucio JP, Phan AC, Ruehlmann DG, Noah AC, Mendias CL. Synergist ablation induces rapid tendon growth through the synthesis of a neotendon matrix. J Appl Physiol 117: 1287–1291, 2014. doi: 10.1152/japplphysiol.00720.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14.Gumucio JP, Sugg KB, Mendias CL. TGF-β superfamily signaling in muscle and tendon adaptation to resistance exercise. Exerc Sport Sci Rev 43: 93–99, 2015. doi: 10.1249/JES.0000000000000041. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15.Hamilton DL, Philp A, MacKenzie MG, Patton A, Towler MC, Gallagher IJ, Bodine SC, Baar K. Molecular brakes regulating mTORC1 activation in skeletal muscle following synergist ablation. Am J Physiol Endocrinol Metab 307: E365–E373, 2014. doi: 10.1152/ajpendo.00674.2013. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 16.Hawke TJ, Garry DJ. Myogenic satellite cells: physiology to molecular biology. J Appl Physiol 91: 534–551, 2001. [DOI] [PubMed] [Google Scholar]
- 17.Hudgens JL, Sugg KB, Grekin JA, Gumucio JP, Bedi A, Mendias CL. Platelet-rich plasma activates proinflammatory signaling pathways and induces oxidative stress in tendon fibroblasts. Am J Sports Med 44: 1931–1940, 2016. doi: 10.1177/0363546516637176. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Huijing PA, Baan GC, Rebel GT. Non-myotendinous force transmission in rat extensor digitorum longus muscle. J Exp Biol 201: 683–691, 1998. [PubMed] [Google Scholar]
- 19.Jackman RW, Kandarian SC. The molecular basis of skeletal muscle atrophy. Am J Physiol Cell Physiol 287: C834–C843, 2004. doi: 10.1152/ajpcell.00579.2003. [DOI] [PubMed] [Google Scholar]
- 20.Joe AW, Yi L, Natarajan A, Le Grand F, So L, Wang J, Rudnicki MA, Rossi FM. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nat Cell Biol 12: 153–163, 2010. doi: 10.1038/ncb2015. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 21.Kandarian SC, White TP. Force deficit during the onset of muscle hypertrophy. J Appl Physiol 67: 2600–2607, 1989. [DOI] [PubMed] [Google Scholar]
- 22.Kjaer M. Role of extracellular matrix in adaptation of tendon and skeletal muscle to mechanical loading. Physiol Rev 84: 649–698, 2004. doi: 10.1152/physrev.00031.2003. [DOI] [PubMed] [Google Scholar]
- 23.Lieber RL, Ward SR. Cellular mechanisms of tissue fibrosis. 4. Structural and functional consequences of skeletal muscle fibrosis. Am J Physiol Cell Physiol 305: C241–C252, 2013. doi: 10.1152/ajpcell.00173.2013. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 24.Light N, Bailey AJ. Collagen cross-links: location of pyridinoline in type I collagen. FEBS Lett 182: 503–508, 1985. doi: 10.1016/0014-5793(85)80363-X. [DOI] [PubMed] [Google Scholar]
- 25.Manders E, Bonta PI, Kloek JJ, Symersky P, Bogaard H-J, Hooijman PE, Jasper JR, Malik FI, Stienen GJM, Vonk-Noordegraaf A, de Man FS, Ottenheijm CA. Reduced force of diaphragm muscle fibers in patients with chronic thromboembolic pulmonary hypertension. Am J Physiol Lung Cell Mol Physiol 311: L20–L28, 2016. doi: 10.1152/ajplung.00113.2016. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 26.McElhinny AS, Kakinuma K, Sorimachi H, Labeit S, Gregorio CC. Muscle-specific RING finger-1 interacts with titin to regulate sarcomeric M-line and thick filament structure and may have nuclear functions via its interaction with glucocorticoid modulatory element binding protein-1. J Cell Biol 157: 125–136, 2002. doi: 10.1083/jcb.200108089. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 27.Mendias CL, Lynch EB, Gumucio JP, Flood MD, Rittman DS, Van Pelt DW, Roche SM, Davis CS. Changes in skeletal muscle and tendon structure and function following genetic inactivation of myostatin in rats. J Physiol 593: 2037–2052, 2015. doi: 10.1113/jphysiol.2014.287144. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 28.Mendias CL, Roche SM, Harning JA, Davis ME, Lynch EB, Sibilsky Enselman ER, Jacobson JA, Claflin DR, Calve S, Bedi A. Reduced muscle fiber force production and disrupted myofibril architecture in patients with chronic rotator cuff tears. J Shoulder Elbow Surg 24: 111–119, 2015. doi: 10.1016/j.jse.2014.06.037. [DOI] [PubMed] [Google Scholar]
- 29.Millay DP, O’Rourke JR, Sutherland LB, Bezprozvannaya S, Shelton JM, Bassel-Duby R, Olson EN. Myomaker is a membrane activator of myoblast fusion and muscle formation. Nature 499: 301–305, 2013. doi: 10.1038/nature12343. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 30.Oak NR, Gumucio JP, Flood MD, Saripalli AL, Davis ME, Harning JA, Lynch EB, Roche SM, Bedi A, Mendias CL. Inhibition of 5-LOX, COX-1, and COX-2 increases tendon healing and reduces muscle fibrosis and lipid accumulation after rotator cuff repair. Am J Sports Med 42: 2860–2868, 2014. doi: 10.1177/0363546514549943. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 31.Philippou A, Maridaki M, Bogdanis GC. Angle-specific impairment of elbow flexors strength after isometric exercise at long muscle length. J Sports Sci 21: 859–865, 2003. doi: 10.1080/0264041031000140356. [DOI] [PubMed] [Google Scholar]
- 32.Ramaswamy KS, Palmer ML, van der Meulen JH, Renoux A, Kostrominova TY, Michele DE, Faulkner JA. Lateral transmission of force is impaired in skeletal muscles of dystrophic mice and very old rats. J Physiol 589: 1195–1208, 2011. doi: 10.1113/jphysiol.2010.201921. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 33.Roberts EW, Deonarine A, Jones JO, Denton AE, Feig C, Lyons SK, Espeli M, Kraman M, McKenna B, Wells RJ, Zhao Q, Caballero OL, Larder R, Coll AP, O’Rahilly S, Brindle KM, Teichmann SA, Tuveson DA, Fearon DT. Depletion of stromal cells expressing fibroblast activation protein-α from skeletal muscle and bone marrow results in cachexia and anemia. J Exp Med 210: 1137–1151, 2013. doi: 10.1084/jem.20122344. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 34.Roche SM, Gumucio JP, Brooks SV, Mendias CL, Claflin DR. Measurement of maximum isometric force generated by permeabilized skeletal muscle fibers. J Vis Exp 100: e52695, 2015. doi: 10.3791/52695. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 35.Sartori R, Gregorevic P, Sandri M. TGFβ and BMP signaling in skeletal muscle: potential significance for muscle-related disease. Trends Endocrinol Metab 25: 464–471, 2014. doi: 10.1016/j.tem.2014.06.002. [DOI] [PubMed] [Google Scholar]
- 36.Sartori R, Schirwis E, Blaauw B, Bortolanza S, Zhao J, Enzo E, Stantzou A, Mouisel E, Toniolo L, Ferry A, Stricker S, Goldberg AL, Dupont S, Piccolo S, Amthor H, Sandri M. BMP signaling controls muscle mass. Nat Genet 45: 1309–1318, 2013. doi: 10.1038/ng.2772. [DOI] [PubMed] [Google Scholar]
- 37.Schwartz AJ, Sarver DC, Sugg KB, Dzierzawski JT, Gumucio JP, Mendias CL. p38 MAPK signaling in postnatal tendon growth and remodeling. PLoS One 10: e0120044, 2015. doi: 10.1371/journal.pone.0120044. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 38.Sharma N, Castorena CM, Cartee GD. Tissue-specific responses of IGF-1/insulin and mTOR signaling in calorie restricted rats. PLoS One 7: e38835, 2012. doi: 10.1371/journal.pone.0038835. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 39.Sugg KB, Lubardic J, Gumucio JP, Mendias CL. Changes in macrophage phenotype and induction of epithelial-to-mesenchymal transition genes following acute Achilles tenotomy and repair. J Orthop Res 32: 944–951, 2014. doi: 10.1002/jor.22624. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 40.Tidball JG, Villalta SA. Regulatory interactions between muscle and the immune system during muscle regeneration. Am J Physiol Regul Integr Comp Physiol 298: R1173–R1187, 2010. doi: 10.1152/ajpregu.00735.2009. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 41.Trappe S, Gallagher P, Harber M, Carrithers J, Fluckey J, Trappe T. Single muscle fibre contractile properties in young and old men and women. J Physiol 552: 47–58, 2003. doi: 10.1113/jphysiol.2003.044966. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 42.Trappe S, Williamson D, Godard M, Porter D, Rowden G, Costill D. Effect of resistance training on single muscle fiber contractile function in older men. J Appl Physiol 89: 143–152, 2000. [DOI] [PubMed] [Google Scholar]
- 43.Trappe S, Trappe T, Gallagher P, Harber M, Alkner B, Tesch P. Human single muscle fibre function with 84 day bed-rest and resistance exercise. J Physiol 557: 501–513, 2004. doi: 10.1113/jphysiol.2004.062166. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 44.White JP, Reecy JM, Washington TA, Sato S, Le ME, Davis JM, Wilson LB, Carson JA. Overload-induced skeletal muscle extracellular matrix remodelling and myofibre growth in mice lacking IL-6. Acta Physiol (Oxf) 197: 321–332, 2009. doi: 10.1111/j.1748-1716.2009.02029.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 45.Widrick JJ, Stelzer JE, Shoepe TC, Garner DP. Functional properties of human muscle fibers after short-term resistance exercise training. Am J Physiol Regul Integr Comp Physiol 283: R408–R416, 2002. doi: 10.1152/ajpregu.00120.2002. [DOI] [PubMed] [Google Scholar]
- 46.Yoon JH, Halper J. Tendon proteoglycans: biochemistry and function. J Musculoskelet Neuronal Interact 5: 22–34, 2005. [PubMed] [Google Scholar]
- 47.Zhong S, Lowe DA, Thompson LV. Effects of hindlimb unweighting and aging on rat semimembranosus muscle and myosin. J Appl Physiol 101: 873–880, 2006. doi: 10.1152/japplphysiol.00526.2005. [DOI] [PubMed] [Google Scholar]
Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.