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. Author manuscript; available in PMC: 2020 Dec 1.
Published in final edited form as: J Immunol. 2019 Oct 25;203(11):3013–3022. doi: 10.4049/jimmunol.1900144

Exercise promotes resolution of acute inflammation by catecholamine-mediated stimulation of resolvin D1 biosynthesis

Jing-Juan Zheng *, Ernesto Pena Calderin *, Bradford G Hill *, Aruni Bhatnagar *, Jason Hellmann *
PMCID: PMC6864295  NIHMSID: NIHMS1541025  PMID: 31653685

Abstract

The mechanisms by which regular exercise prevents the development and progression of chronic inflammatory diseases are largely unknown. We find that exercise enhances resolution of acute inflammation by augmenting resolvin D1 (RvD1) levels and by promoting macrophage phagocytosis. When compared with sedentary controls, mice that performed a 4-week treadmill exercise regimen displayed higher macrophage phagocytic activity, enhanced RvD1 levels and earlier neutrophil clearance following an acute inflammatory challenge. In acute inflammatory cell extracts from exercised mice, we found elevated expression of Alox15 and Alox5 and higher RvD1 levels. Because exercise stimulates release of epinephrine, which has immunomodulatory effects, we questioned if epinephrine exerts proresolving actions on macrophages. Epinephrine-treated macrophages displayed higher RvD1 levels and 15-lipoxygenase-1 protein abundance, which were prevented by incubation with the α1 adrenergic receptor (α1-AR) antagonist, prazosin. Likewise, stimulation of the α1-AR with phenylephrine enhanced macrophage phagocytosis and RvD1 production. During acute inflammation, prazosin abrogated exercise-enhanced neutrophil clearance, macrophage phagocytosis, and RvD1 biosynthesis. These results suggest that exercise-stimulated epinephrine enhances resolution of acute inflammation in an α1-AR-dependent manner. These findings provide new mechanistic insights into the proresolving effects of exercise that could lead to the identification of novel pathways to stimulate resolution.

Introduction

Exercise or regular physical activity prolongs lifespan and enhances resilience against chronic diseases. Conversely, physical inactivity and a sedentary lifestyle are major risk factors for several major non-communicable diseases, including type 2 diabetes (T2D), breast and colon cancers, dementia, depression, and cardiovascular disease (CVD).(1) The World Health Organization (WHO) ranks physical inactivity as the fourth leading cause of mortality, linked to 6% of deaths worldwide.(2) Although many of the health benefits of exercise could be attributed to increased cardiorespiratory fitness and improved CVD risk profile, the beneficial effects of exercise extend beyond reduction in CVD risk factors. Favorable changes in blood pressure, lipids, body mass index and glucose tolerance are major mediators of the inverse relationship between physical activity and CVD risk, but other factors, particularly reductions in inflammation also contribute significantly to the beneficial cardiovascular effects of exercise.(3) In a duration and intensity dependent manner, exercise stimulates the release of catecholamines, which due to their immunoregulatory effects may contribute to the anti-inflammatory effects of exercise. Nevertheless, the underlying mechanisms by which exercise modulates inflammatory processes, and how catecholamines may contribute, remain largely unknown.

Bioactive lipid mediators termed specialized proresolving lipid mediators (SPMs), contribute to resolution of inflammation through receptor-mediated actions on leukocytes. These lipids are produced largely through the biosynthetic conversion of omega-3 fatty acids by 15-lipoxygenase (15-LOX) and 5-lipoxygenase (5-LOX). In mice, the SPM resolvin D1 (RvD1) stimulates macrophage phagocytosis by signaling through its cognate surface receptor ALX/FPR2.(4) (5) Recent work from many laboratories, including our own, has shown that chronic inflammatory diseases such as atherosclerosis and diet-induced insulin resistance are associated with a failure in resolution of acute inflammation, and that treatment with SPMs diminishes chronic inflammation and lessens disease severity.(6) Although acute inflammatory responses caused by bouts of relatively intense exercise likely resolve naturally through the actions of endogenous proresolving mediators that are found to be elevated following exercise(7), it remains unclear how, or if, exercise training affects the magnitude and duration of the resolution response.

In this study, we examined the effect of regular exercise on acute inflammation and resolution in mice. Our results show that exercise training enhances neutrophil clearance in a peritonitis model of acute inflammation. The improvement in resolution in exercise-trained mice was associated with increased biosynthesis of the endogenous SPM, RvD1, as well as increased macrophage phagocytosis. Mechanistically, our findings suggest that exercise-induced enhancement of resolution is due to catecholamine-induced activation of the α1-adrenergic receptor (α1-AR). These findings provide new insights into how exercise regulates acute inflammatory processes and offer new ideas into how exercise may prevent or mitigate disease.

Materials and Methods

Animals and Reagents

Male FVB/NJ mice were purchased from Jackson Laboratories at 8 weeks of age. At 9–10 weeks of age, they were either subjected to exercise training or maintained in a normal, sedentary state. Food and water were provided ad libitum, and the animals were maintained on a 12:12 h light-dark schedule. Adrenergic receptor agonists and antagonists were purchased from Sigma-Aldrich. All animal procedures were approved by the University of Louisville Institutional Animal Care and Use Committee. Primary conjugated antibodies, used for flow cytometry, were purchased from eBiosciences.

Exercise training

Mice were subjected to treadmill exercise as previously described.(8) Animals were randomly assigned to sedentary and exercise (Exe) groups and familiarized for 20 min to the rodent exercise treadmill (Columbus Instruments) and its environment for two days. Following familiarization, a baseline exercise capacity test (ECT) was performed on exercise-trained mice. For ECT, mice were placed on the treadmill for 10 min at 0 m/min and 0° incline with the shock grids on. Mice were then given an initial familiarization/warmup period of 9 min at 8.5 m/min (0° incline). Subsequently, the speed was increased by 2.5 m/min after each 3 min interval, thereby subjecting the mice to a linear increase in speed. Their workload was increased by raising the vertical incline of the treadmill by 5° every 9 min up to a maximum of 15° (all angles with respect to the horizontal). During exercise training, if mice spent more than half the time on the shock grid rather than running on the treadmill, the time, distance and speed were recorded and the animals were removed from the treadmill. To set the initial training regimen for each group, we exercised mice at 75% of the maximum average speed attained during the baseline ECT at a 10° inclination. Mice in the Exe group exercised 5 d a week [Monday through Friday (M-F)] for 4 weeks. The regular exercise routine consisted of a 10 min warmup period at 10 m/min (10° incline), followed by 40 min runs during week 1, 50 min runs during week 2, and 60 min runs during weeks 3 and 4. The following week (week 5), the mice exercised on Monday, and were then allowed to recover for 24 h prior to additional experiments (e.g., acute peritonitis) or tissue harvest.

Acute peritonitis and flow cytometry

Self-resolving, acute peritonitis was induced by intraperitoneal (i.p.) administration of zymosan A (0.04 mg/g; Sigma-Aldrich). Inflammatory and resolving peritoneal exudates were collected from sedentary and exercise-trained mice at 0, 4, 12, 24, and 48 h, or when indicated, after zymosan challenge. In select experiments, mice received the α1-AR antagonist, prazosin (2.5 mg/kg; i.p.), 30 min prior and 12 h after zymosan injection. Exudates were collected by lavaging the peritoneum with 3 mL of DPBS−/−. The total number of leukocytes was counted using light microscopy with trypan blue exclusion. Leukocyte differentials were obtained by flow cytometry (Becton Dickinson, BD LSR II equipped with FacsDiva V6.0). Data were analyzed using FlowJo V10 software. Following enumeration, exudate aliquots were centrifuged and resuspended in a flow cytometric analysis buffer (1% FBS in PBS) and incubated with Fc Block (anti-mouse CD16/CD32) for 10 min on ice. Cells were then treated with fluorescein isothiocyanate (FITC)-CD45, phycoerythrin (PE)-conjugated anti-F4/80, and allophycocyanin (APC)-conjugated Ly6G and allowed to incubate on ice for 30 min protected from light. Samples were then washed using flow cytometry analysis buffer and analyzed.

RT-PCR/Immunoblotting

Bone marrow macrophage and peritoneal cellular RNAs were isolated and purified using RNeasy kit (Qiagen). RNA quantity and purity were assessed by UV absorbance with a Nanodrop (ThermoFisher Scientific). AMV reverse transcriptase and oligo DT primers were used to prepare cDNA by PCR. Real-time amplification was performed using PerfeCTa SYBR Green FastMix with ROX (Quanta Biosciences) with a 7900HT Fast Real-time PCR (Applied Biosystems). Commercially available, validated PCR primers for Alox15, Alox5, and Fpr2 were purchased from SA Biosciences. Housekeeping gene Hprt was considered the control, and relative expression was calculated using the 2−ΔΔCT method. For immunoblotting, bone marrow macrophages were plated (1 × 106 cells/well) in 6-well plates in DMEM supplemented with 10% FBS, 1% HEPES and 1% penicillin/streptomycin and incubated overnight at 37°C with 5% CO2. After 24 h, the media was removed and the cells were treated with epinephrine at the indicated dose in fresh culture media for 2 h. For adrenergic receptor experiments, the mice were pretreated with specific receptor antagonists (prazosin 1 μM; betaxolol 0.3 μM; ICI 118,551 0.3 μM; Sigma Aldrich) for 30 min before the addition of epinephrine (1 μM). After removal of the media, the cells were washed once with ice-cold PBS and immediately lysed by the addition of RIPA buffer (Sigma Aldrich) containing protease and phosphatase inhibitors. Samples were briefly sonicated on ice, and protein levels were measured on a Nanodrop (ThermoFisher Scientific) by following a modified Lowry’s method (Bio-Rad). Equal amounts of reduced sample lysates were separated using 10% SDS-polyacrylamide gels (Bio-Rad) and transferred to 0.2-μm PVDF membrane. Membranes were blocked at room temperature for 1 h in 5% powdered milk suspended in 0.1% Tween 20 (TBS-T), and then incubated overnight at 4°C in TBS-T containing anti-mouse primary antibodies to detect 15-lipoxygenase 1 (15-LO; AbCam) and GAPDH (Cell Signaling Technology). The following day, membranes were washed for 1 h with TBS-T and then incubated with HRP-conjugated secondary antibodies for 1 h. Protein abundance was detected following the addition of Luminata Forte ECL chemiluminescence reagent (Millipore). GAPDH was used as a loading control in all samples.

Macrophage phagocytosis

To assess in vivo phagocytic capacity, exercise-adapted mice were injected (i.p.) with PE-labeled latex beads (Cayman Chemical; in 500 μl of sterile saline) 24 h after their final exercise bout. The animals were euthanized after 2 h, and peritoneal lavage exudates were collected in 3 mL of DPBS−/−. Sample aliquots were incubated with anti-mouse CD16/CD32 antibody on ice as described above for 10 min on ice followed by 30 min incubation in the dark with anti-F4/80 antibody (APC conjugated). Using flow cytometry, the mean fluorescence intensity (MFI) of PE (i.e., latex beads) was measured in macrophage (F4/80+) cells. In select experiments, mice received α1-AR antagonist treatment (prazosin, 2.5 mg/kg; i.p.) 30 min prior to latex bead injection. Macrophage phagocytosis was also measured ex vivo in cells isolated from sedentary and exercised mice 24 h following 4 weeks of training. For this, resident peritoneal macrophages were isolated by lavaging the peritoneal cavity with 5mL of DPBS−/−. Peritoneal macrophages were then seeded in duplicate and allowed to adhere to a 96-well plate at 4 × 105 cells/well in 100 μl of DMEM with 10% fetal bovine serum (FBS). After 1 h, non-adherent cells were removed and fresh, pre-warmed medium (DMEM + 10% FBS) containing IgG-opsonized FITC-zymosan A (ratio 10:1; Life Technologies) particles was added. After incubation for 1 h, the cells were washed with DPBS+/+ to remove non-phagocytosed zymosan and incubated for 1 min with Trypan Blue to quench extracellular fluorescence. Phagocytosis was quantified by measuring fluorescence intensity on a spectrophotometer (Perkin Elmer) BioTek plate reader. In select experiments, bone marrow macrophages were plated overnight in a 96-well plate and treated without or with phenylephrine for 30 min in DMEM supplemented with 10% FBS, 1% HEPES and 1% penicillin/streptomycin. The medium was then replaced, and the cells were incubated for 1 h with DMEM supplemented with 10% FBS, 1% HEPES and 1% penicillin/streptomycin containing FITC-zymosan A (10:1) and assessed for phagocytosis using a spectrophotometer, as above. To test the contribution of 15-lipoxygenase, bone marrow macrophages were incubated with 10 μm ML351 (15-lipoxygenase inhibitor) for 1 h prior to phenylephrine treatment.

Macrophage α1 adrenergic receptor expression

Bone marrow macrophages were plated at 1×105 cells per well in a 96-well plate in DMEM supplemented with 10% FBS, 1% HEPES and 1% penicillin/streptomycin. After 24 h, cell media was replaced with fresh media without or with BODIPY FL-labelled prazosin (2 µM; Life Technologies). After 30 min, cells were washed with pre-warmed DPBS+/+ and imaged using EVOS fl fluorescence digital inverted microscope (Advanced Microscopy Group). For peritoneal macrophages, mice were injected (i.p.) with BODIPY FL-labelled prazosin (2.5 mg/kg) or sterile saline. Following euthanasia, peritoneal cells were isolated using 5 mL DPBS−/− lavage. Following centrifugation and removal of supernatant, the cells were incubated on ice in the dark with Fc block for 10 min and then incubated for 30 min in the dark with anti-F4/80 antibody (APC conjugated) and analyzed using a BD LSR II flow cytometer equipped with FACS Diva V6.0 software.

Quantification of resolvin D1 by targeted LC-MS/MS

Cell-free peritoneal lavage fluid was collected from sedentary and exercised animals during the resolution phase (48 h post-zymosan injection) and immediately stored at −80°C. On the day of extraction, the samples were thawed on ice and incubated for 45 min with two volumes of ice-cold methanol. Protein precipitate was pelleted by centrifugation (1000g, at 4°C for 10 min) and the supernatant was collected into 10 mL glass conical tubes. Samples were dried to ~400 μl under a gentle steady stream of nitrogen gas. Before solid phase extraction, deuterium-labeled, internal standards, i.e. d5-RvD2, d5-LXA4, d4-LTB4, d8-5(S)-HETE and d4-PGE2, were added to determine extraction recovery. Solid phase extraction was performed using C18 columns (Waters). Columns were first equilibrated with 3 mL of methanol and 3 mL of H2O. Samples were acidified by adding 9 mL of pH 3.5 HPLC grade H2O and rapidly loaded onto columns. Following sample loading, the columns were neutralized with 5 mL of H2O. Columns were then treated with 5 mL of hexane, and lipid mediator products were eluted and collected by the addition of 5 mL of methyl formate. Samples were completely dried using a steady stream of N2 gas and immediately resuspended in methanol-water (50:50 vol/vol) for LC-MS/MS. For quantification, a Shimadzu LC-20AD HPLC and SIL-20AC autoinjector (Shimadzu Corporation) coupled to an API2000 (AB Sciex) was used. Targeted LC-MS/MS was carried out using an Eclipse Plus C18 column (100 mm × 4.6 mm × 1.8 μm; Agilent) by eluting with mobile phase consisting of methanol:water:acetic acid of 55:45:0.01 (vol/vol/vol) that was ramped to 80:20:0.01 over 8 min and then to 98:2:0.01 over the next 10 min. The flow rate was maintained at 0.4 mL/min throughout. The API2000 was operated in negative ionization mode and RvD1 was detected using multiple reaction monitoring (MRM) with retention time matching. Using authentic RvD1 synthetic standards (Cayman Chemical), a linear calibration curve was generated under the exact same conditions and used to determine RvD1 abundance. Final pg/mL values were determined following sample recovery determination based on internal deuterium-labeled standards and starting sample volumes.

Analytical catecholamine measurements

For UPLC-MS/MS analysis of catecholamines, mouse urine samples were collected following exercise training at the indicated time points, and frozen immediately. As described before(9), samples were thawed on ice, vortexed, diluted with 0.2% formic acid, and mixed with certified isotope-labeled internal standards [(±)-epinephrine-d6, (±)-norepinephrine-d6, (±)-metanephrine-d3, dopamine-d4, 3-methoxytyramine-d4, (±)-4 hydroxy-3-methoxymandelic acid-d3, (±)-normetanephrine-d3, 5-hydroxyindole-4,6,7-d3-3-acetic-d2 acid, 5-hydroxy-3-methoxyphenyl-d3-acetic-d2 acid; Cerilliant; Serotonin-α,α,β,β-d4; CDN Isotopes]. Samples containing internal standards were then analyzed using an UPLC-MS/MS instrument (Acquity Class-H UPLC and Xevo TQ-S micro Triple Quadrupole Mass Spectrometry with an ESI ionization source, all from Waters Corporation). Sample constituents were separated on an Acquity UPLC HSS PFP column (150 mm × 2.1 mm, 1.8 μm, from Waters Corporation) using a binary gradient comprised of 0.2% formic acid (Solvent A) and methanol (Solvent B). The gradient started with 0.5% solvent B at a flow rate 0.4 ml/min for 1 min, and was then ramped up to 95% solvent B at a flow rate of 0.35 mL/min over a 3 min period, where it was maintained for 0.5 min before recycling back to solvent B 0.5% in 0.1 min. The solvent gradient was then held at 0.5% solvent B at a flow rate of 0.4 mL/min for 5.4 min. The column temperature was maintained throughout the run at 40°C. Samples were injected at a volume of 1 µL. Optimized cone voltage and collision energy were used for each individual analytes. For each sample, three multiple reaction monitoring (MRM) transitions were set up: one for quantification, one for confirmation, and one for labeled internal standard. These MRMs were scheduled around the retention time of the analytes and no less than 12 data points were collected for each peak. Analytes in urine samples were quantified using peak area ratio based on 8 point-standard curves, which were run before and after urine samples. TargetLynx Application Manager (Version 4.1, from Waters Corporation) was used for peak integration, calibration, and quantification.

Statistical analysis

Data are presented as means ± standard error of the mean. For small sample sizes, nonparametric statistical tests were used. For direct comparisons, nonparametric two-tailed Mann-Whitney tests were performed. Kruskal-Wallis tests were performed for multiple comparisons, followed by Dunn’s post-test correction. For direct comparisons with larger sample sizes, normality was tested using the D’Augostino-Pearson omnibus test, followed by unpaired two-tailed Student t tests. Parametric one-way or two-way analysis of variance, followed by Bonferroni or Tukey’s post-tests were used for multi-group comparisons. In all cases, a P < 0.05 value was considered statistically significant and is reported in all figures and figure legends. Statistical analysis was performed using GraphPad Prism 6.0.

Results

Exercise training increases RvD1 biosynthesis and promotes resolution of acute inflammation.

To examine whether exercise training affects innate immune responses during the resolution of acute inflammation, we compared the kinetics of leukocyte trafficking between sedentary mice and mice subjected to a 4-week treadmill training protocol (Fig. 1A). As shown in Fig. 1C, neutrophil (PMN) recruitment peaked 12 h after zymosan injection and were near control levels by 48 h. In both sedentary mice and mice subjected to 4 weeks of exercise training, similar numbers of total leukocytes were recruited in response to zymosan challenge (Supp. Fig. 1A). The total number of Gr-1+ PMN was significantly reduced 48 h after zymosan injection in exercised animals (Fig. 1C).

Figure 1: Exercise training enhances RvD1 biosynthesis and resolution of acute inflammation.

Figure 1:

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(A) Schematic of 4-week exercise training and peritonitis protocol. (B) Representative flow cytometry dot plots of peritoneal lavage PMN (CD45+Ly6G+) and macrophages (CD45+F4/80+). (C) Peritoneal PMN kinetics at 0, 4, 12, 24, and 48h post-zymosan injection in sedentary and exercised animals (n=3–5 per time point). (D) Resolution indices calculations based on PMN kinetics in sedentary and exercised animals (n=3–5 per time point). (E) Linear regression analysis of PMN removal during resolution phase in sedentary and exercised animals (n=3–5 per time point). (F) Peritoneal macrophage kinetics at 0, 4, 12, 24, and 48h post-zymosan injection in sedentary and exercised animals (n=3–5 per time point). (G) Resolving peritoneal lavage exudate RvD1 levels in sedentary and exercised animals (48h; n=6). (H) Resolving peritoneal exudate cellular expression of Alox15 (n=8–9), (I) Alox5 (n=4–5), and (J) RvD1 receptor, Fpr2 (48h; n=5) in sedentary and exercised animals. (K) Peritoneal macrophage phagocytosis of opsonized FITC-zymosan (n=3) with representative image (scale bar =100um). (L) Baseline peritoneal RvD1 levels and Fpr2 expression in sedentary and exercised animals (n=3–4). (M) Phagocytosis of FITC-zymosan by naive bone marrow macrophages treated with either control media or in media with FBS replaced with plasma isolated from sedentary or exercised animals (n=5–8). *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

To quantify differences in resolution in sedentary and exercised mice, we used previously defined indices(10) to calculate changes in resolution phase kinetics. Consistent with previous publications showing enhanced PMN recruitment following exercise(11), exercised mice displayed a robust PMN infiltration that peaked (ψmax = 22.1 ± 3.2 cells/exudate) at 12 h (Tmax) (Fig. 1C, D). In response to an acute inflammatory challenge, sedentary mice also showed PMN infiltration that peaked (ψmax = 13.6 ± 3.9 cells/exudate) at 12 h (Tmax) (Fig. 1C, D). Following peak PMN infiltration, the resolution interval is defined as the time point at which 50% (T50) of max PMN are removed (Ri). Despite an average increase in PMN recruitment, exercised mice displayed a shorter Ri (17 h) when compared with sedentary mice (24 h) (Fig. 1D). Of note, the percentage of apoptotic (Annexin V+) PMN at the peak of infiltration (12 h) were not different between sedentary and exercised mice (Supp. Fig. 1B). Furthermore, linear regression analysis revealed heightened PMN removal in exercised mice (Fig. 1E), consistent with enhanced resolution of acute inflammation. Because macrophage-mediated efferocytosis of apoptotic PMN is critical for resolution of inflammation(12), we assessed the trafficking kinetics of peritoneal macrophages in both sedentary and exercise-trained mice following zymosan challenge. We found that the total number of F4/80+ macrophages in the peritoneal cavity was not different between sedentary and exercised mice throughout the acute inflammatory response (Fig. 1F).

We next investigated whether exercise augments RvD1 biosynthesis. Our LC-MS/MS analysis showed significantly elevated levels of RvD1 in resolving lavage exudates of exercised animals compared with sedentary controls (Fig. 1G). Moreover, cell pellets collected during the resolution phase from exercised mice demonstrated an increase in the mRNA expression of the RvD1 biosynthetic enzymes, 15-lipoxygenase (Alox15) and 5-lipoxygenase (Alox5) (Fig. 1H, I). The expression of the RvD1 receptor, Fpr2, was not different between sedentary and exercise-trained mice (Fig. 1J).

The fact that exercised mice subjected to an acute inflammatory challenge had similar macrophage numbers in the peritoneum, yet displayed enhanced PMN removal and higher RvD1 levels suggests that exercise may promote resolution by increasing macrophage phagocytosis. To test this hypothesis, we isolated peritoneal macrophages from exercised and sedentary mice, and measured their capacity to phagocytize opsonized-zymosan particles. As shown in Fig. 1K, macrophages from exercised mice displayed enhanced phagocytic capacity. This improvement in phagocytic capacity manifested by 1 week of exercise training (Supp. Fig. 1C). We next questioned whether basal macrophage phagocytosis could be enhanced in exercised mice due to higher RvD1 abundance. In support of this idea, basal RvD1 concentrations were higher in peritoneal lavage fluid collected from exercised mice compared with that from sedentary animals (Fig. 1L). The basal mRNA expression of Fpr2 in peritoneal cell extracts was not different between exercised and sedentary animals (Fig. 1L). Furthermore, we found that exposure of naïve bone marrow macrophages to plasma isolated from exercised mice increased their phagocytic capacity; surprisingly, plasma from sedentary mice significantly diminished phagocytosis (Fig. 1M). Collectively, these findings suggest that exercise training enhances proresolving pathways, e.g., RvD1, and promotes the resolution of acute inflammation by stimulating macrophage phagocytosis.

Exercise training stimulates epinephrine release and elevates leukocytic adrenergic receptor expression.

Previous work shows that exercise activates the sympathoadrenal system and increases the production of catecholamines(13), which have direct actions on macrophages.(14–16) Therefore, we examined the effects of our exercise training protocol on catecholamine levels. For this, we measured urinary levels of several catecholamines and their metabolites in sedentary and exercised mice. We found that exercise training increased the levels of both epinephrine and its direct metabolite, metanephrine (Fig. 2A); however, the urinary levels of norepinephrine and its metabolite normetanephrine were similar between sedentary and exercised mice (Fig. 2B). Interestingly, exercise training modestly decreased urinary dopamine and serotonin levels (Fig. 2C, Supp. Fig 2A). Exercise training was also associated with a decrease in the urinary levels of vanillylmandelic acid; however, because this metabolite is derived from monoamine oxidase-catalyzed metabolism of both metanephrine and normetanephrine, no clear reason for its decline could be adduced.

Figure 2: Exercise training stimulates epinephrine release and enhances leukocytic adrenergic receptor expression during resolution of acute inflammation.

Figure 2:

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(A) Urinary levels of epinephrine and metabolites, metanephrine and vanillylmandelic acid, in sedentary and exercised animals (n=15–20). (B) Urinary levels of norepinephrine and normetanephrine in sedentary and exercised animals (n=13–21). (C) Urinary levels of dopamine and metabolites, 3-methoxytyramine and homovanillic acid, in sedentary and exercised animals (n=15–21). (D) Resolving peritoneal exudate cellular expression levels of adrenergic receptors in sedentary and exercised animals (48h; n=8–9). (E) Resolving peritoneal cellular exudate expression of catecholamine metabolizing enzymes Comt (n=6), Maoa (n=6), and Maob (n=7) in sedentary and exercised animal (48 h). *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Given the changes in urinary catecholamines and metabolites following exercise training, we next measured the expression of catecholamine receptors during resolution. For this, we isolated cell pellets from exercised and sedentary mice during the resolution phase (at 48 h) of an acute inflammatory response and measured adrenergic receptor mRNA expression. As shown in Fig. 2D, the expression of Adra1a (α1a), Adra1b (α1b), Adra2a (α2a), and Adrab2 (β2) were higher in cells from exercised mice. Moreover, the cell pellets isolated during resolution from exercised mice displayed higher levels of the catecholamine-metabolizing enzymes catecholamine methyl transferase (Comt) and monoamine oxidase a (Maoa), but not monoamine oxidase b (Maob; Fig. 2E). These data suggest that exercise training increases both the release of epinephrine and the expression of adrenergic receptors in leukocyte-rich resolving exudates.

Epinephrine-induced RvD1 production in macrophages is dependent on the α1 adrenergic receptor.

Next, we examined whether stimulation of bone marrow macrophages by catecholamines affects RvD1 synthesis. Treatment of bone marrow macrophages, which express the RvD1 biosynthetic enzymes 15-Lox-1 and 5-Lox (Fig. 3A), with epinephrine increased RvD1 production (Fig. 3B). Interestingly, stimulation of bone marrow macrophages with epinephrine for 6 h also modestly increased 15-Lox-1 protein expression (Supp Fig. 3A, B). To test whether the increase in macrophage 15-Lox-1 upon epinephrine treatment is adrenergic receptor specific, we incubated bone marrow macrophages with prazosin (α1 antagonist), ICI 118,551 (β2 antagonist) or betaxolol (β1 antagonist) prior to epinephrine treatment. Treatment with prazosin, but not ICI 118,551 or betaxolol, abrogated epinephrine-dependent induction of 15-Lox-1 (Fig 3C, D). Prazosin pre-treatment also inhibited epinephrine-stimulated production of RvD1 in bone marrow macrophages (Fig. 3E). Moreover, direct stimulation of the α1 adrenergic receptor (AR) by phenylephrine increased bone marrow macrophage phagocytosis and enhanced RvD1 production (Fig. 3F, G). We next tested whether 15-lox-1 is involved in α1-AR stimulated phagocytosis. For this we treated bone marrow macrophages with the 15-lipoxygenase antagonist ML351 prior to phenylephrine stimulation and detected an abrogation of phenylephrine-induced phagocytosis (Fig. 3H). Using immunofluorescence and flow cytometry, we confirmed the protein expression of α1-AR on the surface of bone marrow macrophages (Fig. 3I) and peritoneal macrophages (Fig. 3J). Collectively, these data suggest that epinephrine induces RvD1 biosynthesis and phagocytosis in bone marrow macrophages in an α1-AR-dependent manner.

Figure 3: Epinephrine-induced RvD1 production is α1 adrenergic receptor dependent.

Figure 3:

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(A) Western blot targeting 15-Lox-1 and 5-Lox in bone marrow macrophages (n=3). (B) RvD1 levels in bone marrow macrophage supernatants following epinephrine incubation (n=6–13). (C, D) Representative Western blot and quantitation for 15-Lox-1 normalized to GAPDH in bone marrow macrophages treated without or with epinephrine and prazosin (α1-AR antagonist), ICI 118,551 (β2-AR antagonist), or betaxolol (β1-AR antagonist) (n=3–5). (E) Fold change of RvD1 levels in bone marrow macrophage supernatants treated without or with epinephrine and α1-AR antagonist prazosin (n=3–4). (F) Bone marrow macrophage phagocytosis of opsonized FITC-zymosan without or with phenylephrine (α1-AR agonist) at indicated concentration (n=11). (G) RvD1 levels in bone marrow macrophage supernatants treated with phenylephrine (α1-AR agonist) at indicated concentration (n=3–8). (H) Bone marrow macrophage phagocytosis of FITC-zymosan following phenylephrine (α1-AR agonist) without or with ML351 (15-Lox inhibitor) (n=5). (I)Representative fluorescence microscopy image of bone marrow macrophages treated without or with BODIPY-Prazosin (scale bar=100um). (J) Representative flow cytometry dot plots of peritoneal macrophages (F4/80+) treated without or with BODIPY-Prazosin. *P < 0.05; **P < 0.01; ***P < 0.001.

Exercise-enhanced resolution of inflammation is dependent on the α1-AR.

Given that epinephrine-induced RvD1 biosynthesis is dependent on α1-AR signaling in bone marrow macrophages, we questioned whether exercise-enhanced resolution of acute inflammation could also be mediated by the α1-AR. We found that during the resolution phase of acute inflammation, exercised mice treated with prazosin displayed a 4 h delay in the Ri and a significant reduction in PMN removal (Fig. 4A). Moreover, exercised mice treated with prazosin had higher levels of total leukocytes and PMN during the resolution phase of acute inflammation (Fig. 4B). Of note, prazosin treatment had no effect on total leukocyte or PMN levels in sedentary mice (Fig. 4B). To determine whether prazosin-induced abrogation of exercise-enhanced resolution was due to inhibition of RvD1 production, we measured RvD1 in resolving exudates from sedentary and exercised mice pretreated with vehicle or prazosin. As shown in Fig. 4C, resolving exudates from exercised mice contained higher RvD1 levels, and pretreatment with prazosin completely blocked exercise-mediated induction of RvD1 (Fig. 4C). Although prazosin did not affect the phagocytosis of IgG-coated latex beads by macrophages in sedentary mice, it remarkably diminished macrophage phagocytosis in exercised mice (Fig. 4D). Collectively, these results suggest that α1-AR-dependent signaling contributes to exercise-enhanced resolution of acute inflammation.

Figure 4: Exercise-enhanced resolution of inflammation is α1-dependent.

Figure 4:

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(A) PMN (CD45+Ly6G+) enumeration at 12 and 48 h post-zymosan injection during the resolution phase in sedentary and exercised animals treated without or with prazosin (n=3–6 per time point). (B) Enumeration of leukocytes (CD45+), macrophages (CD45+F4/80+) and neutrophils (CD45+Ly6G+) during resolving phase in sedentary and exercised animals treated without or with prazosin (48 h; n=3–6). (C) Resolving peritoneal lavage exudate RvD1 levels in sedentary and exercised animals treated without or with prazosin (48 h; n=5–6). In vivo peritoneal macrophage (F4/80+) phagocytosis of IgG-coated latex beads with representative mean fluorescence intensity histogram in sedentary and exercised animals treated without or with prazosin (n=5–10). *P < 0.05.

Discussion

The results of this study demonstrate that exercise training accelerates the resolution of inflammation in mice. We found that exercise training increases the basal levels of the resolution agonist RvD1, which contributes to exercise-enhanced resolution of acute inflammation by increasing macrophage-mediated efferocytosis of PMN. Our results also show that epinephrine increases RvD1 production in macrophages in an α1-AR-dependent manner. Likewise, α1-AR blockade with prazosin in vivo abrogates exercise enhancement of PMN clearance, macrophage phagocytosis, and RvD1 production. Collectively, these results suggest that exercise training enhances the resolution of acute inflammation in an α1-AR-dependent manner, which supports the notion that the beneficial effects of exercise on innate immune function may relate to enhancement of proresolving pathways.

Although acute inflammation in response to exercise is an area of intense inquiry, few studies address how exercise affects the resolution of inflammation. Some studies have identified augmented levels of SPMs after exercise. Strenuous exercise in humans has been shown to increase urine levels of lipoxin A4, an arachidonic acid-derived SPM.(17) Similarly, Markworth et al. reported that, in humans, post-exercise recovery following a single bout of resistance exercise is characterized by an increase in circulating levels of the SPMs RvD1, RvE1, and the protectin D1 isomer, PDx.(7) An increase in n-3 DPA-derived resolvins has also been reported following intense exercise training.(18) These reports, which are consistent with our results showing increased RvD1 production and upregulation of its biosynthetic enzymes, support the view that acute exercise training stimulates SPM production. Our results, using a well-controlled animal model of acute inflammation, further assess the biological significance of enhanced SPM production following exercise training, and show that the increases in SPM production are accompanied by increased macrophage phagocytosis and resolution of acute inflammation.

Our studies suggest that the proresolving effects of exercise are at least in part due to catecholamines, which increase transiently during exercise.(19) Catecholamine release during exercise is responsible for altering vascular tone, promoting lipolysis in adipose tissue, increasing glucose output from the liver, and diminishing glucose entry into skeletal muscle.(20) In addition, exercise-induced increases in catecholamines prime the immune system and promote leukocytosis through a β-AR-mediated mechanism.(21–25) Catecholamines also exert receptor-specific immunomodulatory effects on macrophages. Activation of the β2 adrenergic receptor on macrophages promotes alternatively activated M2 polarization with dampened TNFα and IL-1β cytokine production.(26) Conversely, β1 activation enhances LPS stimulation and inflammatory cytokine production.(27, 28) Limited data exist regarding the role of α1 adrenergic receptor on macrophage function; however, the α1-AR agonist, phenylephrine, has been reported to stimulate rat peritoneal macrophage phagocytosis, while co-culturing rat microglial cells with LPS and phenylephrine decreases TNFα and IL-6 expression.(29),(30) These findings are consistent with our data showing enhanced phagocytic capacity in α1-AR-stimulated bone marrow macrophages. Additional studies on mouse peritoneal macrophages have shown that norepinephrine reduces LPS-stimulated TNFα production in an α2 dependent manner,(14) suggesting that α-ARs exert an anti-inflammatory effect on macrophages.

Our study adds to this body of knowledge by showing that epinephrine increases 15-Lox-1 expression and RvD1 biosynthesis in macrophages in an α1-AR-dependent manner. Because RvD1 has been shown to repress TNFα production,(31) our results appear consistent with the previously observed anti-inflammatory effects of epinephrine and α1-AR signaling. Our results further suggest that the phagocytic enhancement of α1-AR activation in macrophages is associated with elevated RvD1 production and that enhanced resolution in exercised animals is dependent on α1-AR signaling. These results suggest that resolution enhancement with exercise depends on the adrenergic system. Although we report that exercise increases the mRNA expression of leukocytic Alox15 (15-Lox-1) and that macrophages treated with epinephrine display increased protein expression of 15-Lox-1 and RvD1 biosynthesis, the extent to which exercise-stimulated 15-Lox-1 expression enhances resolution of inflammation remains unclear. Increases in 15-Lox-1 in other conditions can be associated with pathological processes, such as the atherogenic oxidation of LDL.(32)–(33) Nevertheless, our data showing enhanced macrophage phagocytosis with α1-AR activation is dependent upon 15-Lox-1 activation are consistent with the hypothesis that exercise-induced expression of 15-Lox-1 promotes resolution of inflammation through enhanced RvD1 biosynthesis.

The novel link between exercise and the resolution of inflammation suggests that exercise may be an effective intervention not only in promoting the resolution of acute inflammation, but in resolving chronic inflammation as well. Studies from our group and others38 have shown that non-resolved inflammation is associated with the accumulation of apoptotic PMN and defective SPM biosynthesis.(34–36) Such non-resolved inflammation has been linked to diminished macrophage phagocytosis and the development of chronic disease.(36) For example, defective macrophage phagocytosis and delayed resolution is a common feature of atherosclerotic lesions and type 2 diabetes, and non-resolving inflammation in these conditions is associated with defective biosynthesis of SPMs, including RvD1. (37) (38) (34) Importantly, we found that treatment with RvD1 restores macrophage phagocytosis and reduces apoptotic PMN burden in diabetic wounds and reduces lesion size in atherosclerosis-prone mice. (34) (37) Taken together, this work supports the view that delayed resolution, due to defects in SPM production, is linked to non-resolving inflammation in chronic diseases. Given that physical activity and exercise are associated with a decrease in chronic inflammation in both non-diseased and diseased conditions, we propose that exercise-enhanced improvements in resolution may in part be responsible for the salutary changes in inflammation that mediate the beneficial cardiovascular effects of exercise.(3) Our results show that exercise training increases the basal levels of RvD1, a potent SPM that is decreased in states of non-resolving inflammation such as diabetes and atherosclerosis. We also found that exercise stimulates the removal of PMN, which led to a reduction in the resolution interval following zymosan-induced peritonitis. This is consistent with extensive previous work showing that acute exercise promotes recovery from infections and that long-term physical activity and exercise can diminish mortality risk associated with cardiovascular disease and diabetes.(39–41) Although additional work is required to assess the extent to which exercise-induced macrophage α1-AR signaling and resolution of inflammation improve cardiovascular health and insulin sensitivity, our current findings strengthen the rationale for studying the effects of exercise on acute inflammatory responses and on the chronic low-grade inflammation associated with chronic disease.

In summary, our findings suggest that the beneficial effects of exercise on inflammatory processes are due to α1-AR-dependent activation of resolution. The extent to which different SPM species contribute to exercise-induced resolution of inflammation in different tissues or in chronic diseases remains to be determined; however, the findings of the current study suggest that RvD1 is a critical mediator of exercise-induced resolution of acute inflammation. These findings provide critical insights into the health-promoting effect of exercise and could inform future studies to delineate how exercise mitigates chronic inflammation and disease.

Supplementary Material

1

Key Points.

  • Exercise enhances resolution of acute inflammation and RvD1 biosynthesis.

  • Epinephrine stimulates macrophage production of RvD1 through α1 adrenergic receptor.

  • α1 adrenergic receptor blockade in vivo abrogates exercise-enhanced resolution.

Acknowledgements

We would like to thank the University of Louisville Diabetes and Obesity Center Flow Cytometry and Bioanalytical Core for expert technical assistance with leukocyte differentials and urinary catecholamine measurements, respectively.

This work was supported in part by National Institutes of Health grants GM127607 (A.B.), GM127495 (J.H.), HL130174, ES028268, HL078825, and HL147844 (B.H.), and the American Diabetes Association Pathway to Stop Diabetes Grant (1–16-JDF-041; B.H.).

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