Myeloid Cell Responses to Contraction-induced Injury Differ in Muscles of Young and Old Mice

Myeloid Cell Responses to Contraction-induced Injury Differ in Muscles of Young and Old Mice Abstract Myeloid cells play a critical role in regulating muscle degeneration and regeneration. Thus, alterations with aging in the myeloid cell response to muscle damage may affect the progression of the injury in old animals. We hypothesized that neutrophil levels remain elevated and that macrophage accumulation is reduced or delayed in injured muscles of old compared with young animals. Muscles of young and old mice were injured with lengthening contractions and analyzed 2 or 5 days later. Regardless of age, neutrophil (Gr-1+) and macrophage (CD68+) content increased dramatically by Day 2. Between 2 and 5 days, macrophages increased further, whereas neutrophils declined to a level that in old muscles was not different from uninjured controls. M2 macrophages (CD163+) also increased between 2 and 5 days, reaching higher levels in muscles of old mice than in young mice. Although no evidence of persisting neutrophils or reduced M2 accumulation in old muscle was found, total macrophage accumulation was lower in old mice. Furthermore, messenger RNA levels showed age-related changes in macrophage-associated genes that may indicate alterations in myeloid cell function. Overall, differences between muscles of old and young mice in the inflammatory response through the early stages of injury may contribute to defects in muscle regeneration. Skeletal muscle, Macrophages, Neutrophils, Damage, Force deficit Skeletal muscle injuries are common and have a variety of causes, including trauma, ischemia/reperfusion, toxin exposure, and unloading/reloading. Muscles can also be injured by their own contractions, especially during lengthening contractions (LCs) when muscles are stretched while activated. Regardless of the cause, the initial insult is followed by degeneration of damaged muscle fibers and subsequent regeneration and repair. Delayed and/or incomplete regeneration of muscles of old animals has been widely reported. Specifically, muscles of old animals have fewer and smaller regenerating fibers (1,2), more fibrosis (3,4), and a greater deposition of fat (4) compared with young animals following injury. Failed restoration of normal muscle structure is accompanied by persistent deficits in force generation (5,6). Although aging-associated impairments in muscle repair are well established, the underlying mechanisms are incompletely understood. A growing body of evidence supports an important role for myeloid cells in regulating the progression of muscle degeneration and regeneration following injury. Neutrophils are the first to accumulate in injured muscle, arriving within hours of the initial insult to facilitate the removal of damaged or necrotic tissue. Neutrophils can also exacerbate damage to muscle fibers through mechanisms involving reactive oxygen species (7), and preventing neutrophil infiltration to muscle after injury has been reported to reduce force deficits and histological damage to muscle fibers (8,9). Macrophage levels increase later and contribute to muscle repair by mechanisms that depend on macrophage phenotype (10,11). Macrophages are generally defined as proinflammatory (M1) and anti-inflammatory (M2), although these classifications are overly simplistic, especially for cells in vivo. In general, M1 macrophages contribute to recovery from injury by removing damaged tissue and then decline in number as M2 macrophages increase coincident with the regenerative stage of muscle repair (10,11). The accumulating evidence that myeloid cells play a critical role in regulating muscle degeneration and regeneration implies that an inappropriate myeloid cell response could contribute to impairments in muscle repair following injury. Thus, based on known muscle regeneration defects observed in old animals, we asked whether there are differences between young and old mice in the timing, magnitude, and/or characteristics of the myeloid cell response to muscle injury. We specifically hypothesized that neutrophils would persist in injured muscles of old compared with young animals and that the accumulation of macrophages, especially M2 macrophages would be delayed or reduced in old muscles. To test our hypothesis, we injured muscles of young and old mice with LCs in situ and analyzed the muscles 2 or 5 days later, using histological, immunohistochemical, and functional measures, as well as protein and mRNA analyses. Method Animals Young (3–5 months) and old (25–27 months) male C57BL6 mice were maintained in a specific-pathogen-free facility at the University of Michigan (UM) until they were used for experimentation. Between procedures, mice were housed in a separate specific-pathogen-free return room. Animal procedures were approved by the UM Institutional Animal Care and Use Committee. In Situ Evaluation of Contractile Properties Procedures for in situ evaluation of muscle contractile properties were based on the previous studies (12). Mice were anesthetized with 3% isoflurane in oxygen. Anesthesia was maintained with 2% isoflurane in oxygen and confirmed by lack of response to tactile stimuli. Ophthalmic ointment was applied to prevent corneal drying. Hind limb fur was removed, and the skin was disinfected with chlorhexidine. A small incision was made at the ankle to expose the distal tendon of the extensor digitorum longus (EDL) muscle, and 6-0 braided silk was tied around the tendon. Another small incision just distal to the knee exposed the fibular nerve. The mouse was placed on a 37°C platform, and the hind limb was immobilized by pinching the knee and foot with small clamps secured to the platform. Using the tails of the silk suture, the intact tendon was tied to the lever arm of a servomotor (300C-LR-FP, Aurora Scientific) that controlled muscle length and measured force. Custom-designed software controlled stimulus pulses and motor movement and collected force data. The small area of exposed tendon was kept moist by frequent administration of warmed sterile saline. The EDL muscle was activated with platinum electrodes placed under the fibular nerve. Stimulus pulses of 0.2-ms duration were applied using an Aurora Scientific 701C high-power current stimulator, and current and muscle length were adjusted to elicit maximum twitch force. Tetanic contractions were elicited with 200-ms trains of pulses. The frequency of pulses was increased during successive contractions until maximum isometric force (Po) was achieved, typically at 200 Hz. Optimal muscle length (Lo) was measured with calipers, and optimal fiber length (Lf) was determined by multiplying Lo by the previously determined Lf-to-Lo ratio of 0.45 (13). In Situ LC Protocol To induce muscle injury, we used the physiologically relevant approach of exposure to LC. Our group has previously characterized many aspects of the degeneration and regeneration responses following LC-induced injury and demonstrated the ability to administer well-controlled contraction protocols resulting in highly reproducible injury responses (6,8,12,14,15). In the present study, the EDL muscle was exposed to a protocol of 75 LCs spaced 4 seconds apart for a total duration of 5 minutes. Each contraction was 300 ms in duration. A stretch of 20% strain relative to Lf was initiated 100 ms after the onset of stimulation from near maximum isometric force. Muscles were lengthened at 1 Lf/s such that the peak of the stretch coincided with the end of stimulation. After the LC protocol, a final measurement of maximum isometric force was made. Force decreased 49% ± 9% (SD) and 58% ± 12% (SD) for muscles of young and old mice, respectively, reflecting both injury and fatigue. After completion of the in situ experiments, the small incisions at the ankle and knee were closed with 7-0 sterile monofilament nylon suture and bathed with povidone-iodine solution, and mice were monitored until they recovered from anesthesia. In Vitro Evaluation of Contractile Properties Two or five days after LCs, injured and contralateral control muscles were evaluated for Po in vitro as described (13). Mice were anesthetized with an intraperitoneal injection of Avertin (tribromoethanol, 250 mg/kg), and EDL muscles were carefully isolated and removed. Mice were then euthanized with an overdose of Avertin followed by induction of a bilateral pneumothorax. 5-0 silk suture was tied to the proximal and distal tendons of the muscle, which was then placed in a chamber containing Krebs Mammalian Ringer solution composed of (in millimolar) 137 NaCl, 5 KCl, 2 CaCl2·2H2O, 1 MgSO4·7H2O, 1 NaH2PO4, 24 NaHCO3, 11 glucose, and 0.03 tubocurarine chloride. The bath was held at 25°C and bubbled with 95% O2–5% CO2 to maintain a pH of 7.4. The tendons were tied to a force transducer (BG-50, Kulite Semiconductor Products) and a fixed post. Field stimulation (Aurora 701C stimulator) was applied via parallel plate electrodes. Contractile properties were determined as described above except that tetanic contractions were elicited with 300-ms trains of pulses and Po occurred at ~150 Hz. The force deficit induced by the LCs was calculated as the difference between Po measured for the muscle exposed to LCs and the contralateral uninjured control muscle expressed as a percentage of the contralateral control Po. Following force measurements, muscles were trimmed of tendons, weighed, and cut in half. One half was immersed in tissue freezing medium and frozen in isopentane cooled by liquid nitrogen; the other half was submerged in an RNA stabilization reagent (RNAlater, QIAGEN). Histology, Immunohistochemistry, and Immunofluorescence We have previously shown that muscle function, histology, and inflammatory cell levels were not influenced by the surgical procedures associated with the LC protocol (12,16). We have now performed sham operations on old mice and confirmed that no changes were observed in neutrophil or macrophage levels compared with undisturbed control muscles in the days following the surgery. Thus, contralateral muscles from mice in which the ipsilateral muscle had been exposed to LCs served as controls for histology. Cryosections of 10 µm were fixed in cold acetone and stained with hematoxylin (Ricca Chemical Company) and eosin Y (EMD Millipore; Supplementary Figure S1A). In single sections, injured fibers were counted as those with pale or variable staining, swollen appearance, or obvious infiltration by inflammatory cells (8). Basophilic fibers with central nuclei were identified as regenerating (17). Image J was used to count fibers and measure cross-sectional areas of individual fibers and entire sections. Total section area was multiplied by section thickness to calculate volume. Muscle sections were interrogated for neutrophils and macrophages by immunohistochemistry. Others report that mouse CD68 (rat ED1+) labels early-invading macrophages that accumulate in injured skeletal muscle and invade muscle fibers (10,11), whereas mouse CD163 (rat ED2+) labels macrophages that appear later and are associated with repair (10,11). Accordingly, we chose CD68 and CD163 as markers of M1 and M2 macrophages, respectively. To examine the specificity and selectivity of these markers, we used immunofluorescence to label muscles for CD68 and CD163 or CD163 and CD301. CD301 is reported to be a marker of M2 macrophages in mice (18). Sections were fixed in cold acetone, air-dried, and exposed to 1% bovine serum albumin (Jackson ImmunoResearch Laboratories) in phosphate-buffered saline (PBS) to block nonspecific binding. Macrophages were detected with a rat anti-mouse CD68 (FA-11, 1:1,000; AbD Serotec), rabbit anti-mouse CD163 (M-96, 1:200; Santa Cruz Biotechnology), or goat anti-mouse CD301 IgG (1:400; R&D Systems) antibodies. Secondary antibodies were Alexa Fluor 594 goat anti-rat IgG, Alexa Fluor 594 bovine anti-goat IgG, and Alexa Fluor 488 donkey anti-rabbit IgG, all from Jackson ImmunoResearch Laboratories. Last, sections were exposed to 4′,6-diamidino-2-phenylindole (D9564; Sigma–Aldrich, St. Louis, MO) and treated with ProLong Gold (P36930; Life Technologies). These analyses showed that CD163+ cells were also CD68+ (Supplementary Figure S2), indicating that CD68 and CD163 are not distinguishing markers of macrophage phenotype in injured mouse EDL muscle. Rather, CD163 labels a subset of the macrophages also labeled by CD68. CD163+ cells were also positive for CD301 (Supplementary Figure S3), supporting the identification of these cells as M2 macrophages (18). Therefore, CD68 was considered to be a general macrophage marker and CD163 a marker of M2 macrophages. To quantify neutrophils and macrophages, sections were fixed in cold acetone, air-dried, and exposed to BLOXALL solution to block endogenous peroxidase activity followed by 10% normal rabbit or goat serum in PBS (Fisher Scientific) to block nonspecific binding. Neutrophils were detected with a rat anti-mouse Gr-1 antibody (RB6-8C5, 1:50 dilution; BD Pharmingen) diluted in PBS containing 10% rabbit serum, and macrophages were detected with a rat anti-mouse CD68 antibody (FA-11, 1:1,000; AbD Serotec) diluted in PBS containing 10% rabbit serum or with a rabbit anti-mouse CD163 antibody (M-96, 1:1,000; Santa Cruz Biotechnology) diluted in PBS containing 10% goat serum. After incubation in primary antibodies, sections were exposed to biotinylated mouse adsorbed rabbit anti-rat IgG (Gr-1 and CD68) or biotinylated goat anti-rabbit IgG (CD163) in PBS containing 10% rabbit or goat serum, respectively. Sections were treated with VECTASTAIN Elite ABC reagent containing peroxidase followed by peroxidase substrate 3, 3′-diaminobenzidine (ImmPACT DAB). All reagents were from Vector Laboratories unless stated otherwise. Pilot studies showed no regional differences (proximal, central, or distal portion of the muscle) in inflammatory cell infiltration. Therefore, Image J was used to count neutrophils and macrophages in single sections from the belly of each muscle. RB6-8C5 (Gr-1) reacts with both Ly6G and Ly6C (19), and neutrophils display abundant expression of both, with Ly6G levels in peripheral neutrophils directly correlated with the cell’s level of differentiation and maturation (20). Despite the widespread usage of Gr-1 as a neutrophil marker, Ly6C is also expressed at lower levels by a number of other cell types including monocytes, eosinophils, dendritic cells, a wide range of endothelial cells, and subsets of T cells (21,22). Thus, dark brown cells indicating high Gr-1 levels were counted as neutrophils (18), whereas the occasional light brown cell of similar size was not counted (Supplementary Figure S1B). Both light and dark brown cells were counted as macrophages in sections treated with reagents for CD68 (Supplementary Figure S1C) and CD163 (Supplementary Figure S1D) detection. In all cases, cells at the extreme periphery of the section were not counted. Western Blots For analysis of CD68 and CD163 levels, EDL muscles were placed in bead homogenization tubes with RIPA buffer containing 50 mM Tris (pH 7.4), 150 mM NaCl, and protease inhibitors (Thermofisher Scientific, Waltham, MA), whereas for analysis of Ly6G/Ly6C, muscle tissues were homogenized in 20 mM Tris buffer with 0.05% Triton X and the protease inhibitors. All samples were homogenized using a Bead Mill 4 homogenizer (Fisher Scientific, Hampton, NH) for two cycles at 4 m/s for 45 seconds with a 1-minute rest period. After homogenization, samples were centrifuged at 13,000g for 15 minutes and transferred to 1.5-mL microcentrifuge tubes. Protein content was determined using the bicinchoninic acid method (Sigma–Aldrich). For CD68 and CD163, muscle homogenates (20 µg) were fractionated in 4%–20% Mini-PROTEAN TGX precast gels, and gels were transferred to polyvinylidene difluoride membranes. Membranes were stained with Ponceau S before blotting to verify equal loading of the gels. Membranes were blocked in 5% milk in Tris-buffered saline with 0.1% Tween-20 (TBST), for 1–2 hours. Antibodies for CD68 (ab201340; Abcam, Cambridge, UK) or CD163 (ab182422; Abcam, Cambridge, UK) were diluted 1:1,000 in 5% milk in TBST and incubated at 4°C overnight. For Ly6, muscle homogenates (20 µg) were fractionated in 10% native gels and placed in Tris–glycine buffer (pH 8.9) surrounded by Tris–glycine buffer (pH 8.3). Gels were transferred to polyvinylidene difluoride membranes in Tris/Glycine buffer 0.01% sodium dodecyl sulfate. In native conditions, the primary antibody for Ly6G/Ly6C (RB6-8C5; BD Pharmingen, San Jose, CA) was diluted 1:750 in 5% milk in TBST and incubated at 4°C overnight. Anti-rabbit and anti-mouse monoclonal secondary antibodies (Cell Signaling Technologies, Danvers, MA) were diluted 1:2,000 in 5% milk, in TBST, and then incubated at room temperature for 1 hour. Enhanced chemiluminescence was performed using Chemidoc XRS imaging system (Bio-Rad, Hercules, CA) to visualize antibody–antigen interaction. Blotting images were quantified by densitometry using myImageAnalysis software (Thermofisher Scientific). The Ponceau-stained membranes were digitally scanned, and the 45-kDa actin bands were quantified by densitometry and used as a protein loading correction factor for each lane. An internal control was loaded on all gels to make quantitative comparisons between gels. Gene Expression Muscle samples were homogenized in QIAzol lysis reagent (QIAGEN). RNA was isolated using a miRNeasy Mini Kit and treated with DNase I (QIAGEN). RNA concentration was measured with a NanoDrop 2000 Spectrophotometer (Thermo Scientific). RNA (2 µg) was reverse transcribed using a QuantiTect Reverse Transcription Kit (QIAGEN), and cDNA was diluted by fivefold and amplified in a CFX96 real time thermal cycler (Bio-Rad) using a QuantiTect SYBR Green I PCR Kit (QIAGEN) and primers specific for inducible nitric oxide synthase (iNOS) (23), tumor necrosis factor alpha (TNFα) (23), arginase-1 (Arg1) (10), interleukin-10 (IL-10) (10), and β2-microglobulin (15). Primers were purchased from Integrated DNA Technologies and are listed in Supplementary Table S1. mRNA from target genes was undetectable in control muscles, so we completed our analysis using only injured muscles. We were unable, despite multiple trials, to identify a housekeeping gene that remained stable both between age groups and over time following LCs. This is perhaps not surprising, due to the ongoing breakdown of the tissue and the ongoing accumulation in the muscle of different cell types. Because mRNA levels of β2-microglobulin were not different between age groups for any of the experimental conditions and our primary interest was in identifying age-associated differences, we normalized mRNA transcripts to β2-microglobulin and compared young and old samples at each time point. mRNA levels were analyzed by the comparative CT method (24). Blood Analysis Blood (50–100 µL) was collected from a separate group of undisturbed control mice via a scalpel nick of the lateral tail vein. White blood cell counts were determined in whole blood using a cell counter (Hemavet 950FS, Drew Scientific). Data Analysis Data are reported as means ± 1 SEM. Measures of injury and levels of myeloid cells and protein were compared between age groups and across time using a two-factor analysis of variance (ANOVA). In cases when the ANOVA showed significant effects, the Holm-Sidak post hoc test was used to identify individual differences. When comparisons were between only two groups, for example, regenerating fibers in muscles of young and old mice, Student’s t tests were used. In all cases, significance was set a priori at p < .05, and individual p values are included throughout the text and in figure legends and tables. Results Muscle masses were not different between young and old mice, but muscles from old mice were weaker by all measures (Table 1). Although muscles in old animals are reported to be more susceptible to contraction-induced injury (25,26), a common approach for comparing the progression of the injury process between young and old animals is to induce an injury of similar magnitude in both age groups (5,6,12,14). In the present study, the contraction protocol resulted in a similar level of injury in muscles of young and old mice as indicated by no statistically significant differences in the measures of injury after 2 days. Force deficits of ~55% were not different (p = .183) for muscle of young and old mice (Figure 1A), and there was also no difference between the age groups (p = .336) in the number of damaged fibers appearing in muscle cross sections at 2 days (Figure 1B). For young mice, both the force deficit (p = .007) and the number of damaged fibers (p < .001) increased between 2 and 5 days, and the total number of fibers in a cross section decreased (p < .001). The number of damaged fibers increased (p = .007) between 2 and 5 days for old mice, as well, but the number of fibers showing histological evidence of damage was ~35% lower (p = .005) for muscles of old mice compared with young mice at 5 days (Figure 1B) and the total number of fibers remaining in cross section 5 days after LCs was ~15% greater (p = .039) in muscles of old mice compared with young mice (Figure 1C). Table 1. Characteristics of Young and Old Groups Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Note: CSA = cross-sectional area; H&E = hematoxylin and eosin. Forces are maximum forces generated during isometric tetanic contractions. [Two samples are missing from young group for CSA from H&E stained sections and for total number of fibers (ie, n = 16).] aData taken from contralateral muscles. *Significant difference from young group (p < .05). p Values are provided from t tests for each variable. View Large Table 1. Characteristics of Young and Old Groups Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Note: CSA = cross-sectional area; H&E = hematoxylin and eosin. Forces are maximum forces generated during isometric tetanic contractions. [Two samples are missing from young group for CSA from H&E stained sections and for total number of fibers (ie, n = 16).] aData taken from contralateral muscles. *Significant difference from young group (p < .05). p Values are provided from t tests for each variable. View Large Figure 1. View largeDownload slide Lengthening contractions damage muscles of young and old mice. The severity of injury was assessed 2 and 5 d after LCs by (A) the deficit in isometric force generating capacity expressed as a percentage of the contralateral control muscle, (B) the percentage of fibers in a single cross section showing histological evidence of damage, and (C) the total number of fibers appearing in a cross section. Data are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for force deficit (Time: p = .008; Age: p = .004; Time × Age: p = .231) and number of injured fibers (Time: p < .001; Age: p = .008; Time × Age: p = .151) and significant effects of time with significant interactions for the number of fibers in cross sections with no main effect of age (Time: p < .001; Age: p = .319; Time × Age: p = .045). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 1. View largeDownload slide Lengthening contractions damage muscles of young and old mice. The severity of injury was assessed 2 and 5 d after LCs by (A) the deficit in isometric force generating capacity expressed as a percentage of the contralateral control muscle, (B) the percentage of fibers in a single cross section showing histological evidence of damage, and (C) the total number of fibers appearing in a cross section. Data are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for force deficit (Time: p = .008; Age: p = .004; Time × Age: p = .231) and number of injured fibers (Time: p < .001; Age: p = .008; Time × Age: p = .151) and significant effects of time with significant interactions for the number of fibers in cross sections with no main effect of age (Time: p < .001; Age: p = .319; Time × Age: p = .045). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. ANOVA = analysis of variance; LCs = lengthening contractions. Regenerating fibers were clearly present in muscle cross sections 5 days after LCs (Figure 2A and B), with the number of regenerating fibers in injured muscles from old mice nearly 40% lower (p = .050) than in young mice (Figure 2C). In addition to counting the number of regenerating fibers, previous studies have also used the cross-sectional areas of regenerating fibers to compare groups for the rate and extent of regeneration (2,9). Thus, we also analyzed muscle fiber cross-sectional areas as a measure of regeneration. Regenerating fibers in muscle sections from old mice were 20% smaller (p < .001) than those of young mice at Day 5 (Figure 2D). Collectively, these findings are consistent with widely published reports of delayed or deficient repair in muscles of old animals. Figure 2. View largeDownload slide Regeneration is impaired in old mice. Representative partial sections of muscles stained with hematoxylin and eosin from (A) young and (B) old mice show regenerating fibers. Scale bar = 50 µm and applies to both images. Data are shown for (C) the number of regenerating fibers per muscle cross section and (D) average cross-sectional area of regenerating fibers 5 d after lengthening contractions. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Figure 2. View largeDownload slide Regeneration is impaired in old mice. Representative partial sections of muscles stained with hematoxylin and eosin from (A) young and (B) old mice show regenerating fibers. Scale bar = 50 µm and applies to both images. Data are shown for (C) the number of regenerating fibers per muscle cross section and (D) average cross-sectional area of regenerating fibers 5 d after lengthening contractions. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Muscle degeneration and regeneration were accompanied by accumulation of myeloid cells in injured muscles. Two days after LCs, neutrophil content was dramatically elevated relative to control levels in both young and old mice, as evidenced by both numbers of Gr-1+ cells (p < .001; Figure 3A) and Ly6G/Ly6C levels (Figure 4A), with numbers of neutrophils 35% higher (p = .002) in muscles of old than in young mice at this time point. Consistent with previous reports of rapid clearance of neutrophils from injured muscles (9,27), both age groups showed declines in neutrophil numbers of ~70% (p < .001) between 2 and 5 days. The decrease in neutrophils resulted in levels in old mice by the 5-day time point that were not different (p = .088) from baseline values, although baseline neutrophil numbers in old mice were elevated (p = .024) compared with control muscles of young mice (Figure 3A). High variation in the old muscles for Ly6G/Ly6C levels following injury resulted in only a trend (p = .067) for an effect of age by two-factor ANOVA, but consistent with higher baseline numbers of neutrophils in old muscles compared with young muscles, approximately fourfold higher baseline levels of Ly6G/Ly6C in old control muscles were highly significant (p < .001) by t test. Ly6G/Ly6C levels remained elevated at 5 days for both age groups (Figure 4A), probably reflecting Ly6C expression by other cell types, in particular activated macrophages, which are present in 100-fold greater abundance than neutrophils at 5 days (Figure 3A and B). Figure 3. View largeDownload slide Myeloid cell accumulation is altered in injured muscles from young and old mice. Data are shown for (A) neutrophil (Gr-1+ cells), (B) macrophage (CD68+ cells), and (C) M2 macrophage (CD163+ cells) content expressed as the number of cells per volume of muscle in uninjured control muscles and 2 and 5 d after LCs. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for neutrophil (Time: p < .001; Age: p < .001; Time × Age: p = .413) and M2 macrophage (Time: p = .039; Age: p < .001; Time × Age: p = .217) number and significant effects of time and age with significant interactions for the number of CD68+ macrophages (Time: p = .002; Age: p < .001; Time × Age: p < .001). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 3. View largeDownload slide Myeloid cell accumulation is altered in injured muscles from young and old mice. Data are shown for (A) neutrophil (Gr-1+ cells), (B) macrophage (CD68+ cells), and (C) M2 macrophage (CD163+ cells) content expressed as the number of cells per volume of muscle in uninjured control muscles and 2 and 5 d after LCs. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for neutrophil (Time: p < .001; Age: p < .001; Time × Age: p = .413) and M2 macrophage (Time: p = .039; Age: p < .001; Time × Age: p = .217) number and significant effects of time and age with significant interactions for the number of CD68+ macrophages (Time: p = .002; Age: p < .001; Time × Age: p < .001). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 4. View largeDownload slide Protein levels for myeloid cell markers increase following injury and with age. Data are shown for (A) Gr-1 (Ly6G and Ly6C), (B) CD68, and (C) CD163 protein levels in uninjured control muscles and 2 and 5 d after LCs determined by Western blot. Representative blots are shown in the right-hand panels. For analysis, protein levels were normalized to Ponceau S bands, which are also shown. Internal controls, randomly selected aged myotoxin-injected muscles, are included to allow densitometry measures to be compared between different blots. Left panels show average values expressed relative to the level in young control muscles and presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for CD68 levels (Time: p = .001; Age: p < .001; Time × Age: p = .377) and significant effects of time with strong trends for effects of age for Gr-1 (Time: p = .015; Age: p = .067; Time × Age: p = .398) and CD163 (Time: p < .001; Age: p = .078; Time × Age: p = .879) levels. Sample sizes are n = 4 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 4. View largeDownload slide Protein levels for myeloid cell markers increase following injury and with age. Data are shown for (A) Gr-1 (Ly6G and Ly6C), (B) CD68, and (C) CD163 protein levels in uninjured control muscles and 2 and 5 d after LCs determined by Western blot. Representative blots are shown in the right-hand panels. For analysis, protein levels were normalized to Ponceau S bands, which are also shown. Internal controls, randomly selected aged myotoxin-injected muscles, are included to allow densitometry measures to be compared between different blots. Left panels show average values expressed relative to the level in young control muscles and presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for CD68 levels (Time: p = .001; Age: p < .001; Time × Age: p = .377) and significant effects of time with strong trends for effects of age for Gr-1 (Time: p = .015; Age: p = .067; Time × Age: p = .398) and CD163 (Time: p < .001; Age: p = .078; Time × Age: p = .879) levels. Sample sizes are n = 4 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. In contrast to neutrophils, baseline numbers of macrophages (CD68+ cells) were not different (p = .948) in muscles of young and old mice, and both age groups showed similar fivefold increases (p < .001) in CD68+ macrophage content relative to their respective control levels 2 days after LCs (Figure 3B). Between 2 and 5 days, macrophage content increased another threefold to fourfold (p < .001) for both age groups but to a lesser extent in old mice resulting in macrophage numbers in muscles of old mice that were ~25% lower (p < .001) than in young muscles at 5 days (Figure 3B). Although accumulation of macrophages was lower in muscles of old mice at 5 days, CD68 levels were not different (p = .546) in muscles of young and old mice at that time point (Figure 4B), perhaps reflecting a compensatory response to a decrease in function, although this is speculation. The function of CD68 is not clear beyond its role targeting macrophages to tissues. Control levels of M2 macrophages (CD163+ cells) were also not different (p = .666) for muscles of young and old mice and did not increase for either age group (young: control vs. 2 day, p = .629; old: control vs. 2 day, p = .833) until 5 days following LCs (young and old: control vs. 5 day, p < .001) concurrent with the regenerative phase. The increase in M2 macrophages between 2 and 5 days (p < .001) was greater for muscles of old mice resulting in levels of M2 macrophages at 5 days that were ~40% higher (p = .017) in old muscles compared with young muscles (Figure 3C). CD163 protein levels assessed by western blot were generally consistent with the cell numbers determined through immunohistochemistry, increasing fourfold to fivefold (p < .001) for both age groups by Day 5 and showing a trend for increased levels (p = .078) in muscles of old mice compared with young mice (Figure 4C). The effect of differences in myeloid cell content in muscles of young and old mice may either be exacerbated or mitigated by alterations in myeloid cell function. Alternatively, differences in cellular accumulation may represent compensatory responses to poorly functioning myeloid cells. To begin to explore these questions, we examined messenger RNA levels for genes expressed by M1 (iNOS, TNFα) or M2 (Arg1, IL-10) macrophages in injured muscles from young and old mice (Figure 5). In old injured muscles, mRNA levels for iNOS (p = .621) and TNFα (p = .667) were not different from young muscles 2 days after LCs (Figure 5A and C), but were significantly elevated (iNOS; p = .021; TNFα, p = .014) over young levels at 5 days (Figure 5B and D). mRNA levels for Arg1 were also not different (p = .439) between young and old muscles 2 days after LCs (Figure 5E). After 5 days, Arg1 mRNA level in old muscles appeared to increase relative to young, but values from old animals were highly variable and the difference was not significant (Figure 5F, p = .130). Two days after LCs, IL-10 mRNA levels were lower (p = .009) in old muscles compared with young muscles (Figure 5G) with no differences (p = .280) between age groups at 5 days (Figure 5H). Figure 5. View largeDownload slide Messenger RNA levels of M1 (iNOS, TNFα) and M2 (Arg1, IL-10) macrophage genes are altered with age. Shown are messenger RNA levels for iNOS (A, B), TNFα (C, D), Arg1 (E, F), and IL-10 (G, H) normalized to β2-microglobulin mRNA 2 d (A, C, E, G) and 5 d (B, D, F, H) after lengthening contractions for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Arg1 = arginase-1; IL-10 = interleukin-10; iNOS = inducible nitric oxide synthase; LC = lengthening contraction; TNFα = tumor necrosis factor alpha. Figure 5. View largeDownload slide Messenger RNA levels of M1 (iNOS, TNFα) and M2 (Arg1, IL-10) macrophage genes are altered with age. Shown are messenger RNA levels for iNOS (A, B), TNFα (C, D), Arg1 (E, F), and IL-10 (G, H) normalized to β2-microglobulin mRNA 2 d (A, C, E, G) and 5 d (B, D, F, H) after lengthening contractions for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Arg1 = arginase-1; IL-10 = interleukin-10; iNOS = inducible nitric oxide synthase; LC = lengthening contraction; TNFα = tumor necrosis factor alpha. Finally, we questioned whether neutrophil and macrophage content in injured muscles from young and old mice reflected the baseline circulating levels of neutrophils and monocytes with age. In mice that were not subjected to LCs, circulating levels of neutrophils and monocytes were significantly greater (neutrophils, p = .008; monocytes, p = .016) in old animals relative to young animals (Figure 6). Figure 6. View largeDownload slide Circulating levels of neutrophils and monocytes are elevated in old mice. Mean (A) neutrophil and (B) monocyte levels expressed as thousand cells per microliter blood ± SEM are shown for young (black bars) and old (gray bars) mice. Sample sizes are n = 6 per group. Differences (p < .05) between individual groups are indicated by brackets. Figure 6. View largeDownload slide Circulating levels of neutrophils and monocytes are elevated in old mice. Mean (A) neutrophil and (B) monocyte levels expressed as thousand cells per microliter blood ± SEM are shown for young (black bars) and old (gray bars) mice. Sample sizes are n = 6 per group. Differences (p < .05) between individual groups are indicated by brackets. Discussion The goal was to examine the effects of aging on the timing and magnitude of the myeloid cell response to muscle injury. Injury was induced in muscles of young and old mice by exposure to LCs. The amount of injury observed in the present study was comparable qualitatively and quantitatively to previous studies from our laboratory that used the same contraction protocol (28). Moreover, the use of a contraction protocol that resulted in similar magnitudes of injury in muscles of young and old mice 2 days following LCs, as indicated by no differences between the age groups in force deficit or histological evidence of damage, allowed us to evaluate and compare the progression of degeneration and regeneration in the two age groups starting from roughly equivalent initial levels of damage. This is a strategy that has been used previously (5,6,12,14). Overall, our results support alterations in the myeloid cell response to LC-induced injury in old mice relative to young mice. Consistent with previous reports (8,9), we observed a robust increase in neutrophils following LCs in both young and old muscles. We did not, however, observe a persistence of neutrophils in old muscles, as expected. This conclusion is based on our findings that neutrophils declined dramatically between 2 and 5 days in both young and old mice. This interpretation is complicated somewhat by our observation that Gr-1 protein levels did not show a similar decline between 2 and 5 days. The significance of the continued elevation of Ly6G/Ly6C protein levels above control levels is not clear, but it indicates that protein levels are not a good indicator of neutrophil numbers when large numbers of macrophages and/or other cell types that also express Ly6C are also present. Our observation that the neutrophil level remained higher in muscles of old mice compared with young mice 5 days after LCs is not interpreted as a persistence of the neutrophils induced by injury, because the tissue neutrophil level in old mice had returned at this time point to the level observed in control muscles. Although control levels of neutrophils were rapidly reestablished in muscles of old mice following LCs, the elevated baseline neutrophil content in muscles of old mice compared with young mice suggests either underlying injury in old mice (29,30) or the presence of elevated proinflammatory signals in muscles in the absence of injury (31–33). The impact of an age-associated increase in baseline inflammation has been hypothesized to contribute to age-associated muscle wasting through effects on myogenesis as well as the balance between protein synthesis and degradation (34,35) and may also influence repair from damage. A second unexpected finding was that accumulation of M2 macrophages was not impaired in muscles of old mice. We had predicted delays or reductions in old animals in the accumulation of M2 macrophages following LCs based on the role of M2 macrophages to promote muscle regeneration (10,36,37) and the generally accepted view that muscle regeneration is impaired with aging (1,2,4–6,14,38–43). M2 macrophages have been reported to stimulate proliferation and differentiation of muscle precursor cells in vitro (10,36,37). Thus, our observation of fewer and smaller regenerating fibers in injured muscles of old mice despite no impairment in the accumulation of M2 macrophages suggests that macrophage function may be impaired in old muscles. Although we did not observe reduced M2 macrophage accumulation in old muscles, we cannot rule out the possibility that the levels continue to increase beyond 5 days and old mice would ultimately display a defect in M2 macrophage accumulation. However, Wang and colleagues (44) recently reported higher levels of CD163+ M2a macrophages in muscles of old mice compared with young mice, a finding that is potentially consistent with the present study and our own data showing ~20% higher baseline levels of M2 macrophages in old muscles (young: 10,130 ± 348 CD163+ cells per mm3 tissue vs. old: 12,070 ± 421 cells per mm3 tissue), although the difference was not statistically significant (p = .666). The finding of increases in young mice between 2 and 5 days for both force deficit and the number of damaged fibers along with a loss of fibers from cross sections indicates that muscle fiber degeneration continued in young mice throughout the time period studied. In contrast, no change in the force deficit for old mice along with the smaller increase in the percentage of fibers showing histological evidence of damage in muscles of old mice compared with young mice between 2 and 5 days is consistent with a diminished degeneration response for old mice. In addition, the total number of fibers remaining in cross section at 5 days was slightly greater in muscles of old mice compared with young mice, suggesting that fewer muscle fibers had completely degenerated in old mice at this time point. This apparent blunting of degeneration in old mice was associated with a reduction in overall macrophage accumulation, as evidenced by our observation of 25% fewer CD68+ cells in muscles of old mice compared with young mice 5 days following exposure to LCs. Although based on the present data set, we cannot definitively establish whether the fewer macrophages observed in muscles of old mice compared with young mice 5 days following LCs are a cause or consequence of the reduced level of injury observed at that time point, the similarity in force deficits, total fiber number, and number of damaged fibers in muscles of young and old mice at 2 days argues against the possibility that less severe injury was a major driver of lower macrophage accumulation in old muscles. Based on the known phagocytic function of M1 macrophages (11,36), the observation of fewer CD68+ macrophages in muscles of old mice is consistent with the hypothesis that the phagocytic removal of damaged tissue would be less effective in muscles of old mice (36). Higher CD68 levels observed in muscles of old compared with young mice may reflect a compensatory response to an age-related decrease in cell number, but this is entirely speculative at this stage and future studies are necessary to examine the phagocytic activity of macrophages from old muscles. Previous investigations comparing leukocyte accumulation in muscles from young and old animals in response to injury have generated conflicting conclusions. For example, muscles from old rats displayed elevated CD68 protein content compared with muscles of young rats 3 days after a contusion injury (3), and using immunohistochemistry, Koh and colleagues (12) reported almost twofold higher numbers of neutrophils (Ly6G+ cells) and 50% more macrophages (F4/80+ cells) in muscles of old mice compared with young mice 3 days after LCs. Similarly, following ischemia/reperfusion injury, plantaris muscles of old rats showed greater leukocyte infiltration than that observed in young rats, but specific cell types were not identified (45). Finally, prolonged expression (mRNA) of the leukocyte marker CD18 was reported in muscles of old rats after myotoxin injection (43), although this marker cannot distinguish between persistence of neutrophils and an age-related increase in macrophage levels. Contrary to these prior reports that generally support an age-related increase in myeloid cell content after injury, Paliwal and colleagues (41) found no difference in the numbers of CD11b+ cells, as assessed by flow cytometry, in muscles of young and old mice 3 and 5 days after cardiotoxin injection, and Hamada and colleagues (46) suggested delayed or reduced leukocyte accumulation with advanced age based on their observation of a blunted increase in CD18 mRNA levels in vastus lateralis muscles of old mice relative to young mice 3 days after downhill running. Finally, Sadeh (2) reported delayed recruitment of macrophages in muscles of old rats compared with young rats injured with myotoxin, but macrophage numbers were estimated based on counting nuclei in muscle sections stained with H&E, which would also include other cell types. The bases for these disparate findings are not known, but they highlight the importance of quantifying the extent of damage induced and combining approaches for assessing leukocyte numbers and function as we have begun to do here. Full interpretation of our findings must also consider the levels of myeloid cells within tissues before the injury. Indeed, we observed substantially more neutrophils in uninjured control muscles of old mice relative to young mice. In contrast, the greater LC-induced accumulation of M2 macrophages in muscles of old mice occurred with no significant difference between muscles of young and old mice in M2 macrophage levels either at baseline or 2 days following LCs. Thus, the higher levels of M2 macrophages are not merely a reflection of differing control conditions. Numerous factors could play a role in differences in inflammatory cell numbers in old mice versus young mice including differences in chemoattractant release by the injured muscle, effectiveness of the homing mechanisms for leukocytes to injured sites, or alterations in the life span of inflammatory cells in the tissue. Our observation of elevated CD68 expression in muscles of old mice compared with young mice may represent a compensatory response to impairments in the ability of macrophages to effectively target the injured tissue. The precise signals that trigger inflammatory cell accumulation in muscle are not known, but TNFα, monocyte chemotactic protein-1 (MCP-1, CCL2), IL-8, granulocyte-macrophage colony stimulating factor, and cyclooxygenase (COX)-2 have all been reported to recruit neutrophils and macrophages to injured muscle (47). In addition, resident myeloid cells recruit other inflammatory cells to injured muscle (48). Additional experiments exploring age-associated changes in the levels of chemoattractant signals are warranted. Finally, consistent with our observation of elevated levels of circulating leukocytes in old mice compared with young mice, neutrophil and macrophage content in tissues may also be influenced by a larger pool of cells available to respond to the recruitment signals. Consistent with previous reports of elevations in iNOS, TNFα, Arg1, and IL-10 in injured muscles (10,23) and in myeloid cells isolated from injured muscles (23,36), we also detected iNOS, TNFα, Arg1, and IL-10 mRNA in injured muscles at both 2 and 5 days after LCs. Our finding that at 5 days, mRNA levels of iNOS and TNFα were elevated in muscles from old mice relative to young mice suggests that iNOS and TNFα were produced in excess by macrophages in the old mice. In support of this conclusion, macrophages have been identified as a source of TNFα and iNOS after muscle injury (23,36), and in the present study, iNOS and TNFα mRNA levels were elevated in muscles of old mice despite fewer CD68+ cells present in injured muscles from old compared with young mice at this time point. TNFα is reported to inhibit myogenic differentiation in vitro and in vivo, to lead to the degradation of transcription factors critical for muscle regeneration and to induce apoptosis (35,49), whereas iNOS-expressing macrophages can injure muscle cells in vitro (50) and in vivo (51). Therefore, excessive TNFα or iNOS production by macrophages in muscles of old mice could undermine or impair muscle regeneration in aged animals, although low levels of TNFα and iNOS expression are also reported to benefit muscle repair (23,35). Anti-inflammatory M2 macrophages have been identified as the primary source of IL-10 after muscle injury (36,52). We found less IL-10 mRNA in muscles of old mice relative to young mice at 2 days, in spite of no difference in the levels of CD163+ cells in young and old muscles at this time point. Thus, the age-related decrease in IL-10 mRNA is consistent with impaired IL-10 production in aged macrophages. Impaired IL-10 production can be detrimental to muscle repair. Mice lacking IL-10 showed persistent fiber damage and slowed regeneration and growth after an unloading/reloading muscle injury (10). IL-10 can contribute to repair by several mechanisms. IL-10-stimulated macrophages promote the proliferation and differentiation of myogenic precursors in vitro and in vivo (51,52). IL-10 can also play a regulatory role, suppressing the production of proinflammatory cytokines IL-1, IL-6, and TNFα (48), whereas stimulating production of the growth factor IGF-1 (52). Therefore, our observation of elevated levels of M2 macrophage content in injured muscles from old mice may reflect a compensatory effect of insufficient IL-10 production by these cells. Overall, our findings suggest age-related changes in the expression of macrophage-associated genes that have the potential to undermine or impair muscle regeneration in aged animals. Our study is not without limitations. We did not follow muscles out to full recovery. We cannot therefore definitively state that in the present study muscles of old mice would have failed to recover as quickly or completely as those in young mice; however, age-related impairments in regeneration after muscle injury resulting from a wide range of insults have been firmly established (1–4,38–43), including in studies from our own group using LCs (5,6,14). Thus, sacrificing additional animals to once again confirm this result does not appear justified. A second limitation is that our examination of mRNA levels for myeloid cell genes in the whole tissue provides no definitive information regarding the cellular source of the mRNA. Furthermore, altered mRNA levels do not necessarily reflect changes in protein content. Current studies are aimed at addressing these limitations by isolating different myeloid cell populations from injured muscles of young and old mice and examining mRNA and protein levels of purified cells. Despite some limitations, this study addresses gaps in our knowledge of aging-related changes in the myeloid cell response to injury, and our findings provide support for future studies specifically addressing the hypothesis that an effective degenerative phase is necessary to provide an environment amenable to effective regeneration. Supplementary Material Supplementary data is available at The Journals of Gerontology, Series A: Biological Sciences and Medical Sciences online. Funding This work was supported by the National Institute on Aging at the National Institutes of Health (AG020591 and AG051442 to S.V.B. and AG000114 to D.D.S. and L.A.B.). Conflict of Interest None reported. Acknowledgments We thank the Unit for Laboratory Animal Medicine (ULAM) Pathology Core for Animal Research (PCAR) at the University of Michigan for performing the blood cell counts. References 1. Conboy IM, Conboy MJ, Wagers AJ, Girma ER, Weissman IL, Rando TA. Rejuvenation of aged progenitor cells by exposure to a young systemic environment. Nature . 2005; 433: 760– 764. doi: 10.1038/nature03260 Google Scholar CrossRef Search ADS PubMed  2. Sadeh M. Effects of aging on skeletal muscle regeneration. J Neurol Sci . 1988; 87: 67– 74. doi:10.1016/0022-510X(88)90055-X Google Scholar CrossRef Search ADS PubMed  3. Ghaly A, Marsh DR. Aging-associated oxidative stress modulates the acute inflammatory response in skeletal muscle after contusion injury. Exp Gerontol . 2010; 45: 381– 388. doi: 10.1016/j.exger.2010.03.004 Google Scholar CrossRef Search ADS PubMed  4. Zhou Y, Lovell D, Bethea Met al.   Age-dependent changes cooperatively impact skeletal muscle regeneration after compartment syndrome injury. Am J Pathol . 2014; 184: 2225– 2236. doi: 10.1016/j.ajpath.2014.03.018 Google Scholar CrossRef Search ADS PubMed  5. McArdle A, Dillmann WH, Mestril R, Faulkner JA, Jackson MJ. Overexpression of HSP70 in mouse skeletal muscle protects against muscle damage and age-related muscle dysfunction. FASEB J . 2004; 18: 355– 357. doi: 10.1096/fj.03-0395fje Google Scholar CrossRef Search ADS PubMed  6. Rader EP, Faulkner JA. Recovery from contraction-induced injury is impaired in weight-bearing muscles of old male mice. J Appl Physiol (1985) . 2006; 100: 656– 661. doi: 10.1152/japplphysiol.00663.2005 Google Scholar CrossRef Search ADS PubMed  7. Nguyen HX, Tidball JG. Null mutation of gp91phox reduces muscle membrane lysis during muscle inflammation in mice. J Physiol . 2003; 553: 833– 841. doi: 10.1113/jphysiol.2003.051912 Google Scholar CrossRef Search ADS PubMed  8. Lockhart NC, Brooks SV. Neutrophil accumulation following passive stretches contributes to adaptations that reduce contraction-induced skeletal muscle injury in mice. J Appl Physiol (1985) . 2008; 104: 1109– 1115. doi: 10.1152/japplphysiol.00850.2007 Google Scholar CrossRef Search ADS PubMed  9. Pizza FX, Peterson JM, Baas JH, Koh TJ. Neutrophils contribute to muscle injury and impair its resolution after lengthening contractions in mice. J Physiol . 2005; 562: 899– 913. doi: 10.1113/jphysiol.2004.073965 Google Scholar CrossRef Search ADS PubMed  10. Deng B, Wehling-Henricks M, Villalta SA, Wang Y, Tidball JG. IL-10 triggers changes in macrophage phenotype that promote muscle growth and regeneration. J Immunol . 2012; 189: 3669– 3680. doi: 10.4049/jimmunol.1103180 Google Scholar CrossRef Search ADS PubMed  11. McLennan IS. Degenerating and regenerating skeletal muscles contain several subpopulations of macrophages with distinct spatial and temporal distributions. J Anat . 1996; 188: 17– 28. Google Scholar PubMed  12. Koh TJ, Peterson JM, Pizza FX, Brooks SV. Passive stretches protect skeletal muscle of adult and old mice from lengthening contraction-induced injury. J Gerontol A Biol Sci Med Sci . 2003; 58: 592– 597. doi:10.1093/gerona/58.7.B592 Google Scholar CrossRef Search ADS PubMed  13. Brooks SV, Faulkner JA. Contractile properties of skeletal muscles from young, adult and aged mice. J Physiol . 1988; 404: 71– 82. doi:10.1113/jphysiol.1988.sp017279 Google Scholar CrossRef Search ADS PubMed  14. Brooks SV, Faulkner JA. Contraction-induced injury: recovery of skeletal muscles in young and old mice. Am J Physiol . 1990; 258: C436– C442. doi: 10.1152/ajpcell.1990.258.3.C436 Google Scholar CrossRef Search ADS PubMed  15. Gumucio JP, Flood MD, Phan AC, Brooks SV, Mendias CL. Targeted inhibition of TGF-beta results in an initial improvement but long-term deficit in force production after contraction-induced skeletal muscle injury. J Appl Physiol (1985) . 2013; 115: 539– 545. doi: 10.1152/japplphysiol.00374.2013 Google Scholar CrossRef Search ADS PubMed  16. Pizza FX, Koh TJ, McGregor SJ, Brooks SV. Muscle inflammatory cells after passive stretches, isometric contractions, and lengthening contractions. J Appl Physiol (1985) . 2002; 92: 1873– 1878. doi: 10.1152/japplphysiol.01055.2001 Google Scholar CrossRef Search ADS PubMed  17. Shortreed K, Johnston A, Hawke TJ. Satellite cells and muscle repair. In: Tiidus PM, ed. Skeletal Muscle Damage and Repair . Champaign, IL: Human Kinetics; 2008: 77– 88. 18. Lumeng CN, DelProposto JB, Westcott DJ, Saltiel AR. Phenotypic switching of adipose tissue macrophages with obesity is generated by spatiotemporal differences in macrophage subtypes. Diabetes . 2008; 57: 3239– 3246. doi: 10.2337/db08-0872 Google Scholar CrossRef Search ADS PubMed  19. Fleming TJ, Fleming ML, Malek TR. Selective expression of Ly-6G on myeloid lineage cells in mouse bone marrow. RB6-8C5 mAb to granulocyte-differentiation antigen (Gr-1) detects members of the Ly-6 family. J Immunol . 1993; 151: 2399– 2408. Google Scholar PubMed  20. Serbina NV, Pamer EG. Monocyte emigration from bone marrow during bacterial infection requires signals mediated by chemokine receptor CCR2. Nat Immunol . 2006; 7: 311– 317. doi: 10.1038/ni1309 Google Scholar CrossRef Search ADS PubMed  21. Lee PY, Wang JX, Parisini E, Dascher CC, Nigrovic PA. Ly6 family proteins in neutrophil biology. J Leukoc Biol . 2013; 94: 585– 594. doi: 10.1189/jlb.0113014 Google Scholar CrossRef Search ADS PubMed  22. Lagasse E, Weissman IL. Flow cytometric identification of murine neutrophils and monocytes. J Immunol Methods . 1996; 197: 139– 150. doi:10.1016/0022-1759(96)00138-X Google Scholar CrossRef Search ADS PubMed  23. Rigamonti E, Touvier T, Clementi E, Manfredi AA, Brunelli S, Rovere-Querini P. Requirement of inducible nitric oxide synthase for skeletal muscle regeneration after acute damage. J Immunol . 2013; 190: 1767– 1777. doi: 10.4049/jimmunol.1202903 Google Scholar CrossRef Search ADS PubMed  24. Schmittgen TD, Livak KJ. Analyzing real-time PCR data by the comparative C(T) method. Nat Protoc . 2008; 3: 1101– 1108. doi:10.1038/nprot.2008.73 Google Scholar CrossRef Search ADS PubMed  25. Manfredi TG, Fielding RA, O’Reilly KP, Meredith CN, Lee HY, Evans WJ. Plasma creatine kinase activity and exercise-induced muscle damage in older men. Med Sci Sports Exerc . 1991; 23: 1028– 1034. doi:10.1249/00005768-199109000-00006 Google Scholar CrossRef Search ADS PubMed  26. Zerba E, Komorowski TE, Faulkner JA. Free radical injury to skeletal muscles of young, adult, and old mice. Am J Physiol . 1990; 258: C429– C435. doi: 10.1152/ajpcell.1990.258.3.C429 Google Scholar CrossRef Search ADS PubMed  27. Roche JA, Tulapurkar ME, Mueller ALet al.   Myofiber damage precedes macrophage infiltration after in vivo injury in dysferlin-deficient A/J mouse skeletal muscle. Am J Pathol . 2015; 185: 1686– 1698. doi: 10.1016/j.ajpath.2015.02.020 Google Scholar CrossRef Search ADS PubMed  28. Lockhart NC, Brooks SV. Protection from contraction-induced injury provided to skeletal muscles of young and old mice by passive stretch is not due to a decrease in initial mechanical damage. J Gerontol A Biol Sci Med Sci . 2006; 61: 527– 533. doi: 10.1093/gerona/61.6.527 Google Scholar CrossRef Search ADS PubMed  29. Li Y, Lee Yi, Thompson WJ. Changes in aging mouse neuromuscular junctions are explained by degeneration and regeneration of muscle fiber segments at the synapse. J Neurosci . 2011; 31: 14910– 14919. doi: 10.1523/JNEUROSCI.3590-11.2011 Google Scholar CrossRef Search ADS PubMed  30. Valdez G, Tapia JC, Kang Het al.   Attenuation of age-related changes in mouse neuromuscular synapses by caloric restriction and exercise. Proc Natl Acad Sci USA . 2010; 107: 14863– 14868. doi: 10.1073/pnas.1002220107 Google Scholar CrossRef Search ADS PubMed  31. Greiwe JS, Cheng B, Rubin DC, Yarasheski KE, Semenkovich CF. Resistance exercise decreases skeletal muscle tumor necrosis factor alpha in frail elderly humans. FASEB J . 2001; 15: 475– 482. doi: 10.1096/fj.00-0274com Google Scholar CrossRef Search ADS PubMed  32. Jozsi AC, Dupont-Versteegden EE, Taylor-Jones JMet al.   Molecular characteristics of aged muscle reflect an altered ability to respond to exercise. Int J Sport Nutr Exerc Metab . 2001; 11( suppl): S9– S15. doi:10.1123/ ijsnem.11.s1.s9 Google Scholar CrossRef Search ADS PubMed  33. Przybyla B, Gurley C, Harvey JFet al.   Aging alters macrophage properties in human skeletal muscle both at rest and in response to acute resistance exercise. Exp Gerontol . 2006; 41: 320– 327. doi: 10.1016/j.exger.2005.12.007 Google Scholar CrossRef Search ADS PubMed  34. Costamagna D, Costelli P, Sampaolesi M, Penna F. Role of inflammation in muscle homeostasis and myogenesis. Mediators Inflamm . 2015; 2015: 805172. doi: 10.1155/2015/805172 Google Scholar CrossRef Search ADS PubMed  35. Degens H. The role of systemic inflammation in age-related muscle weakness and wasting. Scand J Med Sci Sports . 2010; 20: 28– 38. doi: 10.1111/j.1600-0838.2009.01018.x Google Scholar CrossRef Search ADS PubMed  36. Arnold L, Henry A, Poron Fet al.   Inflammatory monocytes recruited after skeletal muscle injury switch into anti-inflammatory macrophages to support myogenesis. J Exp Med . 2007; 204: 1057– 1069. doi: 10.1084/jem.20070075 Google Scholar CrossRef Search ADS PubMed  37. Massimino ML, Rapizzi E, Cantini Met al.   ED2+ macrophages increase selectively myoblast proliferation in muscle cultures. Biochem Biophys Res Commun . 1997; 235: 754– 759. doi: 10.1006/bbrc.1997.6823 Google Scholar CrossRef Search ADS PubMed  38. Carlson BM, Faulkner JA. Muscle transplantation between young and old rats: age of host determines recovery. Am J Physiol . 1989; 256: C1262– C1266. doi: 10.1152/ajpcell.1989.256.6.C1262 Google Scholar CrossRef Search ADS PubMed  39. Conboy IM, Conboy MJ, Smythe GM, Rando TA. Notch-mediated restoration of regenerative potential to aged muscle. Science . 2003; 302: 1575– 1577. doi: 10.1126/science.1087573 Google Scholar CrossRef Search ADS PubMed  40. McBride TA, Gorin FA, Carlsen RC. Prolonged recovery and reduced adaptation in aged rat muscle following eccentric exercise. Mech Ageing Dev . 1995; 83: 185– 200. doi: 10.1016/0047-6374(95)01629-E Google Scholar CrossRef Search ADS PubMed  41. Paliwal P, Pishesha N, Wijaya D, Conboy IM. Age dependent increase in the levels of osteopontin inhibits skeletal muscle regeneration. Aging (Albany NY) . 2012; 4: 553– 566. doi: 10.18632/aging.100477 Google Scholar CrossRef Search ADS PubMed  42. Ullman M, Ullman A, Sommerland H, Skottner A, Oldfors A. Effects of growth hormone on muscle regeneration and IGF-I concentration in old rats. Acta Physiol Scand . 1990; 140: 521– 525. doi: 10.1111/j.1748-1716.1990.tb09029.x Google Scholar CrossRef Search ADS PubMed  43. van der Poel C, Gosselin LE, Schertzer JDet al.   Ageing prolongs inflammatory marker expression in regenerating rat skeletal muscles after injury. J Inflamm (Lond) . 2011; 8: 41. doi: 10.1186/1476-9255-8-41 Google Scholar CrossRef Search ADS PubMed  44. Wang Y, Wehling-Henricks M, Samengo G, Tidball JG. Increases of M2a macrophages and fibrosis in aging muscle are influenced by bone marrow aging and negatively regulated by muscle-derived nitric oxide. Aging Cell . 2015; 14: 678– 688. doi: 10.1111/acel.12350 Google Scholar CrossRef Search ADS PubMed  45. Hammers DW, Merritt EK, Matheny RWJret al.   Functional deficits and insulin-like growth factor-I gene expression following tourniquet-induced injury of skeletal muscle in young and old rats. J Appl Physiol (1985) . 2008; 105: 1274– 1281. doi: 10.1152/japplphysiol.90418.2008 Google Scholar CrossRef Search ADS PubMed  46. Hamada K, Vannier E, Sacheck JM, Witsell AL, Roubenoff R. Senescence of human skeletal muscle impairs the local inflammatory cytokine response to acute eccentric exercise. FASEB J . 2005; 19: 264– 266. doi: 10.1096/fj.03-1286fje Google Scholar CrossRef Search ADS PubMed  47. Peake J, Della Gatta P, Cameron-Smith D. Aging and its effects on inflammation in skeletal muscle at rest and following exercise-induced muscle injury. Am J Physiol Regul Integr Comp Physiol . 2010; 298: R1485– R1495. doi: 10.1152/ajpregu.00467.2009 Google Scholar CrossRef Search ADS PubMed  48. Brigitte M, Schilte C, Plonquet Aet al.   Muscle resident macrophages control the immune cell reaction in a mouse model of notexin-induced myoinjury. Arthritis Rheum . 2010; 62: 268– 279. doi: 10.1002/art.27183 Google Scholar CrossRef Search ADS PubMed  49. Langen RC, Van Der Velden JL, Schols AM, Kelders MC, Wouters EF, Janssen-Heininger YM. Tumor necrosis factor-alpha inhibits myogenic differentiation through MyoD protein destabilization. FASEB J . 2004; 18: 227– 237. doi: 10.1096/fj.03-0251com Google Scholar CrossRef Search ADS PubMed  50. Nguyen HX, Tidball JG. Expression of a muscle-specific, nitric oxide synthase transgene prevents muscle membrane injury and reduces muscle inflammation during modified muscle use in mice. J Physiol . 2003; 550: 347– 356. doi: 10.1113/jphysiol.2003.040907 Google Scholar CrossRef Search ADS PubMed  51. Villalta SA, Nguyen HX, Deng B, Gotoh T, Tidball JG. Shifts in macrophage phenotypes and macrophage competition for arginine metabolism affect the severity of muscle pathology in muscular dystrophy. Hum Mol Genet . 2009; 18: 482– 496. doi: 10.1093/hmg/ddn376 Google Scholar CrossRef Search ADS PubMed  52. Bosurgi L, Corna G, Vezzoli Met al.   Transplanted mesoangioblasts require macrophage IL-10 for survival in a mouse model of muscle injury. J Immunol . 2012; 188: 6267– 6277. doi: 10.4049/jimmunol.1102680 Google Scholar CrossRef Search ADS PubMed  © The Author(s) 2018. Published by Oxford University Press on behalf of The Gerontological Society of America. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png The Journals of Gerontology Series A: Biomedical Sciences and Medical Sciences Oxford University Press

Myeloid Cell Responses to Contraction-induced Injury Differ in Muscles of Young and Old Mice

Loading next page...
 
/lp/ou_press/myeloid-cell-responses-to-contraction-induced-injury-differ-in-muscles-Bs2a3smP7u
Publisher
Oxford University Press
Copyright
© The Author(s) 2018. Published by Oxford University Press on behalf of The Gerontological Society of America. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com.
ISSN
1079-5006
eISSN
1758-535X
D.O.I.
10.1093/gerona/gly086
Publisher site
See Article on Publisher Site

Abstract

Abstract Myeloid cells play a critical role in regulating muscle degeneration and regeneration. Thus, alterations with aging in the myeloid cell response to muscle damage may affect the progression of the injury in old animals. We hypothesized that neutrophil levels remain elevated and that macrophage accumulation is reduced or delayed in injured muscles of old compared with young animals. Muscles of young and old mice were injured with lengthening contractions and analyzed 2 or 5 days later. Regardless of age, neutrophil (Gr-1+) and macrophage (CD68+) content increased dramatically by Day 2. Between 2 and 5 days, macrophages increased further, whereas neutrophils declined to a level that in old muscles was not different from uninjured controls. M2 macrophages (CD163+) also increased between 2 and 5 days, reaching higher levels in muscles of old mice than in young mice. Although no evidence of persisting neutrophils or reduced M2 accumulation in old muscle was found, total macrophage accumulation was lower in old mice. Furthermore, messenger RNA levels showed age-related changes in macrophage-associated genes that may indicate alterations in myeloid cell function. Overall, differences between muscles of old and young mice in the inflammatory response through the early stages of injury may contribute to defects in muscle regeneration. Skeletal muscle, Macrophages, Neutrophils, Damage, Force deficit Skeletal muscle injuries are common and have a variety of causes, including trauma, ischemia/reperfusion, toxin exposure, and unloading/reloading. Muscles can also be injured by their own contractions, especially during lengthening contractions (LCs) when muscles are stretched while activated. Regardless of the cause, the initial insult is followed by degeneration of damaged muscle fibers and subsequent regeneration and repair. Delayed and/or incomplete regeneration of muscles of old animals has been widely reported. Specifically, muscles of old animals have fewer and smaller regenerating fibers (1,2), more fibrosis (3,4), and a greater deposition of fat (4) compared with young animals following injury. Failed restoration of normal muscle structure is accompanied by persistent deficits in force generation (5,6). Although aging-associated impairments in muscle repair are well established, the underlying mechanisms are incompletely understood. A growing body of evidence supports an important role for myeloid cells in regulating the progression of muscle degeneration and regeneration following injury. Neutrophils are the first to accumulate in injured muscle, arriving within hours of the initial insult to facilitate the removal of damaged or necrotic tissue. Neutrophils can also exacerbate damage to muscle fibers through mechanisms involving reactive oxygen species (7), and preventing neutrophil infiltration to muscle after injury has been reported to reduce force deficits and histological damage to muscle fibers (8,9). Macrophage levels increase later and contribute to muscle repair by mechanisms that depend on macrophage phenotype (10,11). Macrophages are generally defined as proinflammatory (M1) and anti-inflammatory (M2), although these classifications are overly simplistic, especially for cells in vivo. In general, M1 macrophages contribute to recovery from injury by removing damaged tissue and then decline in number as M2 macrophages increase coincident with the regenerative stage of muscle repair (10,11). The accumulating evidence that myeloid cells play a critical role in regulating muscle degeneration and regeneration implies that an inappropriate myeloid cell response could contribute to impairments in muscle repair following injury. Thus, based on known muscle regeneration defects observed in old animals, we asked whether there are differences between young and old mice in the timing, magnitude, and/or characteristics of the myeloid cell response to muscle injury. We specifically hypothesized that neutrophils would persist in injured muscles of old compared with young animals and that the accumulation of macrophages, especially M2 macrophages would be delayed or reduced in old muscles. To test our hypothesis, we injured muscles of young and old mice with LCs in situ and analyzed the muscles 2 or 5 days later, using histological, immunohistochemical, and functional measures, as well as protein and mRNA analyses. Method Animals Young (3–5 months) and old (25–27 months) male C57BL6 mice were maintained in a specific-pathogen-free facility at the University of Michigan (UM) until they were used for experimentation. Between procedures, mice were housed in a separate specific-pathogen-free return room. Animal procedures were approved by the UM Institutional Animal Care and Use Committee. In Situ Evaluation of Contractile Properties Procedures for in situ evaluation of muscle contractile properties were based on the previous studies (12). Mice were anesthetized with 3% isoflurane in oxygen. Anesthesia was maintained with 2% isoflurane in oxygen and confirmed by lack of response to tactile stimuli. Ophthalmic ointment was applied to prevent corneal drying. Hind limb fur was removed, and the skin was disinfected with chlorhexidine. A small incision was made at the ankle to expose the distal tendon of the extensor digitorum longus (EDL) muscle, and 6-0 braided silk was tied around the tendon. Another small incision just distal to the knee exposed the fibular nerve. The mouse was placed on a 37°C platform, and the hind limb was immobilized by pinching the knee and foot with small clamps secured to the platform. Using the tails of the silk suture, the intact tendon was tied to the lever arm of a servomotor (300C-LR-FP, Aurora Scientific) that controlled muscle length and measured force. Custom-designed software controlled stimulus pulses and motor movement and collected force data. The small area of exposed tendon was kept moist by frequent administration of warmed sterile saline. The EDL muscle was activated with platinum electrodes placed under the fibular nerve. Stimulus pulses of 0.2-ms duration were applied using an Aurora Scientific 701C high-power current stimulator, and current and muscle length were adjusted to elicit maximum twitch force. Tetanic contractions were elicited with 200-ms trains of pulses. The frequency of pulses was increased during successive contractions until maximum isometric force (Po) was achieved, typically at 200 Hz. Optimal muscle length (Lo) was measured with calipers, and optimal fiber length (Lf) was determined by multiplying Lo by the previously determined Lf-to-Lo ratio of 0.45 (13). In Situ LC Protocol To induce muscle injury, we used the physiologically relevant approach of exposure to LC. Our group has previously characterized many aspects of the degeneration and regeneration responses following LC-induced injury and demonstrated the ability to administer well-controlled contraction protocols resulting in highly reproducible injury responses (6,8,12,14,15). In the present study, the EDL muscle was exposed to a protocol of 75 LCs spaced 4 seconds apart for a total duration of 5 minutes. Each contraction was 300 ms in duration. A stretch of 20% strain relative to Lf was initiated 100 ms after the onset of stimulation from near maximum isometric force. Muscles were lengthened at 1 Lf/s such that the peak of the stretch coincided with the end of stimulation. After the LC protocol, a final measurement of maximum isometric force was made. Force decreased 49% ± 9% (SD) and 58% ± 12% (SD) for muscles of young and old mice, respectively, reflecting both injury and fatigue. After completion of the in situ experiments, the small incisions at the ankle and knee were closed with 7-0 sterile monofilament nylon suture and bathed with povidone-iodine solution, and mice were monitored until they recovered from anesthesia. In Vitro Evaluation of Contractile Properties Two or five days after LCs, injured and contralateral control muscles were evaluated for Po in vitro as described (13). Mice were anesthetized with an intraperitoneal injection of Avertin (tribromoethanol, 250 mg/kg), and EDL muscles were carefully isolated and removed. Mice were then euthanized with an overdose of Avertin followed by induction of a bilateral pneumothorax. 5-0 silk suture was tied to the proximal and distal tendons of the muscle, which was then placed in a chamber containing Krebs Mammalian Ringer solution composed of (in millimolar) 137 NaCl, 5 KCl, 2 CaCl2·2H2O, 1 MgSO4·7H2O, 1 NaH2PO4, 24 NaHCO3, 11 glucose, and 0.03 tubocurarine chloride. The bath was held at 25°C and bubbled with 95% O2–5% CO2 to maintain a pH of 7.4. The tendons were tied to a force transducer (BG-50, Kulite Semiconductor Products) and a fixed post. Field stimulation (Aurora 701C stimulator) was applied via parallel plate electrodes. Contractile properties were determined as described above except that tetanic contractions were elicited with 300-ms trains of pulses and Po occurred at ~150 Hz. The force deficit induced by the LCs was calculated as the difference between Po measured for the muscle exposed to LCs and the contralateral uninjured control muscle expressed as a percentage of the contralateral control Po. Following force measurements, muscles were trimmed of tendons, weighed, and cut in half. One half was immersed in tissue freezing medium and frozen in isopentane cooled by liquid nitrogen; the other half was submerged in an RNA stabilization reagent (RNAlater, QIAGEN). Histology, Immunohistochemistry, and Immunofluorescence We have previously shown that muscle function, histology, and inflammatory cell levels were not influenced by the surgical procedures associated with the LC protocol (12,16). We have now performed sham operations on old mice and confirmed that no changes were observed in neutrophil or macrophage levels compared with undisturbed control muscles in the days following the surgery. Thus, contralateral muscles from mice in which the ipsilateral muscle had been exposed to LCs served as controls for histology. Cryosections of 10 µm were fixed in cold acetone and stained with hematoxylin (Ricca Chemical Company) and eosin Y (EMD Millipore; Supplementary Figure S1A). In single sections, injured fibers were counted as those with pale or variable staining, swollen appearance, or obvious infiltration by inflammatory cells (8). Basophilic fibers with central nuclei were identified as regenerating (17). Image J was used to count fibers and measure cross-sectional areas of individual fibers and entire sections. Total section area was multiplied by section thickness to calculate volume. Muscle sections were interrogated for neutrophils and macrophages by immunohistochemistry. Others report that mouse CD68 (rat ED1+) labels early-invading macrophages that accumulate in injured skeletal muscle and invade muscle fibers (10,11), whereas mouse CD163 (rat ED2+) labels macrophages that appear later and are associated with repair (10,11). Accordingly, we chose CD68 and CD163 as markers of M1 and M2 macrophages, respectively. To examine the specificity and selectivity of these markers, we used immunofluorescence to label muscles for CD68 and CD163 or CD163 and CD301. CD301 is reported to be a marker of M2 macrophages in mice (18). Sections were fixed in cold acetone, air-dried, and exposed to 1% bovine serum albumin (Jackson ImmunoResearch Laboratories) in phosphate-buffered saline (PBS) to block nonspecific binding. Macrophages were detected with a rat anti-mouse CD68 (FA-11, 1:1,000; AbD Serotec), rabbit anti-mouse CD163 (M-96, 1:200; Santa Cruz Biotechnology), or goat anti-mouse CD301 IgG (1:400; R&D Systems) antibodies. Secondary antibodies were Alexa Fluor 594 goat anti-rat IgG, Alexa Fluor 594 bovine anti-goat IgG, and Alexa Fluor 488 donkey anti-rabbit IgG, all from Jackson ImmunoResearch Laboratories. Last, sections were exposed to 4′,6-diamidino-2-phenylindole (D9564; Sigma–Aldrich, St. Louis, MO) and treated with ProLong Gold (P36930; Life Technologies). These analyses showed that CD163+ cells were also CD68+ (Supplementary Figure S2), indicating that CD68 and CD163 are not distinguishing markers of macrophage phenotype in injured mouse EDL muscle. Rather, CD163 labels a subset of the macrophages also labeled by CD68. CD163+ cells were also positive for CD301 (Supplementary Figure S3), supporting the identification of these cells as M2 macrophages (18). Therefore, CD68 was considered to be a general macrophage marker and CD163 a marker of M2 macrophages. To quantify neutrophils and macrophages, sections were fixed in cold acetone, air-dried, and exposed to BLOXALL solution to block endogenous peroxidase activity followed by 10% normal rabbit or goat serum in PBS (Fisher Scientific) to block nonspecific binding. Neutrophils were detected with a rat anti-mouse Gr-1 antibody (RB6-8C5, 1:50 dilution; BD Pharmingen) diluted in PBS containing 10% rabbit serum, and macrophages were detected with a rat anti-mouse CD68 antibody (FA-11, 1:1,000; AbD Serotec) diluted in PBS containing 10% rabbit serum or with a rabbit anti-mouse CD163 antibody (M-96, 1:1,000; Santa Cruz Biotechnology) diluted in PBS containing 10% goat serum. After incubation in primary antibodies, sections were exposed to biotinylated mouse adsorbed rabbit anti-rat IgG (Gr-1 and CD68) or biotinylated goat anti-rabbit IgG (CD163) in PBS containing 10% rabbit or goat serum, respectively. Sections were treated with VECTASTAIN Elite ABC reagent containing peroxidase followed by peroxidase substrate 3, 3′-diaminobenzidine (ImmPACT DAB). All reagents were from Vector Laboratories unless stated otherwise. Pilot studies showed no regional differences (proximal, central, or distal portion of the muscle) in inflammatory cell infiltration. Therefore, Image J was used to count neutrophils and macrophages in single sections from the belly of each muscle. RB6-8C5 (Gr-1) reacts with both Ly6G and Ly6C (19), and neutrophils display abundant expression of both, with Ly6G levels in peripheral neutrophils directly correlated with the cell’s level of differentiation and maturation (20). Despite the widespread usage of Gr-1 as a neutrophil marker, Ly6C is also expressed at lower levels by a number of other cell types including monocytes, eosinophils, dendritic cells, a wide range of endothelial cells, and subsets of T cells (21,22). Thus, dark brown cells indicating high Gr-1 levels were counted as neutrophils (18), whereas the occasional light brown cell of similar size was not counted (Supplementary Figure S1B). Both light and dark brown cells were counted as macrophages in sections treated with reagents for CD68 (Supplementary Figure S1C) and CD163 (Supplementary Figure S1D) detection. In all cases, cells at the extreme periphery of the section were not counted. Western Blots For analysis of CD68 and CD163 levels, EDL muscles were placed in bead homogenization tubes with RIPA buffer containing 50 mM Tris (pH 7.4), 150 mM NaCl, and protease inhibitors (Thermofisher Scientific, Waltham, MA), whereas for analysis of Ly6G/Ly6C, muscle tissues were homogenized in 20 mM Tris buffer with 0.05% Triton X and the protease inhibitors. All samples were homogenized using a Bead Mill 4 homogenizer (Fisher Scientific, Hampton, NH) for two cycles at 4 m/s for 45 seconds with a 1-minute rest period. After homogenization, samples were centrifuged at 13,000g for 15 minutes and transferred to 1.5-mL microcentrifuge tubes. Protein content was determined using the bicinchoninic acid method (Sigma–Aldrich). For CD68 and CD163, muscle homogenates (20 µg) were fractionated in 4%–20% Mini-PROTEAN TGX precast gels, and gels were transferred to polyvinylidene difluoride membranes. Membranes were stained with Ponceau S before blotting to verify equal loading of the gels. Membranes were blocked in 5% milk in Tris-buffered saline with 0.1% Tween-20 (TBST), for 1–2 hours. Antibodies for CD68 (ab201340; Abcam, Cambridge, UK) or CD163 (ab182422; Abcam, Cambridge, UK) were diluted 1:1,000 in 5% milk in TBST and incubated at 4°C overnight. For Ly6, muscle homogenates (20 µg) were fractionated in 10% native gels and placed in Tris–glycine buffer (pH 8.9) surrounded by Tris–glycine buffer (pH 8.3). Gels were transferred to polyvinylidene difluoride membranes in Tris/Glycine buffer 0.01% sodium dodecyl sulfate. In native conditions, the primary antibody for Ly6G/Ly6C (RB6-8C5; BD Pharmingen, San Jose, CA) was diluted 1:750 in 5% milk in TBST and incubated at 4°C overnight. Anti-rabbit and anti-mouse monoclonal secondary antibodies (Cell Signaling Technologies, Danvers, MA) were diluted 1:2,000 in 5% milk, in TBST, and then incubated at room temperature for 1 hour. Enhanced chemiluminescence was performed using Chemidoc XRS imaging system (Bio-Rad, Hercules, CA) to visualize antibody–antigen interaction. Blotting images were quantified by densitometry using myImageAnalysis software (Thermofisher Scientific). The Ponceau-stained membranes were digitally scanned, and the 45-kDa actin bands were quantified by densitometry and used as a protein loading correction factor for each lane. An internal control was loaded on all gels to make quantitative comparisons between gels. Gene Expression Muscle samples were homogenized in QIAzol lysis reagent (QIAGEN). RNA was isolated using a miRNeasy Mini Kit and treated with DNase I (QIAGEN). RNA concentration was measured with a NanoDrop 2000 Spectrophotometer (Thermo Scientific). RNA (2 µg) was reverse transcribed using a QuantiTect Reverse Transcription Kit (QIAGEN), and cDNA was diluted by fivefold and amplified in a CFX96 real time thermal cycler (Bio-Rad) using a QuantiTect SYBR Green I PCR Kit (QIAGEN) and primers specific for inducible nitric oxide synthase (iNOS) (23), tumor necrosis factor alpha (TNFα) (23), arginase-1 (Arg1) (10), interleukin-10 (IL-10) (10), and β2-microglobulin (15). Primers were purchased from Integrated DNA Technologies and are listed in Supplementary Table S1. mRNA from target genes was undetectable in control muscles, so we completed our analysis using only injured muscles. We were unable, despite multiple trials, to identify a housekeeping gene that remained stable both between age groups and over time following LCs. This is perhaps not surprising, due to the ongoing breakdown of the tissue and the ongoing accumulation in the muscle of different cell types. Because mRNA levels of β2-microglobulin were not different between age groups for any of the experimental conditions and our primary interest was in identifying age-associated differences, we normalized mRNA transcripts to β2-microglobulin and compared young and old samples at each time point. mRNA levels were analyzed by the comparative CT method (24). Blood Analysis Blood (50–100 µL) was collected from a separate group of undisturbed control mice via a scalpel nick of the lateral tail vein. White blood cell counts were determined in whole blood using a cell counter (Hemavet 950FS, Drew Scientific). Data Analysis Data are reported as means ± 1 SEM. Measures of injury and levels of myeloid cells and protein were compared between age groups and across time using a two-factor analysis of variance (ANOVA). In cases when the ANOVA showed significant effects, the Holm-Sidak post hoc test was used to identify individual differences. When comparisons were between only two groups, for example, regenerating fibers in muscles of young and old mice, Student’s t tests were used. In all cases, significance was set a priori at p < .05, and individual p values are included throughout the text and in figure legends and tables. Results Muscle masses were not different between young and old mice, but muscles from old mice were weaker by all measures (Table 1). Although muscles in old animals are reported to be more susceptible to contraction-induced injury (25,26), a common approach for comparing the progression of the injury process between young and old animals is to induce an injury of similar magnitude in both age groups (5,6,12,14). In the present study, the contraction protocol resulted in a similar level of injury in muscles of young and old mice as indicated by no statistically significant differences in the measures of injury after 2 days. Force deficits of ~55% were not different (p = .183) for muscle of young and old mice (Figure 1A), and there was also no difference between the age groups (p = .336) in the number of damaged fibers appearing in muscle cross sections at 2 days (Figure 1B). For young mice, both the force deficit (p = .007) and the number of damaged fibers (p < .001) increased between 2 and 5 days, and the total number of fibers in a cross section decreased (p < .001). The number of damaged fibers increased (p = .007) between 2 and 5 days for old mice, as well, but the number of fibers showing histological evidence of damage was ~35% lower (p = .005) for muscles of old mice compared with young mice at 5 days (Figure 1B) and the total number of fibers remaining in cross section 5 days after LCs was ~15% greater (p = .039) in muscles of old mice compared with young mice (Figure 1C). Table 1. Characteristics of Young and Old Groups Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Note: CSA = cross-sectional area; H&E = hematoxylin and eosin. Forces are maximum forces generated during isometric tetanic contractions. [Two samples are missing from young group for CSA from H&E stained sections and for total number of fibers (ie, n = 16).] aData taken from contralateral muscles. *Significant difference from young group (p < .05). p Values are provided from t tests for each variable. View Large Table 1. Characteristics of Young and Old Groups Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Group  n  Body Weight (g)  Preinjury Force In Situ (mN)  Absolute Force In Vitro (mN)a  Specific Force In Vitro (mN/mm2)a  Muscle Mass (mg)a  CSA From H&E Stained Sections (mm2)a  Total Number of Fibersa  Young  18  32.0 ± 0.8  407 ± 8  473 ± 8  250 ± 4  12.0 ± 0.2  1.9 ± 0.1  951 ± 26  Old  19  32.7 ± 0.6  343 ± 9*  436 ± 13*  232 ± 6*  11.9 ± 0.3  1.6 ± 0.1*  936 ± 23  p Value    .470  <.001  .018  .026  .765  .009  .875  Note: CSA = cross-sectional area; H&E = hematoxylin and eosin. Forces are maximum forces generated during isometric tetanic contractions. [Two samples are missing from young group for CSA from H&E stained sections and for total number of fibers (ie, n = 16).] aData taken from contralateral muscles. *Significant difference from young group (p < .05). p Values are provided from t tests for each variable. View Large Figure 1. View largeDownload slide Lengthening contractions damage muscles of young and old mice. The severity of injury was assessed 2 and 5 d after LCs by (A) the deficit in isometric force generating capacity expressed as a percentage of the contralateral control muscle, (B) the percentage of fibers in a single cross section showing histological evidence of damage, and (C) the total number of fibers appearing in a cross section. Data are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for force deficit (Time: p = .008; Age: p = .004; Time × Age: p = .231) and number of injured fibers (Time: p < .001; Age: p = .008; Time × Age: p = .151) and significant effects of time with significant interactions for the number of fibers in cross sections with no main effect of age (Time: p < .001; Age: p = .319; Time × Age: p = .045). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 1. View largeDownload slide Lengthening contractions damage muscles of young and old mice. The severity of injury was assessed 2 and 5 d after LCs by (A) the deficit in isometric force generating capacity expressed as a percentage of the contralateral control muscle, (B) the percentage of fibers in a single cross section showing histological evidence of damage, and (C) the total number of fibers appearing in a cross section. Data are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for force deficit (Time: p = .008; Age: p = .004; Time × Age: p = .231) and number of injured fibers (Time: p < .001; Age: p = .008; Time × Age: p = .151) and significant effects of time with significant interactions for the number of fibers in cross sections with no main effect of age (Time: p < .001; Age: p = .319; Time × Age: p = .045). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. ANOVA = analysis of variance; LCs = lengthening contractions. Regenerating fibers were clearly present in muscle cross sections 5 days after LCs (Figure 2A and B), with the number of regenerating fibers in injured muscles from old mice nearly 40% lower (p = .050) than in young mice (Figure 2C). In addition to counting the number of regenerating fibers, previous studies have also used the cross-sectional areas of regenerating fibers to compare groups for the rate and extent of regeneration (2,9). Thus, we also analyzed muscle fiber cross-sectional areas as a measure of regeneration. Regenerating fibers in muscle sections from old mice were 20% smaller (p < .001) than those of young mice at Day 5 (Figure 2D). Collectively, these findings are consistent with widely published reports of delayed or deficient repair in muscles of old animals. Figure 2. View largeDownload slide Regeneration is impaired in old mice. Representative partial sections of muscles stained with hematoxylin and eosin from (A) young and (B) old mice show regenerating fibers. Scale bar = 50 µm and applies to both images. Data are shown for (C) the number of regenerating fibers per muscle cross section and (D) average cross-sectional area of regenerating fibers 5 d after lengthening contractions. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Figure 2. View largeDownload slide Regeneration is impaired in old mice. Representative partial sections of muscles stained with hematoxylin and eosin from (A) young and (B) old mice show regenerating fibers. Scale bar = 50 µm and applies to both images. Data are shown for (C) the number of regenerating fibers per muscle cross section and (D) average cross-sectional area of regenerating fibers 5 d after lengthening contractions. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Muscle degeneration and regeneration were accompanied by accumulation of myeloid cells in injured muscles. Two days after LCs, neutrophil content was dramatically elevated relative to control levels in both young and old mice, as evidenced by both numbers of Gr-1+ cells (p < .001; Figure 3A) and Ly6G/Ly6C levels (Figure 4A), with numbers of neutrophils 35% higher (p = .002) in muscles of old than in young mice at this time point. Consistent with previous reports of rapid clearance of neutrophils from injured muscles (9,27), both age groups showed declines in neutrophil numbers of ~70% (p < .001) between 2 and 5 days. The decrease in neutrophils resulted in levels in old mice by the 5-day time point that were not different (p = .088) from baseline values, although baseline neutrophil numbers in old mice were elevated (p = .024) compared with control muscles of young mice (Figure 3A). High variation in the old muscles for Ly6G/Ly6C levels following injury resulted in only a trend (p = .067) for an effect of age by two-factor ANOVA, but consistent with higher baseline numbers of neutrophils in old muscles compared with young muscles, approximately fourfold higher baseline levels of Ly6G/Ly6C in old control muscles were highly significant (p < .001) by t test. Ly6G/Ly6C levels remained elevated at 5 days for both age groups (Figure 4A), probably reflecting Ly6C expression by other cell types, in particular activated macrophages, which are present in 100-fold greater abundance than neutrophils at 5 days (Figure 3A and B). Figure 3. View largeDownload slide Myeloid cell accumulation is altered in injured muscles from young and old mice. Data are shown for (A) neutrophil (Gr-1+ cells), (B) macrophage (CD68+ cells), and (C) M2 macrophage (CD163+ cells) content expressed as the number of cells per volume of muscle in uninjured control muscles and 2 and 5 d after LCs. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for neutrophil (Time: p < .001; Age: p < .001; Time × Age: p = .413) and M2 macrophage (Time: p = .039; Age: p < .001; Time × Age: p = .217) number and significant effects of time and age with significant interactions for the number of CD68+ macrophages (Time: p = .002; Age: p < .001; Time × Age: p < .001). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 3. View largeDownload slide Myeloid cell accumulation is altered in injured muscles from young and old mice. Data are shown for (A) neutrophil (Gr-1+ cells), (B) macrophage (CD68+ cells), and (C) M2 macrophage (CD163+ cells) content expressed as the number of cells per volume of muscle in uninjured control muscles and 2 and 5 d after LCs. Values are presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for neutrophil (Time: p < .001; Age: p < .001; Time × Age: p = .413) and M2 macrophage (Time: p = .039; Age: p < .001; Time × Age: p = .217) number and significant effects of time and age with significant interactions for the number of CD68+ macrophages (Time: p = .002; Age: p < .001; Time × Age: p < .001). Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 4. View largeDownload slide Protein levels for myeloid cell markers increase following injury and with age. Data are shown for (A) Gr-1 (Ly6G and Ly6C), (B) CD68, and (C) CD163 protein levels in uninjured control muscles and 2 and 5 d after LCs determined by Western blot. Representative blots are shown in the right-hand panels. For analysis, protein levels were normalized to Ponceau S bands, which are also shown. Internal controls, randomly selected aged myotoxin-injected muscles, are included to allow densitometry measures to be compared between different blots. Left panels show average values expressed relative to the level in young control muscles and presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for CD68 levels (Time: p = .001; Age: p < .001; Time × Age: p = .377) and significant effects of time with strong trends for effects of age for Gr-1 (Time: p = .015; Age: p = .067; Time × Age: p = .398) and CD163 (Time: p < .001; Age: p = .078; Time × Age: p = .879) levels. Sample sizes are n = 4 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. Figure 4. View largeDownload slide Protein levels for myeloid cell markers increase following injury and with age. Data are shown for (A) Gr-1 (Ly6G and Ly6C), (B) CD68, and (C) CD163 protein levels in uninjured control muscles and 2 and 5 d after LCs determined by Western blot. Representative blots are shown in the right-hand panels. For analysis, protein levels were normalized to Ponceau S bands, which are also shown. Internal controls, randomly selected aged myotoxin-injected muscles, are included to allow densitometry measures to be compared between different blots. Left panels show average values expressed relative to the level in young control muscles and presented as means ± SEM for muscles of young (black bars) and old (gray bars) mice. Two-factor ANOVA indicated significant effects of time and age without significant interactions for CD68 levels (Time: p = .001; Age: p < .001; Time × Age: p = .377) and significant effects of time with strong trends for effects of age for Gr-1 (Time: p = .015; Age: p = .067; Time × Age: p = .398) and CD163 (Time: p < .001; Age: p = .078; Time × Age: p = .879) levels. Sample sizes are n = 4 per group. Differences (p < .05) between individual groups are indicated by brackets. *Differences within an age group from the control value. †Differences within an age group from the 2-d value. ANOVA = analysis of variance; LCs = lengthening contractions. In contrast to neutrophils, baseline numbers of macrophages (CD68+ cells) were not different (p = .948) in muscles of young and old mice, and both age groups showed similar fivefold increases (p < .001) in CD68+ macrophage content relative to their respective control levels 2 days after LCs (Figure 3B). Between 2 and 5 days, macrophage content increased another threefold to fourfold (p < .001) for both age groups but to a lesser extent in old mice resulting in macrophage numbers in muscles of old mice that were ~25% lower (p < .001) than in young muscles at 5 days (Figure 3B). Although accumulation of macrophages was lower in muscles of old mice at 5 days, CD68 levels were not different (p = .546) in muscles of young and old mice at that time point (Figure 4B), perhaps reflecting a compensatory response to a decrease in function, although this is speculation. The function of CD68 is not clear beyond its role targeting macrophages to tissues. Control levels of M2 macrophages (CD163+ cells) were also not different (p = .666) for muscles of young and old mice and did not increase for either age group (young: control vs. 2 day, p = .629; old: control vs. 2 day, p = .833) until 5 days following LCs (young and old: control vs. 5 day, p < .001) concurrent with the regenerative phase. The increase in M2 macrophages between 2 and 5 days (p < .001) was greater for muscles of old mice resulting in levels of M2 macrophages at 5 days that were ~40% higher (p = .017) in old muscles compared with young muscles (Figure 3C). CD163 protein levels assessed by western blot were generally consistent with the cell numbers determined through immunohistochemistry, increasing fourfold to fivefold (p < .001) for both age groups by Day 5 and showing a trend for increased levels (p = .078) in muscles of old mice compared with young mice (Figure 4C). The effect of differences in myeloid cell content in muscles of young and old mice may either be exacerbated or mitigated by alterations in myeloid cell function. Alternatively, differences in cellular accumulation may represent compensatory responses to poorly functioning myeloid cells. To begin to explore these questions, we examined messenger RNA levels for genes expressed by M1 (iNOS, TNFα) or M2 (Arg1, IL-10) macrophages in injured muscles from young and old mice (Figure 5). In old injured muscles, mRNA levels for iNOS (p = .621) and TNFα (p = .667) were not different from young muscles 2 days after LCs (Figure 5A and C), but were significantly elevated (iNOS; p = .021; TNFα, p = .014) over young levels at 5 days (Figure 5B and D). mRNA levels for Arg1 were also not different (p = .439) between young and old muscles 2 days after LCs (Figure 5E). After 5 days, Arg1 mRNA level in old muscles appeared to increase relative to young, but values from old animals were highly variable and the difference was not significant (Figure 5F, p = .130). Two days after LCs, IL-10 mRNA levels were lower (p = .009) in old muscles compared with young muscles (Figure 5G) with no differences (p = .280) between age groups at 5 days (Figure 5H). Figure 5. View largeDownload slide Messenger RNA levels of M1 (iNOS, TNFα) and M2 (Arg1, IL-10) macrophage genes are altered with age. Shown are messenger RNA levels for iNOS (A, B), TNFα (C, D), Arg1 (E, F), and IL-10 (G, H) normalized to β2-microglobulin mRNA 2 d (A, C, E, G) and 5 d (B, D, F, H) after lengthening contractions for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Arg1 = arginase-1; IL-10 = interleukin-10; iNOS = inducible nitric oxide synthase; LC = lengthening contraction; TNFα = tumor necrosis factor alpha. Figure 5. View largeDownload slide Messenger RNA levels of M1 (iNOS, TNFα) and M2 (Arg1, IL-10) macrophage genes are altered with age. Shown are messenger RNA levels for iNOS (A, B), TNFα (C, D), Arg1 (E, F), and IL-10 (G, H) normalized to β2-microglobulin mRNA 2 d (A, C, E, G) and 5 d (B, D, F, H) after lengthening contractions for muscles of young (black bars) and old (gray bars) mice. Sample sizes are n = 8–10 per group. Differences (p < .05) between individual groups are indicated by brackets. Arg1 = arginase-1; IL-10 = interleukin-10; iNOS = inducible nitric oxide synthase; LC = lengthening contraction; TNFα = tumor necrosis factor alpha. Finally, we questioned whether neutrophil and macrophage content in injured muscles from young and old mice reflected the baseline circulating levels of neutrophils and monocytes with age. In mice that were not subjected to LCs, circulating levels of neutrophils and monocytes were significantly greater (neutrophils, p = .008; monocytes, p = .016) in old animals relative to young animals (Figure 6). Figure 6. View largeDownload slide Circulating levels of neutrophils and monocytes are elevated in old mice. Mean (A) neutrophil and (B) monocyte levels expressed as thousand cells per microliter blood ± SEM are shown for young (black bars) and old (gray bars) mice. Sample sizes are n = 6 per group. Differences (p < .05) between individual groups are indicated by brackets. Figure 6. View largeDownload slide Circulating levels of neutrophils and monocytes are elevated in old mice. Mean (A) neutrophil and (B) monocyte levels expressed as thousand cells per microliter blood ± SEM are shown for young (black bars) and old (gray bars) mice. Sample sizes are n = 6 per group. Differences (p < .05) between individual groups are indicated by brackets. Discussion The goal was to examine the effects of aging on the timing and magnitude of the myeloid cell response to muscle injury. Injury was induced in muscles of young and old mice by exposure to LCs. The amount of injury observed in the present study was comparable qualitatively and quantitatively to previous studies from our laboratory that used the same contraction protocol (28). Moreover, the use of a contraction protocol that resulted in similar magnitudes of injury in muscles of young and old mice 2 days following LCs, as indicated by no differences between the age groups in force deficit or histological evidence of damage, allowed us to evaluate and compare the progression of degeneration and regeneration in the two age groups starting from roughly equivalent initial levels of damage. This is a strategy that has been used previously (5,6,12,14). Overall, our results support alterations in the myeloid cell response to LC-induced injury in old mice relative to young mice. Consistent with previous reports (8,9), we observed a robust increase in neutrophils following LCs in both young and old muscles. We did not, however, observe a persistence of neutrophils in old muscles, as expected. This conclusion is based on our findings that neutrophils declined dramatically between 2 and 5 days in both young and old mice. This interpretation is complicated somewhat by our observation that Gr-1 protein levels did not show a similar decline between 2 and 5 days. The significance of the continued elevation of Ly6G/Ly6C protein levels above control levels is not clear, but it indicates that protein levels are not a good indicator of neutrophil numbers when large numbers of macrophages and/or other cell types that also express Ly6C are also present. Our observation that the neutrophil level remained higher in muscles of old mice compared with young mice 5 days after LCs is not interpreted as a persistence of the neutrophils induced by injury, because the tissue neutrophil level in old mice had returned at this time point to the level observed in control muscles. Although control levels of neutrophils were rapidly reestablished in muscles of old mice following LCs, the elevated baseline neutrophil content in muscles of old mice compared with young mice suggests either underlying injury in old mice (29,30) or the presence of elevated proinflammatory signals in muscles in the absence of injury (31–33). The impact of an age-associated increase in baseline inflammation has been hypothesized to contribute to age-associated muscle wasting through effects on myogenesis as well as the balance between protein synthesis and degradation (34,35) and may also influence repair from damage. A second unexpected finding was that accumulation of M2 macrophages was not impaired in muscles of old mice. We had predicted delays or reductions in old animals in the accumulation of M2 macrophages following LCs based on the role of M2 macrophages to promote muscle regeneration (10,36,37) and the generally accepted view that muscle regeneration is impaired with aging (1,2,4–6,14,38–43). M2 macrophages have been reported to stimulate proliferation and differentiation of muscle precursor cells in vitro (10,36,37). Thus, our observation of fewer and smaller regenerating fibers in injured muscles of old mice despite no impairment in the accumulation of M2 macrophages suggests that macrophage function may be impaired in old muscles. Although we did not observe reduced M2 macrophage accumulation in old muscles, we cannot rule out the possibility that the levels continue to increase beyond 5 days and old mice would ultimately display a defect in M2 macrophage accumulation. However, Wang and colleagues (44) recently reported higher levels of CD163+ M2a macrophages in muscles of old mice compared with young mice, a finding that is potentially consistent with the present study and our own data showing ~20% higher baseline levels of M2 macrophages in old muscles (young: 10,130 ± 348 CD163+ cells per mm3 tissue vs. old: 12,070 ± 421 cells per mm3 tissue), although the difference was not statistically significant (p = .666). The finding of increases in young mice between 2 and 5 days for both force deficit and the number of damaged fibers along with a loss of fibers from cross sections indicates that muscle fiber degeneration continued in young mice throughout the time period studied. In contrast, no change in the force deficit for old mice along with the smaller increase in the percentage of fibers showing histological evidence of damage in muscles of old mice compared with young mice between 2 and 5 days is consistent with a diminished degeneration response for old mice. In addition, the total number of fibers remaining in cross section at 5 days was slightly greater in muscles of old mice compared with young mice, suggesting that fewer muscle fibers had completely degenerated in old mice at this time point. This apparent blunting of degeneration in old mice was associated with a reduction in overall macrophage accumulation, as evidenced by our observation of 25% fewer CD68+ cells in muscles of old mice compared with young mice 5 days following exposure to LCs. Although based on the present data set, we cannot definitively establish whether the fewer macrophages observed in muscles of old mice compared with young mice 5 days following LCs are a cause or consequence of the reduced level of injury observed at that time point, the similarity in force deficits, total fiber number, and number of damaged fibers in muscles of young and old mice at 2 days argues against the possibility that less severe injury was a major driver of lower macrophage accumulation in old muscles. Based on the known phagocytic function of M1 macrophages (11,36), the observation of fewer CD68+ macrophages in muscles of old mice is consistent with the hypothesis that the phagocytic removal of damaged tissue would be less effective in muscles of old mice (36). Higher CD68 levels observed in muscles of old compared with young mice may reflect a compensatory response to an age-related decrease in cell number, but this is entirely speculative at this stage and future studies are necessary to examine the phagocytic activity of macrophages from old muscles. Previous investigations comparing leukocyte accumulation in muscles from young and old animals in response to injury have generated conflicting conclusions. For example, muscles from old rats displayed elevated CD68 protein content compared with muscles of young rats 3 days after a contusion injury (3), and using immunohistochemistry, Koh and colleagues (12) reported almost twofold higher numbers of neutrophils (Ly6G+ cells) and 50% more macrophages (F4/80+ cells) in muscles of old mice compared with young mice 3 days after LCs. Similarly, following ischemia/reperfusion injury, plantaris muscles of old rats showed greater leukocyte infiltration than that observed in young rats, but specific cell types were not identified (45). Finally, prolonged expression (mRNA) of the leukocyte marker CD18 was reported in muscles of old rats after myotoxin injection (43), although this marker cannot distinguish between persistence of neutrophils and an age-related increase in macrophage levels. Contrary to these prior reports that generally support an age-related increase in myeloid cell content after injury, Paliwal and colleagues (41) found no difference in the numbers of CD11b+ cells, as assessed by flow cytometry, in muscles of young and old mice 3 and 5 days after cardiotoxin injection, and Hamada and colleagues (46) suggested delayed or reduced leukocyte accumulation with advanced age based on their observation of a blunted increase in CD18 mRNA levels in vastus lateralis muscles of old mice relative to young mice 3 days after downhill running. Finally, Sadeh (2) reported delayed recruitment of macrophages in muscles of old rats compared with young rats injured with myotoxin, but macrophage numbers were estimated based on counting nuclei in muscle sections stained with H&E, which would also include other cell types. The bases for these disparate findings are not known, but they highlight the importance of quantifying the extent of damage induced and combining approaches for assessing leukocyte numbers and function as we have begun to do here. Full interpretation of our findings must also consider the levels of myeloid cells within tissues before the injury. Indeed, we observed substantially more neutrophils in uninjured control muscles of old mice relative to young mice. In contrast, the greater LC-induced accumulation of M2 macrophages in muscles of old mice occurred with no significant difference between muscles of young and old mice in M2 macrophage levels either at baseline or 2 days following LCs. Thus, the higher levels of M2 macrophages are not merely a reflection of differing control conditions. Numerous factors could play a role in differences in inflammatory cell numbers in old mice versus young mice including differences in chemoattractant release by the injured muscle, effectiveness of the homing mechanisms for leukocytes to injured sites, or alterations in the life span of inflammatory cells in the tissue. Our observation of elevated CD68 expression in muscles of old mice compared with young mice may represent a compensatory response to impairments in the ability of macrophages to effectively target the injured tissue. The precise signals that trigger inflammatory cell accumulation in muscle are not known, but TNFα, monocyte chemotactic protein-1 (MCP-1, CCL2), IL-8, granulocyte-macrophage colony stimulating factor, and cyclooxygenase (COX)-2 have all been reported to recruit neutrophils and macrophages to injured muscle (47). In addition, resident myeloid cells recruit other inflammatory cells to injured muscle (48). Additional experiments exploring age-associated changes in the levels of chemoattractant signals are warranted. Finally, consistent with our observation of elevated levels of circulating leukocytes in old mice compared with young mice, neutrophil and macrophage content in tissues may also be influenced by a larger pool of cells available to respond to the recruitment signals. Consistent with previous reports of elevations in iNOS, TNFα, Arg1, and IL-10 in injured muscles (10,23) and in myeloid cells isolated from injured muscles (23,36), we also detected iNOS, TNFα, Arg1, and IL-10 mRNA in injured muscles at both 2 and 5 days after LCs. Our finding that at 5 days, mRNA levels of iNOS and TNFα were elevated in muscles from old mice relative to young mice suggests that iNOS and TNFα were produced in excess by macrophages in the old mice. In support of this conclusion, macrophages have been identified as a source of TNFα and iNOS after muscle injury (23,36), and in the present study, iNOS and TNFα mRNA levels were elevated in muscles of old mice despite fewer CD68+ cells present in injured muscles from old compared with young mice at this time point. TNFα is reported to inhibit myogenic differentiation in vitro and in vivo, to lead to the degradation of transcription factors critical for muscle regeneration and to induce apoptosis (35,49), whereas iNOS-expressing macrophages can injure muscle cells in vitro (50) and in vivo (51). Therefore, excessive TNFα or iNOS production by macrophages in muscles of old mice could undermine or impair muscle regeneration in aged animals, although low levels of TNFα and iNOS expression are also reported to benefit muscle repair (23,35). Anti-inflammatory M2 macrophages have been identified as the primary source of IL-10 after muscle injury (36,52). We found less IL-10 mRNA in muscles of old mice relative to young mice at 2 days, in spite of no difference in the levels of CD163+ cells in young and old muscles at this time point. Thus, the age-related decrease in IL-10 mRNA is consistent with impaired IL-10 production in aged macrophages. Impaired IL-10 production can be detrimental to muscle repair. Mice lacking IL-10 showed persistent fiber damage and slowed regeneration and growth after an unloading/reloading muscle injury (10). IL-10 can contribute to repair by several mechanisms. IL-10-stimulated macrophages promote the proliferation and differentiation of myogenic precursors in vitro and in vivo (51,52). IL-10 can also play a regulatory role, suppressing the production of proinflammatory cytokines IL-1, IL-6, and TNFα (48), whereas stimulating production of the growth factor IGF-1 (52). Therefore, our observation of elevated levels of M2 macrophage content in injured muscles from old mice may reflect a compensatory effect of insufficient IL-10 production by these cells. Overall, our findings suggest age-related changes in the expression of macrophage-associated genes that have the potential to undermine or impair muscle regeneration in aged animals. Our study is not without limitations. We did not follow muscles out to full recovery. We cannot therefore definitively state that in the present study muscles of old mice would have failed to recover as quickly or completely as those in young mice; however, age-related impairments in regeneration after muscle injury resulting from a wide range of insults have been firmly established (1–4,38–43), including in studies from our own group using LCs (5,6,14). Thus, sacrificing additional animals to once again confirm this result does not appear justified. A second limitation is that our examination of mRNA levels for myeloid cell genes in the whole tissue provides no definitive information regarding the cellular source of the mRNA. Furthermore, altered mRNA levels do not necessarily reflect changes in protein content. Current studies are aimed at addressing these limitations by isolating different myeloid cell populations from injured muscles of young and old mice and examining mRNA and protein levels of purified cells. Despite some limitations, this study addresses gaps in our knowledge of aging-related changes in the myeloid cell response to injury, and our findings provide support for future studies specifically addressing the hypothesis that an effective degenerative phase is necessary to provide an environment amenable to effective regeneration. Supplementary Material Supplementary data is available at The Journals of Gerontology, Series A: Biological Sciences and Medical Sciences online. Funding This work was supported by the National Institute on Aging at the National Institutes of Health (AG020591 and AG051442 to S.V.B. and AG000114 to D.D.S. and L.A.B.). Conflict of Interest None reported. Acknowledgments We thank the Unit for Laboratory Animal Medicine (ULAM) Pathology Core for Animal Research (PCAR) at the University of Michigan for performing the blood cell counts. References 1. Conboy IM, Conboy MJ, Wagers AJ, Girma ER, Weissman IL, Rando TA. Rejuvenation of aged progenitor cells by exposure to a young systemic environment. Nature . 2005; 433: 760– 764. doi: 10.1038/nature03260 Google Scholar CrossRef Search ADS PubMed  2. Sadeh M. Effects of aging on skeletal muscle regeneration. J Neurol Sci . 1988; 87: 67– 74. doi:10.1016/0022-510X(88)90055-X Google Scholar CrossRef Search ADS PubMed  3. Ghaly A, Marsh DR. Aging-associated oxidative stress modulates the acute inflammatory response in skeletal muscle after contusion injury. Exp Gerontol . 2010; 45: 381– 388. doi: 10.1016/j.exger.2010.03.004 Google Scholar CrossRef Search ADS PubMed  4. Zhou Y, Lovell D, Bethea Met al.   Age-dependent changes cooperatively impact skeletal muscle regeneration after compartment syndrome injury. Am J Pathol . 2014; 184: 2225– 2236. doi: 10.1016/j.ajpath.2014.03.018 Google Scholar CrossRef Search ADS PubMed  5. McArdle A, Dillmann WH, Mestril R, Faulkner JA, Jackson MJ. Overexpression of HSP70 in mouse skeletal muscle protects against muscle damage and age-related muscle dysfunction. FASEB J . 2004; 18: 355– 357. doi: 10.1096/fj.03-0395fje Google Scholar CrossRef Search ADS PubMed  6. Rader EP, Faulkner JA. Recovery from contraction-induced injury is impaired in weight-bearing muscles of old male mice. J Appl Physiol (1985) . 2006; 100: 656– 661. doi: 10.1152/japplphysiol.00663.2005 Google Scholar CrossRef Search ADS PubMed  7. Nguyen HX, Tidball JG. Null mutation of gp91phox reduces muscle membrane lysis during muscle inflammation in mice. J Physiol . 2003; 553: 833– 841. doi: 10.1113/jphysiol.2003.051912 Google Scholar CrossRef Search ADS PubMed  8. Lockhart NC, Brooks SV. Neutrophil accumulation following passive stretches contributes to adaptations that reduce contraction-induced skeletal muscle injury in mice. J Appl Physiol (1985) . 2008; 104: 1109– 1115. doi: 10.1152/japplphysiol.00850.2007 Google Scholar CrossRef Search ADS PubMed  9. Pizza FX, Peterson JM, Baas JH, Koh TJ. Neutrophils contribute to muscle injury and impair its resolution after lengthening contractions in mice. J Physiol . 2005; 562: 899– 913. doi: 10.1113/jphysiol.2004.073965 Google Scholar CrossRef Search ADS PubMed  10. Deng B, Wehling-Henricks M, Villalta SA, Wang Y, Tidball JG. IL-10 triggers changes in macrophage phenotype that promote muscle growth and regeneration. J Immunol . 2012; 189: 3669– 3680. doi: 10.4049/jimmunol.1103180 Google Scholar CrossRef Search ADS PubMed  11. McLennan IS. Degenerating and regenerating skeletal muscles contain several subpopulations of macrophages with distinct spatial and temporal distributions. J Anat . 1996; 188: 17– 28. Google Scholar PubMed  12. Koh TJ, Peterson JM, Pizza FX, Brooks SV. Passive stretches protect skeletal muscle of adult and old mice from lengthening contraction-induced injury. J Gerontol A Biol Sci Med Sci . 2003; 58: 592– 597. doi:10.1093/gerona/58.7.B592 Google Scholar CrossRef Search ADS PubMed  13. Brooks SV, Faulkner JA. Contractile properties of skeletal muscles from young, adult and aged mice. J Physiol . 1988; 404: 71– 82. doi:10.1113/jphysiol.1988.sp017279 Google Scholar CrossRef Search ADS PubMed  14. Brooks SV, Faulkner JA. Contraction-induced injury: recovery of skeletal muscles in young and old mice. Am J Physiol . 1990; 258: C436– C442. doi: 10.1152/ajpcell.1990.258.3.C436 Google Scholar CrossRef Search ADS PubMed  15. Gumucio JP, Flood MD, Phan AC, Brooks SV, Mendias CL. Targeted inhibition of TGF-beta results in an initial improvement but long-term deficit in force production after contraction-induced skeletal muscle injury. J Appl Physiol (1985) . 2013; 115: 539– 545. doi: 10.1152/japplphysiol.00374.2013 Google Scholar CrossRef Search ADS PubMed  16. Pizza FX, Koh TJ, McGregor SJ, Brooks SV. Muscle inflammatory cells after passive stretches, isometric contractions, and lengthening contractions. J Appl Physiol (1985) . 2002; 92: 1873– 1878. doi: 10.1152/japplphysiol.01055.2001 Google Scholar CrossRef Search ADS PubMed  17. Shortreed K, Johnston A, Hawke TJ. Satellite cells and muscle repair. In: Tiidus PM, ed. Skeletal Muscle Damage and Repair . Champaign, IL: Human Kinetics; 2008: 77– 88. 18. Lumeng CN, DelProposto JB, Westcott DJ, Saltiel AR. Phenotypic switching of adipose tissue macrophages with obesity is generated by spatiotemporal differences in macrophage subtypes. Diabetes . 2008; 57: 3239– 3246. doi: 10.2337/db08-0872 Google Scholar CrossRef Search ADS PubMed  19. Fleming TJ, Fleming ML, Malek TR. Selective expression of Ly-6G on myeloid lineage cells in mouse bone marrow. RB6-8C5 mAb to granulocyte-differentiation antigen (Gr-1) detects members of the Ly-6 family. J Immunol . 1993; 151: 2399– 2408. Google Scholar PubMed  20. Serbina NV, Pamer EG. Monocyte emigration from bone marrow during bacterial infection requires signals mediated by chemokine receptor CCR2. Nat Immunol . 2006; 7: 311– 317. doi: 10.1038/ni1309 Google Scholar CrossRef Search ADS PubMed  21. Lee PY, Wang JX, Parisini E, Dascher CC, Nigrovic PA. Ly6 family proteins in neutrophil biology. J Leukoc Biol . 2013; 94: 585– 594. doi: 10.1189/jlb.0113014 Google Scholar CrossRef Search ADS PubMed  22. Lagasse E, Weissman IL. Flow cytometric identification of murine neutrophils and monocytes. J Immunol Methods . 1996; 197: 139– 150. doi:10.1016/0022-1759(96)00138-X Google Scholar CrossRef Search ADS PubMed  23. Rigamonti E, Touvier T, Clementi E, Manfredi AA, Brunelli S, Rovere-Querini P. Requirement of inducible nitric oxide synthase for skeletal muscle regeneration after acute damage. J Immunol . 2013; 190: 1767– 1777. doi: 10.4049/jimmunol.1202903 Google Scholar CrossRef Search ADS PubMed  24. Schmittgen TD, Livak KJ. Analyzing real-time PCR data by the comparative C(T) method. Nat Protoc . 2008; 3: 1101– 1108. doi:10.1038/nprot.2008.73 Google Scholar CrossRef Search ADS PubMed  25. Manfredi TG, Fielding RA, O’Reilly KP, Meredith CN, Lee HY, Evans WJ. Plasma creatine kinase activity and exercise-induced muscle damage in older men. Med Sci Sports Exerc . 1991; 23: 1028– 1034. doi:10.1249/00005768-199109000-00006 Google Scholar CrossRef Search ADS PubMed  26. Zerba E, Komorowski TE, Faulkner JA. Free radical injury to skeletal muscles of young, adult, and old mice. Am J Physiol . 1990; 258: C429– C435. doi: 10.1152/ajpcell.1990.258.3.C429 Google Scholar CrossRef Search ADS PubMed  27. Roche JA, Tulapurkar ME, Mueller ALet al.   Myofiber damage precedes macrophage infiltration after in vivo injury in dysferlin-deficient A/J mouse skeletal muscle. Am J Pathol . 2015; 185: 1686– 1698. doi: 10.1016/j.ajpath.2015.02.020 Google Scholar CrossRef Search ADS PubMed  28. Lockhart NC, Brooks SV. Protection from contraction-induced injury provided to skeletal muscles of young and old mice by passive stretch is not due to a decrease in initial mechanical damage. J Gerontol A Biol Sci Med Sci . 2006; 61: 527– 533. doi: 10.1093/gerona/61.6.527 Google Scholar CrossRef Search ADS PubMed  29. Li Y, Lee Yi, Thompson WJ. Changes in aging mouse neuromuscular junctions are explained by degeneration and regeneration of muscle fiber segments at the synapse. J Neurosci . 2011; 31: 14910– 14919. doi: 10.1523/JNEUROSCI.3590-11.2011 Google Scholar CrossRef Search ADS PubMed  30. Valdez G, Tapia JC, Kang Het al.   Attenuation of age-related changes in mouse neuromuscular synapses by caloric restriction and exercise. Proc Natl Acad Sci USA . 2010; 107: 14863– 14868. doi: 10.1073/pnas.1002220107 Google Scholar CrossRef Search ADS PubMed  31. Greiwe JS, Cheng B, Rubin DC, Yarasheski KE, Semenkovich CF. Resistance exercise decreases skeletal muscle tumor necrosis factor alpha in frail elderly humans. FASEB J . 2001; 15: 475– 482. doi: 10.1096/fj.00-0274com Google Scholar CrossRef Search ADS PubMed  32. Jozsi AC, Dupont-Versteegden EE, Taylor-Jones JMet al.   Molecular characteristics of aged muscle reflect an altered ability to respond to exercise. Int J Sport Nutr Exerc Metab . 2001; 11( suppl): S9– S15. doi:10.1123/ ijsnem.11.s1.s9 Google Scholar CrossRef Search ADS PubMed  33. Przybyla B, Gurley C, Harvey JFet al.   Aging alters macrophage properties in human skeletal muscle both at rest and in response to acute resistance exercise. Exp Gerontol . 2006; 41: 320– 327. doi: 10.1016/j.exger.2005.12.007 Google Scholar CrossRef Search ADS PubMed  34. Costamagna D, Costelli P, Sampaolesi M, Penna F. Role of inflammation in muscle homeostasis and myogenesis. Mediators Inflamm . 2015; 2015: 805172. doi: 10.1155/2015/805172 Google Scholar CrossRef Search ADS PubMed  35. Degens H. The role of systemic inflammation in age-related muscle weakness and wasting. Scand J Med Sci Sports . 2010; 20: 28– 38. doi: 10.1111/j.1600-0838.2009.01018.x Google Scholar CrossRef Search ADS PubMed  36. Arnold L, Henry A, Poron Fet al.   Inflammatory monocytes recruited after skeletal muscle injury switch into anti-inflammatory macrophages to support myogenesis. J Exp Med . 2007; 204: 1057– 1069. doi: 10.1084/jem.20070075 Google Scholar CrossRef Search ADS PubMed  37. Massimino ML, Rapizzi E, Cantini Met al.   ED2+ macrophages increase selectively myoblast proliferation in muscle cultures. Biochem Biophys Res Commun . 1997; 235: 754– 759. doi: 10.1006/bbrc.1997.6823 Google Scholar CrossRef Search ADS PubMed  38. Carlson BM, Faulkner JA. Muscle transplantation between young and old rats: age of host determines recovery. Am J Physiol . 1989; 256: C1262– C1266. doi: 10.1152/ajpcell.1989.256.6.C1262 Google Scholar CrossRef Search ADS PubMed  39. Conboy IM, Conboy MJ, Smythe GM, Rando TA. Notch-mediated restoration of regenerative potential to aged muscle. Science . 2003; 302: 1575– 1577. doi: 10.1126/science.1087573 Google Scholar CrossRef Search ADS PubMed  40. McBride TA, Gorin FA, Carlsen RC. Prolonged recovery and reduced adaptation in aged rat muscle following eccentric exercise. Mech Ageing Dev . 1995; 83: 185– 200. doi: 10.1016/0047-6374(95)01629-E Google Scholar CrossRef Search ADS PubMed  41. Paliwal P, Pishesha N, Wijaya D, Conboy IM. Age dependent increase in the levels of osteopontin inhibits skeletal muscle regeneration. Aging (Albany NY) . 2012; 4: 553– 566. doi: 10.18632/aging.100477 Google Scholar CrossRef Search ADS PubMed  42. Ullman M, Ullman A, Sommerland H, Skottner A, Oldfors A. Effects of growth hormone on muscle regeneration and IGF-I concentration in old rats. Acta Physiol Scand . 1990; 140: 521– 525. doi: 10.1111/j.1748-1716.1990.tb09029.x Google Scholar CrossRef Search ADS PubMed  43. van der Poel C, Gosselin LE, Schertzer JDet al.   Ageing prolongs inflammatory marker expression in regenerating rat skeletal muscles after injury. J Inflamm (Lond) . 2011; 8: 41. doi: 10.1186/1476-9255-8-41 Google Scholar CrossRef Search ADS PubMed  44. Wang Y, Wehling-Henricks M, Samengo G, Tidball JG. Increases of M2a macrophages and fibrosis in aging muscle are influenced by bone marrow aging and negatively regulated by muscle-derived nitric oxide. Aging Cell . 2015; 14: 678– 688. doi: 10.1111/acel.12350 Google Scholar CrossRef Search ADS PubMed  45. Hammers DW, Merritt EK, Matheny RWJret al.   Functional deficits and insulin-like growth factor-I gene expression following tourniquet-induced injury of skeletal muscle in young and old rats. J Appl Physiol (1985) . 2008; 105: 1274– 1281. doi: 10.1152/japplphysiol.90418.2008 Google Scholar CrossRef Search ADS PubMed  46. Hamada K, Vannier E, Sacheck JM, Witsell AL, Roubenoff R. Senescence of human skeletal muscle impairs the local inflammatory cytokine response to acute eccentric exercise. FASEB J . 2005; 19: 264– 266. doi: 10.1096/fj.03-1286fje Google Scholar CrossRef Search ADS PubMed  47. Peake J, Della Gatta P, Cameron-Smith D. Aging and its effects on inflammation in skeletal muscle at rest and following exercise-induced muscle injury. Am J Physiol Regul Integr Comp Physiol . 2010; 298: R1485– R1495. doi: 10.1152/ajpregu.00467.2009 Google Scholar CrossRef Search ADS PubMed  48. Brigitte M, Schilte C, Plonquet Aet al.   Muscle resident macrophages control the immune cell reaction in a mouse model of notexin-induced myoinjury. Arthritis Rheum . 2010; 62: 268– 279. doi: 10.1002/art.27183 Google Scholar CrossRef Search ADS PubMed  49. Langen RC, Van Der Velden JL, Schols AM, Kelders MC, Wouters EF, Janssen-Heininger YM. Tumor necrosis factor-alpha inhibits myogenic differentiation through MyoD protein destabilization. FASEB J . 2004; 18: 227– 237. doi: 10.1096/fj.03-0251com Google Scholar CrossRef Search ADS PubMed  50. Nguyen HX, Tidball JG. Expression of a muscle-specific, nitric oxide synthase transgene prevents muscle membrane injury and reduces muscle inflammation during modified muscle use in mice. J Physiol . 2003; 550: 347– 356. doi: 10.1113/jphysiol.2003.040907 Google Scholar CrossRef Search ADS PubMed  51. Villalta SA, Nguyen HX, Deng B, Gotoh T, Tidball JG. Shifts in macrophage phenotypes and macrophage competition for arginine metabolism affect the severity of muscle pathology in muscular dystrophy. Hum Mol Genet . 2009; 18: 482– 496. doi: 10.1093/hmg/ddn376 Google Scholar CrossRef Search ADS PubMed  52. Bosurgi L, Corna G, Vezzoli Met al.   Transplanted mesoangioblasts require macrophage IL-10 for survival in a mouse model of muscle injury. J Immunol . 2012; 188: 6267– 6277. doi: 10.4049/jimmunol.1102680 Google Scholar CrossRef Search ADS PubMed  © The Author(s) 2018. Published by Oxford University Press on behalf of The Gerontological Society of America. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices)

Journal

The Journals of Gerontology Series A: Biomedical Sciences and Medical SciencesOxford University Press

Published: Apr 19, 2018

There are no references for this article.

You’re reading a free preview. Subscribe to read the entire article.


DeepDyve is your
personal research library

It’s your single place to instantly
discover and read the research
that matters to you.

Enjoy affordable access to
over 18 million articles from more than
15,000 peer-reviewed journals.

All for just $49/month

Explore the DeepDyve Library

Search

Query the DeepDyve database, plus search all of PubMed and Google Scholar seamlessly

Organize

Save any article or search result from DeepDyve, PubMed, and Google Scholar... all in one place.

Access

Get unlimited, online access to over 18 million full-text articles from more than 15,000 scientific journals.

Your journals are on DeepDyve

Read from thousands of the leading scholarly journals from SpringerNature, Elsevier, Wiley-Blackwell, Oxford University Press and more.

All the latest content is available, no embargo periods.

See the journals in your area

DeepDyve

Freelancer

DeepDyve

Pro

Price

FREE

$49/month
$360/year

Save searches from
Google Scholar,
PubMed

Create lists to
organize your research

Export lists, citations

Read DeepDyve articles

Abstract access only

Unlimited access to over
18 million full-text articles

Print

20 pages / month

PDF Discount

20% off