PENCE, BRANDT D.1,2; DIPIETRO, LUISA A.3,4; WOODS, JEFFREY A.1,2,5,6
Obesity is a major health concern in the United States. From 13.4% prevalence in 1962, the percentage of US adults considered obese reached 30.9% in the year 2000 (8). This trend has continued, because the most recent national data indicates that nearly 34% of Americans can be considered obese, with more than 30% of the remainder now classified as overweight (7). Obesity and obesity-related disorders account for more than 300,000 deaths per year in the United States alone (1,13) and are estimated to be responsible for nearly $150 billion in health care costs per year in the United States (6).
Obesity and obesity-associated disorders such as cardiovascular disease (25), infectious disease (20), metabolic syndrome/diabetes (26), cancer (17,29), and chronic kidney disease (14) are known to involve a dysregulation of the immune response including up-regulated inflammation (9,10) at both the systemic and local adipose tissue levels. This aberrant inflammatory response is associated with impairments in wound healing in obese individuals (38). This includes delays in healing of acute wounds and failure to heal chronic wounds; the latter of which leaves the individual prone to infection and other complications. Type 2 diabetes, a frequent comorbidity of obesity, is a leading cause of amputations in the United States annually (33).
Some studies have shown that delayed healing in animal models of obesity is associated with dysregulated inflammation. In genetically obese mice, delayed healing has been shown to be associated with increased wound levels of proinflammatory chemokines (37) and cytokines (12). Moreover, depletion of inflammatory cells, including neutrophils (2) and macrophages (12), as well as treatment with neutralizing antibodies against proinflammatory cytokines such as tumor necrosis factor (TNF)-α (12) has been shown to speed healing in genetically obese animal models. Delayed healing may be mediated by prolonged induction of inflammation through macrophage dysfunction, because wound-associated macrophages in obese animals have been shown to fail to perform essential functions such as clearance of apoptotic neutrophils (22), a process that is necessary to switch the macrophage from a proinflammatory phenotype to an anti-inflammatory, prohealing phenotype. Thus, interventions that target increased wound-associated inflammation and speed healing in obese individuals are of major public health and research interest.
Exercise has been shown to decrease inflammation in obese individuals at both systemic (4,30) and adipose tissue-specific (35) levels. However, no studies to date have examined the effects of exercise training on healing rate or wound inflammation using an obesity model of delayed healing. Interestingly, exercise has been shown to speed healing in aged mice (21) and older humans (3) and reduce inflammation in the wounded tissue of aged mice (21). Because aging increases basal levels of inflammation similar to that seen in obese individuals (23), similar mechanisms might allow exercise to speed healing in obese and diabetic individuals.
Therefore, the goal of this study is to determine the effects of exercise on the wound healing process in obese mice. We hypothesized that exercise would speed healing rate in obese mice similar to that shown in aged mice and that this would be associated with a reduced level of inflammation in the wound tissue as in our previous aging study (21). A diet-induced obesity (DIO) model in which obesity is induced by feeding of a high-fat diet (HFD) (45% kcal from fat) was chosen because evidence suggests that DIO models are most appropriate for studies of acute wounds, whereas genetically obese models are most appropriate for chronic, nonhealing wound models (32). The intent of the exercise model used in this study as well as in the Keylock et al. (21) study in aged mice is to determine whether a short-duration exercise training program performed within a known time frame before and after wounding can speed healing and lessen wound inflammation. The exercise paradigm and time points for the analyses described both in Keylock et al. (21) and in the present study are intended to contrast similar studies, which show an impaired healing in mice subjected to restraint stress (19).
This intervention is targeted as a type of “prehabilitation” for obese individuals who are scheduled to undergo procedures such as bariatric surgery that will create cutaneous wounds. That is, should exercise be shown to speed healing rates in obese humans in future studies, short-term exercise training such as that described in this study may be prescribed to patients who will be undergoing surgery to help their wounds to heal faster after surgery. Because wounds from such procedures are known to heal more slowly in obese individuals (38), results from this preclinical study may have clinical potential.
Six-week-old female C57Bl/6J mice (Jackson Labs, Bar Harbor, ME) were maintained individually in standard wire-top cages in an AAALAC-accredited animal care facility for 1 wk before the start of experiments. Mice were allowed ad libitum access to water and to the diets. The mice were housed on a 12-h reverse light–dark cycle with the dark period from 1000 to 2200 h. HFD-fed mice were housed for 16 wk and given an experimental diet consisting of 45% total kilocalorie from fat (Research Diets, New Brunswick, NJ). Food intake and body weight were measured every 2 wk during the feeding portion of the study. In addition, food intake and body weight were assessed at day −3, day 0 (when wounds were applied), and day 6 during the exercise portion of the study. For characterization of the exercise effect on healing kinetics in chow-fed mice, 20-wk-old female C57Bl/6J mice (Jackson Labs) were maintained as described previously for 4 wk before wounding. The four groups hereinafter are called HFD-exercise (HFD-Ex), HFD-sedentary (HFD-Sed), chow-exercise (Chow-Ex), and chow-sedentary (Chow-Sed). All procedures were approved by the Institutional Animal Care and Use Committee of the University of Illinois at Urbana-Champaign.
Mice in the exercised groups ran on a motorized treadmill (Jog-a-Dog, Ottawa Lake, MI) in individual lanes at 12 m·min−1 and 5% incline for the final 30 min of the light period (0930–1000 h) starting at 3 d (day −3) before wounding and continuing until 5 d (day 5) after wounding for a total of 9 d of exercise. Gentle prodding was used to encourage mice to continue running. This exercise protocol is similar to that previously used by our laboratory to study wound healing kinetics with exercise in aged mice (21). Nonexercised mice remained sedentary during this time but were exposed to noise and handling similar to that of the exercisers.
The wounding procedure, which results in two full-thickness excisional cutaneous wounds, is well established and has been used in our laboratory in a previous study (21). Briefly, mice were anesthetized with isoflurane in oxygen at a rate of 2–3 L·min−1 starting at 1 h after exercise on day 0. Although anesthetized, mice were shaved and treated with three alternating scrubs of povidone–iodine (Purdue Pharma, Stamford, CT) and isopropyl alcohol. The skin was then folded over and excised using a 6.0-mm-diameter sterile, disposable punch biopsy instrument (HealthLink, Jacksonville, FL) to create two full-thickness dermal wounds. Mice were immediately photographed for assessment of baseline wound size and then returned to their home cage to recover. Return to consciousness after isoflurane administration takes about 30 s using this method. Mice were monitored until full recovery from the wounding procedure. No postsurgical pain medication, bandaging, or hemorrhage control was provided to avoid any potential effects of such treatment on healing rate.
Analysis of wound closure
Mice were photographed each day from day 0 to 10 after wounding to assess wound area. Briefly, mice were anesthetized with isoflurane administered with oxygen at a flow rate of 2–3 L·min−1, and wounds were photographed before recovery from anesthesia. Wound size was analyzed by photoplanimetry as previously described (21). Wound photograph files were recoded by an investigator otherwise uninvolved with the study for blinding purposes before analysis.
Analysis of gene and protein expression
Mice used for gene and protein expression analysis were run in the same manner as described and were euthanized at day 1, day 3, or day 5 after wounding. Mice were euthanized via rapid CO2 asphyxiation, and wounds were immediately harvested using an 8-mm punch biopsy needle (HealthLink) and frozen on dry ice. Frozen wound tissue was stored at −80°C until processing.
Total ribonucleic acid (RNA) was isolated from frozen tissue using Trizol® (Invitrogen, Carlsbad, CA) according to the manufacturer’s instructions. RNA purity was assayed using an automated microvolume NanoDrop spectrophotometer (Thermo Fisher, Waltham, MA). RNA samples with 260:280 ratios near 2.0 were used for further analyses. Isolated RNA was reverse transcribed to complementary deoxyribonucleic acid (cDNA) using a high-capacity cDNA reverse transcription kit (Applied Biosystems, Carlsbad, CA) according to manufacturer’s instructions. The resulting cDNA was stored at −20°C until real-time polymerase chain reaction (RT-PCR) analysis for gene expression of selected inflammatory markers. RT-PCR analysis was performed on a high-throughput RT-PCR machine (Applied Biosystems) according to the manufacturer’s instructions. All samples were run in duplicate.
Cytokines and chemokines tested for gene expression were interleukin (IL)-1β, IL-10, TNF-α, monocyte chemoattractant protein (MCP)-1, and keratinocyte chemoattractant (KC). Gene expression of platelet-derived growth factor (PDGF) was also assessed. TaqMan (Applied Biosystems) master-mix reagent and commercially available, prevalidated primer-probe sets (Applied Biosystems) were used for all analyses. Expression of β-actin was used as the housekeeping gene, and expression of all genes was related to the housekeeping gene and to the referent (HFD-Sed day 1) group by the 2−ΔΔCt method (39).
Wound protein analyses of cytokines IL-1β, IL-10, and TNF-α were carried out by enzyme-linked immunosorbant assay (ELISA). Briefly, wounds were homogenized in ice-cold phosphate-buffered saline with 0.5% v/v protease inhibitor (Sigma-Aldrich, St. Louis, MO) and centrifuged at 10,000g to remove insoluble material. Supernatant was removed and assayed for total protein concentration by a commercially available protein assay (DC Protein Assay; Bio-Rad, Hercules, CA). Protein was then stored at −20°C until analysis. Commercial ELISA matched antibody sets (murine IL-10 (R&D Systems, Minneapolis, MN), murine IL-1β and murine TNF-α (PeproTech, Rocky Hill, NJ)) were used for all analyses. Dilutions for protein samples were as follows: IL-1β, 1:3; IL-10, no dilution; and TNF-α, 1:1. All samples were diluted in phosphate-buffered saline supplemented with 1% bovine serum albumin (Sigma-Aldrich, St. Louis, MO).
ELISAs were performed according to manufacturer’s instructions with the following modifications. For the IL-1β and TNF-α assays, streptavidin-conjugated horseradish peroxidase (#890803, R&D Systems) was substituted for the provided streptavidin-conjugated horseradish peroxidase reagent and diluted 1:200 for use. For all assays, color development was carried out using a commercial substrate reagent set (BD OptEIA TMB Substrate Reagent Set; BD Biosciences, San Diego, CA), and reactions were stopped with 2 N H2SO4. Color development was read at 450 nm on a spectrophotometric plate reader (BioTek, Winooski, VT). Protein concentrations are expressed relative to total protein isolated from each wound sample (picograms of cytokine per milligram of total protein).
Glucose tolerance testing
Mice were assessed for insulin sensitivity via intraperitoneal glucose tolerance testing (GTT). GTT was performed on a subset of mice in each group 1 wk before the exercise intervention as previously described (35). Mice were fasted overnight before GTT. At time 0, blood glucose was measured by tail nick using an automated glucometer (Johnson & Johnson, Langhorne, PA); after which, mice were given 1 g·kg−1 glucose intraperitoneally. Blood glucose was similarly measured at 30, 60, 90, and 120 min after injection. For the tail nick, a small portion of the tip of the tail was cut off using scissors, and blood (one to two drops) was collected. Clotting takes place within less than 2 min after nick, and only a small amount of blood is lost. GTT values were compared between HFD- and chow-fed groups as a marker of metabolic derangement induced by the HFD. To determine whether exercise-induced alterations in glucose tolerance were related to wound healing, a subset of mice underwent GTT at day 1 after wounding in addition to the baseline testing. GTT was performed after an overnight fast, 24 h after the previous exercise bout.
All results are expressed as mean ± SEM. Significance level was set at α = 0.05. Body weight and food intake differences in HFD-fed and chow-fed mice were assessed by unpaired Student’s t-tests. Changes in body weight during the exercise intervention were analyzed by paired t-tests. Glucose tolerance tests were compared by repeated-measures ANOVA (RM-ANOVA) with Tukey HSD post hoc analysis. GTT differences were also expressed as a comparison of area under the glucose response curve (AUC). RM-ANOVA (time × treatment – activity or diet) was used to analyze wound healing kinetics from days 0–10 and or from days 0–5 after wounding. Gene and protein expression levels were analyzed by a 3 × 2 factorial ANOVA (day × activity). GTT response to exercise was analyzed by a 2 × 2 factorial ANOVA (time point × activity). In the instance of a significant interaction or treatment main effect, mean separation was carried out using Tukey HSD post hoc analysis. All analyses were carried out using SPSS v. 19.0 software (IBM, Somers, NY).
Body Weight and Food Intake
By week 16 of the feeding intervention, HFD body weights were significantly higher than those of the chow group, as expected (Table 1). Food intake in grams per day was significantly lower in HFD compared with chow animals (Table 1). However, when expressed as energy intake (kcal·d−1), HFD had a greater energy intake compared with chow animals, which explains their greater weight gain after 16 wk of the study. There were no differences between exercised and sedentary mice in body weight or food intake, and the short (e.g., 9 d) exercise intervention did not result in significantly different food intake or weight loss compared with sedentary controls in either diet interventions (data not shown). In mice used for wound cytokine analysis, there was a significant reduction of body weight in HFD-Ex mice euthanized at day 1, although the magnitude of the weight loss was small (0.9 g) and similar to nonsignificant changes in HFD-Ex mice euthanized at other time points as well as in HFD-Sed mice euthanized at all time points (Table 2). There were no significant differences in pre– or post–body weight or in food intake either between groups (Ex vs. Sed) or within groups at different time points (Table 2).
GTT was performed 1 wk before the wounding (week 15 of feeding) on a subset of HFD-fed (n = 10) and chow-fed (n = 5) mice. As expected, HFD-fed mice had an impaired response to glucose injection (Fig. 1A), indicating a higher degree of insulin resistance in these mice (diet × time interaction, F1.9,24.3 = tolerance (HFD AUC, 39,132 ± 2445; chow AUC, 28,422 ± 1244; P = 0.002). There were no differences between HFD-Ex and HFD-Sed mice at the 15-wk baseline GTT measure (data not shown).
On a subset of mice (n = 5 HFD-Ex, n = 5 HFD-Sed), GTT was measured at day 1 after wounding in addition to baseline (Fig. 1B). Although HFD-Ex had slightly higher glucose intolerance at baseline (before exercise), there was no significant interaction (activity × time point, F1,8 = 0.160, P = 0.699), nor were there significant time point (baseline vs. day 1, F1,8 = 3.072, P = 0.188) or activity (Ex vs. Sed, F1,8 = 2.971, P = 0.123) main effects, indicating that the exercise intervention had little effect on insulin resistance.
As in previous studies (27,28), HFD feeding slowed wound healing rate (Fig. 2A). When comparing wound healing kinetics in sedentary HFD-fed and chow-fed mice for 10 d after wounding, there was a significant time × diet interaction (F2.9,39.4 = 3.655, P = 0.021) and significant main effects for both diet (F1,13 = 6.495, P = 0.024) and time (F2.9,39.4 = 59.955, P = 0.000). Post hoc analysis revealed that chow-fed mice had significantly smaller wounds starting at day 3 and continuing to day 10 after wounding (Fig. 2A, P < 0.05). We also tested whether exercise training could speed healing in both chow-fed and HFD-fed mice. There was no additional effect of exercise on wound healing in chow-fed mice (Fig. 2B, time × activity interaction, F3.1,43.1 = 0.690, P = 0.567). In HFD-fed mice, RM-ANOVA indicated a nonsignificant time × activity interaction (Fig. 2C, F3.0,33.4 = 1.669, P = 0.192) but a main effect of activity that trended toward significance (Fig. 2C, F1,11 = 3.267, P = 0.098). It should be noted that chow and HFD mice in Figure 2A are identical with Chow-Sed and HFD-Sed in Figures 2B and C, respectively. We have arranged the figure in this fashion because we have found that display of the figure in a 2 × 2 factorial arrangement obscures some of the important comparisons that we have highlighted here.
However, in our previous study in aged mice (21), the major effect of exercise was seen early (within 5 d) after wounding. A qualitative evaluation of our healing data suggested that this might have been the case in the present study as well, because the differences in wound size between HFD-Ex and HFD-Sed were greater in the early part (e.g., day 1–5) compared with that in the late part (e.g., day 8–10). Because of this, and because our original hypothesis was exercise would alter wound inflammation early after wounding, we analyzed the effect of exercise on healing kinetics early (day 0 through day 5) after wounding. When RM-ANOVA was performed only on days 0–5, analysis revealed a nearly significant time × activity interaction (Fig. 2C, F2.2,24.1 = 3.229, P = 0.053) and a significant main effect of activity (Fig. 2C, F1,11 = 4.856, P = .050). Post hoc analysis revealed significant differences in wound size between HFD-Ex and HFD-Sed mice at day 1, day 4, and day 5 after wounding (Fig. 2C, P < 0.05).
Wound gene and protein expression
Healing kinetics data described previously demonstrated a) that exercise effects healing of only HFD-fed and not chow-fed mice and b) that the major effects of the exercise bout happened early (within 5 d) after the wound was applied. This is similar to previous research in our laboratory using aged mice (21). Early responses to wounding consist primarily of inflammation (15), and obesity is known to impair the resolution of the inflammatory state prolonging the healing process (27,28). Thus, as in our previous study, we focused our analyses on day 1, day 3, and day 5 after wounding and limited our analyses to HFD-fed mice because exercise did not appear to affect wound healing in chow-fed mice in any way.
We measured gene and protein expression of proinflammatory cytokines IL-1β and TNF-α as well as the anti-inflammatory cytokine IL-10. Previous research has demonstrated that obesity increases levels of proinflammatory cytokines and reduces levels of anti-inflammatory cytokines in the wound environment (5,12,31,37). Surprisingly, exercise training did not affect gene expression of any of the cytokine markers measured in this study. There was neither day × activity interaction (F2,59 = 1.114, P = 0.335) nor main effect for activity (F1,59 = 0.042, P = 0.839) or day (F2,59 = 0.449, P = 0.640) for TNF-α gene expression (Fig. 3A). Likewise, there was neither day × activity interaction (F2,59 = 1.058, P = 0.729) nor main effect for activity (F1,59 = 0.022, P = 0.882) or day (F2,59 = 0.290, P = 0.749) for IL-1β gene expression (Fig. 3B). Finally, there was neither day × activity interaction (F2,59 = 0.448, P = 0.641) nor main effect for activity (F1,59 = 0.711, P = 0.139) or day (F2,59 = 1.093, P = 0.342) for IL-10 gene expression (Fig. 3C).
Although gene expression was unaltered, we hypothesized that posttranscriptional processes might alter the expression of inflammatory proteins. Therefore, we tested wound protein levels of IL-1β, TNF-α, and IL-10 via ELISA at the same time points. Similar to the results of the gene expression assays, exercise training had no effect on protein expression of these cytokines. There was no significant day × activity interaction (F2,58 = 0.921, P = 0.404) and no significant main effect for activity (F1,58 = 0.285, P = 0.596) for TNF-α (Fig. 4A), although there was a significant main effect of day (F2,59 = 15.152, P = 0.000). Likewise, there was no significant day × activity interaction (F2,56 = 0.060, P = 0.942) and no significant main effect of activity (F1,56 = 0.379, P = 0.540) for IL-1β (Fig. 4B), although again, there was a significant main effect of day (F2,56 = 9.021, P = 0.000). Finally, there was no significant day × activity interaction (F2,55 = 0.304, P = 0.739) and no significant main effect of either activity (F1,55 = 0.329, P = 0.568) or day (F2,55 = 0.150, P = 0.861) for IL-10 (Fig. 4C).
We additionally hypothesized that exercise might affect influx of inflammatory cells by modulating chemokine expression in the wound tissue. Therefore, we measured gene expression of macrophage chemokine MCP-1 and neutrophil chemokine KC. There was no significant day × activity interaction (F2,57 = 0.470, P = 0.627) and no significant main effect for activity (F1,57 = 0.010, P = 0.920) for MCP-1 (Fig. 5A), although there was a significant main effect of day (F2,57 = 13.039, P = 0.000). Likewise, there was no significant day × activity interaction (F2,59 = 0.312, P = 0.733) and no significant main effect of activity (F1,59 = 0.796, P = 0.376) for KC (Fig. 5B), although again, there was a significant main effect of day (F2,59 = 8.480, P = 0.001).
Because we found no effect of exercise on wound inflammation in obese mice, we hypothesized that exercise might affect other aspects of the early healing response such as hemostasis. Therefore, we tested gene expression of PDGF, a growth factor released upon platelet activation. There was no significant day × activity interaction (F2,59 = 0.091, P = 0.913) and no significant main effect of activity (F1,59 = 0.218, P = 0.642) for PDGF (Fig. 5C). There was a significant main effect of day (F2,59 = 11.859, P = 0.000).
The major finding of this study was that short-term treadmill exercise (3 d before and 5 d after wounding) speeds wound healing rate in obese, HFD-fed female mice. This is in line with previous studies in aged mice (21) and older adults (3), but to the best of our knowledge, this is the first report of an exercise effect on cutaneous wound healing using an obesity model. Obesity is known to impede wound healing (38), and this has been demonstrated in animal models of obesity (32).
Because the effect of exercise was seemingly limited to early (day 0 to day 5) postwounding, we hypothesized that exercise would reduce wound inflammation, which has been previously shown to be excessive in obese mice (5,12,31,37) and has been shown to be ameliorated by exercise in aged mice (21). Surprisingly, unlike our previous study in aged mice, exercise seemed not to affect wound site gene or protein expression of inflammatory cytokines TNF-α and IL-1β or of anti-inflammatory cytokine IL-10, nor wound site gene expression of chemokines MCP-1 and KC. This is to our knowledge the first report of an exercise effect on wound healing that is unrelated to alterations in wound site inflammation. This finding warrants further study.
There are some potential limitations to this study, primary of which is the use of only female mice. This was done to remain consistent with previous studies in aged mice (21) and in restraint-stressed mice (19) because these studies were major sources of inspiration for the study reported here. However, both previous studies used mouse strains different from the C57Bl/6J strain used in this study (Balb/cByJ and SKH-1 mice, respectively). The choice of C57Bl/6J mice was made because of their suitability as a model of DIO (36), and this strain of mouse is commonly used in obesity research. It is possible that exercise induces differential effects on wound tissue in these mouse strains, because previous research showed an exercise effect on healing rate that approached significance in young, lean Balb/cByJ mice (21), whereas we detected no difference in lean C57Bl/6J mice.
In female mice, estrogen plays a major role in protection from obesity-impaired wound healing (18). In the aged mouse study previously performed by our laboratory (21), the mice were postmenopausal, and thus, most of the estrogen effect was removed. However, our mice were much younger and premenopausal; thus, estrogen may be playing a protective role in reducing inflammation and speeding healing. This is partially supported by the finding that 6-wk-old male C57Bl/6J mice heal more slowly than their female counterparts (unpublished data). It is possible that an extension of this study to male mice might demonstrate an exercise effect on healing rate and wound inflammation similar to that seen in postmenopausal female mice in our previous aging study. This is a possibility that needs future study.
Several other potential mechanisms for the exercise effect on wound healing in obese mice require further investigation. Clotting and hemostasis represent the earliest responses to wounding (15), happening generally within 30 min after trauma. Exercise is known to positively influence hemostasis, possibly by increasing activity of coagulation factors as well as by increasing reactivity of platelets (24). Obesity and diabetes are linked to a dysregulated hemostasis through the maintenance of a procoagulant state (11). Both obesity and sedentary behavior have been shown to be risk factors for the development of inflammation and hemostatic imbalances (16). Thus, it is possible that exercise-induced alterations in hemostasis may explain the effects of exercise in HFD-fed mice seen in this study, and this potential mechanism should be evaluated. Although we saw no differences in PDGF in this study, it is likely that our first measure at 24 h after healing was taken too late to capture differences in hemostasis because this process generally occurs during the first 30–60 min of healing. The increase in PDGF expression seen at days 3 and 5 after wounding (compared with day 1) in this study likely reflects increased PDGF production by cells such as wound-associated macrophages rather than PDGF production resulting from platelet activation.
A second area that bears consideration for future research is the role of exercise in promoting myofibroblast migration and wound contraction in obese mice. Myofibroblasts are specialized cells that induce wound site contraction and are important in the healing of rodent tissues (15). Treadmill running has been previously shown to induce migration of myofibroblasts into the patellar tendon of female mice (34), a finding that lends some credence to this hypothesis. Recent evidence indicates that DIO causes a delay in myofibroblast differentiation in the wound site of HFD-fed rats (28). Thus, the effect on myofibroblast migration and differentiation may be an important mechanism by which exercise exerts its prohealing effect in obese, HFD-fed mice.
This study is the first to demonstrate an effect of exercise on cutaneous wound healing in a model of obesity. Although exercise sped healing rate early after wounding, contrary to previous aging studies, this was seemingly unrelated to alterations in wound inflammation. Future research in this area should focus on the role of early events after wounding (hemostasis and myofibroblast activity) as well as the differential roles of mouse strain and sex because these are thought to have a major effect on healing kinetics.
This project was partially supported by an American College of Sports Medicine Foundation Doctoral Student Research Grant and a Midwest American College of Sports Medicine Student Research Project Award to BDP.
The authors would like to acknowledge Dr. K. Todd Keylock, assistant professor at Bowling Green State University, who consulted on experimental design.
The authors declare no conflicts of interest.
The results of the present study do not constitute endorsement by the American College of Sports Medicine.
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