Resistance Training Attenuates Oxidative Stress and Muscle Damage and Improves the Quality of Induced Skin Lesions in Rats

Abstract

Resistance training (RT) can help with injury recovery and the healing process. Still, high-intensity exercise can cause ischemia and reperfusion, resulting in exacerbated production of reactive oxygen species and oxidative stress. This study aimed to evaluate the effects of RT with progressive loads on markers of tissue damage and oxidative stress in rats subjected to skin lesions. Forty male Wistar rats were used, divided into four groups (n = 10): Control (CG): no intervention; Sedentary Injury (SHAM): subjected to injury, no training; Training + Injury 1 (G1): injury after one week of training; Training + Injury 2 (G2): injury followed by training. The protocol consisted of climbing a vertical ladder three times a week, 48 h apart, using progressive loads (50%, 65%, and 80%). After euthanasia, markers of tissue damage (CK, LDH, ALT, AST), oxidative stress (MDA/TBARS, SH, uric acid), and histological analysis of collagen deposition in the injured tissue were assessed. Groups G1 and G2 showed a significant increase (p < 0.0001) in CK, LDH, ALT, and AST levels compared to GC and SHAM. Oxidative stress markers, such as MDA and SH, were also elevated in the G1 and G2 groups (p < 0.0001). Uric acid concentrations increased significantly in the exercised groups compared to the controls (p < 0.0001). Histology revealed an inflammatory infiltrate and disorganized collagen fibers in the SHAM group, while G1 and G2 showed tissue with greater cellular maturity and organization. Although RT induced muscle damage and an increase in pro-oxidant markers, it also favored cellular organization and scar tissue quality.

1. Introduction

Resistance training (RT) is performed through contractions in muscle groups against an external resistance [1]. The manipulation of variables such as range of motion, volume, intensity, and rest promotes physiological and metabolic changes that improve physical fitness [2] Its benefits include improvements in cardiovascular physiology [2,3], glycemic profile, and insulin sensitivity [4], as well as muscle strengthening, increased bone density and cartilage preservation.

However, excessive exercise or high-intensity activities can cause tissue microlesions and promote the production of reactive oxygen species (ROS), leading to oxidative damage to lipid membranes and DNA [5]. This process can result in oxidative stress (OS), characterized by an imbalance between pro-oxidant and antioxidant systems, when antioxidant capacity is insufficient to neutralize free radicals [6]. Despite this, high-intensity exercise also stimulates metabolic adaptations, including an increase in endogenous antioxidant defenses [7]. Thus, RT tends to favor homeostasis since small amounts of ROS are essential for muscle strength, contributing to increased muscle mass and modulating redox reactions [1]. In addition, RT can positively influence the wound healing process, whose recovery is modulated by the balance of pro-oxidant and antioxidant systems [8]. Wounds, characterized by the rupture of skin tissue due to damage, compromise skin function and increase vulnerability to infection if the healing process is not effective [9,10].

The scientific literature describes different therapeutic strategies aimed at optimizing the wound healing process. Preclinical study shows that low-intensity aerobic training accelerates the wound-healing process [11]. However, the effects of RT on oxidative stress and tissue damage, as well as the mechanisms involved in the healing process, are unclear. Therefore, this study aims to evaluate the effects of resistance training on oxidative stress, muscle damage markers, and collagen deposition in rats with skin lesions, providing new perspectives on wound healing treatment.

2. Materials and Methods

2.1. Animals

Forty adults male Wistar rats (90 days old), weighing between 250 and 350 g, were selected from the Sector Animal Facility of the Intracellular Signaling Research Center (NUPESIN) at the Federal University of Sergipe. The animals were randomly housed in collective cages at an ambient temperature of 22 ± 3 °C, with a 12 h light–dark cycle.

2.2. Experimental Groups

The animals were adapted to the laboratory environment for a week before the start of the experimental protocol. After the adaptation period, the animals were weighed and randomly assigned. They were divided into four groups, n = 10: Control (CG): animals that did not undergo any intervention; Sedentary Injury (SHAM): subjected to the injury and did not undergo resistance training; Training + Injury Group 1 (G1): after the first week of training, the skin lesion was made, and then they underwent the training program for another three weeks, totaling 12 sessions; Training + Injury Group 2 (G2): Initially they were subjected to the skin lesion, and subsequently to a four-week training program also comprising 12 sessions. Prior to the experiment, the research was approved by the Animal Research Ethics Committee of the Federal University of Sergipe/UFS under protocol 08/2019.

2.3. Training Protocol

The training protocol was adapted from [12] and consisted of climbing a vertical ladder with a load apparatus attached to the base of the tail. They were initially trained for two weeks to climb the ladder. After the two-week adaptation period, the animals in groups G1 and G2 began the training protocol. The training was carried out three times a week, with 48 h intervals between training sessions, over four weeks, following a pyramidal training proposal, where the loads were 50%, 65%, and 80% of the Maximum Load Test (MCT), and 50% of the second week of training was adjusted according to performance in the second MCT. The initial series (initial two weeks) had a higher volume and lower load, and the final series (final two weeks) had a higher intensity and lower volume.

2.4. Maximum Load Test

This test was adapted from [13] to determine the training load. Three maximum load tests (MCTs) were carried out. The test protocol consisted of repetitions, with an initial load of 75% of body weight, adding 30 g to each climb, and two minutes of rest between repetitions. This procedure was repeated successively until the animal was exhausted, with the highest load being considered the final result of the MCT.

2.5. Induction of Skin Damage

The animals underwent epilation by manual traction in the dorsal costal region in an area of approximately 4 cm, with a Rhosse brand metal dermatological punch (ANVISA Reg. No. 80310620001, date of manufacture: 7 December 2018) containing a cutting blade on its lower edge; similar to that used in plastic surgery procedures. With this instrument, two skin fragments were excised in the center of the depilated area until the dorsal muscular fascia was exposed [14].

2.6. Preparation of Biological Material

Twenty-four hours after the last exercise session, the animals were anesthetized with ketamine/xylazine (75 mg/kg + 10 mg/kg i.p.) [15], and blood (±5 mL) was collected via cardiac puncture; then, they were euthanized by exsanguination under anesthesia. After collection, the blood was immediately centrifuged at 4000× g for 15 min at ±4 °C, and the supernatant was stored at ±−70 °C.

2.7. Tissue Damage Analysis

Muscle and liver damage markers were analyzed: cretin kinase (CK), lactate dehydrogenase (LDH), aspartate aminotransferase (AST), and alanine aminotransferase (ALT), using commercial kits (Labtest ^®, Santa Lagoa, Minas Gerais, Brazil) in which the plasma from each animal was homogenized in specific reagents at 37 °C and read using a spectrophotometer (BioespectroModel SP-22 UV/Visible, Minas Gerais, Uberaba, Brazil).

2.8. Determination of TBARS In Vivo

Gastrocnemius and liver tissues were harvested and processed according to the method described previsioly [16]; lipid oxidation was determined by measuring thiobarbituric acid-reactive substances (TBARS).

2.9. Determination of Total Sulfhydryls (Thiols)

Quantification of the antioxidant level of plasma and gastrocnemius and liver tissues was measured by determining the sulfhydryl groups, carried out according to the methodology described previsioly [17], in which aliquots of 50 μL of samples (blood and organs) were mixed in 1 mL of tris-EDTA buffer, pH 8.2.

2.10. Determination of Plasma Uric Acid

To determine uric acid (enzymatic UV uricase-peroxidase), the commercial kit (Labtest^®, BioespectroModel SP-22 UV/Visible, Uberaba, Minas Gerais, Brazil) was used, where (20 μL of plasma from each animal was homogenized in specific reagents at 37 ± 0.2 °C, and readings were taken using a spectrophotometer at a wavelength of 540 nm.

2.11. Histomorphological Analysis

Three histological sections stained in HE from each case were used to carry out the histomorphological analysis of the scar areas.

2.12. Analysis of the Average Thickness of the Epidermis

Histological images of the epidermal tissue on the scarred area (HE) were obtained using an Olympus CX31 microscope and an Olympus Camedia C-5060 digital camera, 5.1 megapixels (400 × magnification; 0.045 mm² analytical area). The epidermal thickness measurements for each sample were taken using ImageJ for Windows software, version 1.8.0 (National Institute of Health, Bethesda, ML, USA).

2.13. Analysis of Collagen Deposition

Analysis of collagen fiber deposition was carried out using two special stains. Masson’s trichrome, which highlights collagen fibers in blue, was used for descriptive analysis of the morphological pattern of collagenization. The density of collagen deposition was estimated by calculating the optical density of the image analysis system in an area of the dermis equivalent to 0.825 mm² (100× magnification) for each section. Photomicrographs of the selected areas were obtained using an Olympus Camedia C-5060 digital camera, 5.1 megapixels, coupled to an Olympus CX31 trinocular microscope. Using a scanning system (Olympus C-7070 WideZoom), the images were loaded onto a computer (Pentium 133 MHz) and processed using ImageJ for Windows software, version 1.8.0 (National Institute of Health, USA).

2.14. Statistics

Descriptive statistics were carried out using measures of central tendency, mean (X) ± Standard Deviation (SD). The Shapiro–Wilk test was used to check the normality of the variables, given the size of the sample. The ANOVA test (One Way) with Bonferroni Post Hoc was used to assess performance between the groups. Differences between means were considered significant when the “p” value was less than 0.05 (α = 5%). These analyses were carried out using the GraphPad Prism version 8.0 statistical program (GraphPad Software, San Diego, CA, USA). To check the size of the effect, values of low effect (0.1 and 0.24), medium effect (0.25 and 0.39), and high effect (greater than 0.40) were adopted [18].

3. Results

3.1. Tissue Damage Markers

There were statistically significant differences in plasma CK enzyme levels, with increased concentrations in groups G1 (628 ± 70.4 95% CI 563–693) and G2 (693 ± 87.3 95% CI 612–773) when compared to GC (326 ± 123 95% CI 231–420) and SHAM (360 ± 100 95% CI 283–437). No differences were identified between the SHAM and CG groups. In addition, significant differences were found between LDH concentrations, showing an increase in groups G1 (312 ± 44.4 95% CI 257–367) and G2 (278 ± 40.1 95% CI 236–320) when compared to GC (174 ± 16.5 95% CI 161–186) and SHAM (196 ± 24.8 95% CI 120–222) (Figure 1B). This indicates that the training induced muscle damage.

ALT concentrations increased in groups G1 (136 ± 15.6 95% CI 122–150) and G2 (126 ± 18.3 95% CI 109–143) when compared to the CG (57 ± 13.9 95% CI 44.1–69.9) and SHAM group (61.2 ± 13.3 95% CI 49–73.5) (Figure 2A). Similarly, AST enzyme concentrations were also high in groups G1 (97.3 ± 8.58 95% CI 88.3–106) and G2 (120 ± 23.2 95% CI 95.2–144) when compared to the CG (55.6 ± 10.2 95% CI 46.2–65.1) and SHAM (57.7 ± 2.72 95% CI 54.8–60.1) (Figure 2B). There were no significant differences between the trained groups (G1 and G2).

3.2. Oxidative Stress

Plasma malonaldehyde concentrations (Figure 3A) showed a significant increase in groups G1 (346 ± 54 CI 95% 296–36) and G2 (353 ± 51.6, CI 95% 306–401) when compared to GC (195 ± 55.9 CI 95% 114.3–247), p < 0.0001 and SHAM (263 ± 32.2 CI 95% 233–293), p < 0.05. The CG and SHAM groups had no significant differences (p > 0.05). There were also statistically significant differences in hepatic MDA concentrations, with an increase in the G1 (556 ± 83 CI 95% 469–643) and G2 (562 ± 64.4 CI 95% 494–629) groups when compared to the CG (310 ± 69.3 CI 95% 252–368), p < 0.0001, and the SHAM group (402 ± 74.9 CI 95% 332–471) (Figure 3B). MDA concentrations in muscle tissue were high in groups G1 (268 ± 41.5 CI 95% 234–303) and G2 (269 ± 18.4 CI 95% 253–284) when compared to GC (140 ± 46.7 CI 95% 90.8–189), with p < 0.0001, and SHAM (174 ± 49.7 CI 95% 128–220), with p < 0.001 (Figure 3C).

Analysis of the sulfhydryl (SH) groups showed an increase in plasma concentrations in groups G1 (242 ± 32.5 CI 95% 212–372) and G2 (226 ± 18.4 CI 95% 207–245) when compared to GC (142 ± 17.3 CI 95% 124–161) and the SHAM group (146 ± 31.4 CI 95% 113–179) (Figure 4A). Furthermore, when we assessed the levels of HS in liver tissue, we saw an increase in groups G1 (250 ± 65.2 CI 95% 182–319) and G2 (273 ± 55.7 CI 95% 215–332) when compared to the CG group (95 ± 47.3 CI 95% 45.3–145), with p < 0.0001. There was a significant increase in the G2 group when compared to SHAM (170 ± 24.3 95% CI 145–196) with p < 0.05 (Figure 4B).

In addition, there were statistically significant differences in muscle HS concentrations, showing an increase in groups G1 (327 ± 54.6 95% CI 259–394) and G2 (368 ± 69.5 95% CI 281–454) when compared to GC (122 ± 35.6 95% CI 89.3–155) and SHAM (155 ± 52.2 95% CI 107–204) (Figure 4C).

3.3. Plasma Uric Acid

Uric acid levels were elevated in the G1 trained group (4.04 ± 0.465 95% CI 3.61–4.47) and in the G2 group (4.28 ± 0.117 95% CI 4.17–4.39) when compared to the CG (3.12 ± 0.271 95% CI 2.91–3.33) and SHAM (3.21 ± 0.246 95% CI 3.03–3.4) groups (Figure 5).

3.4. Histological Analysis

In the GC group, no cytological or histological alterations were observed in the epidermal or dermal tissues (Figure 6A–C). The epidermis consisted of a thin, uniformly thick keratinized stratified squamous epithelium. Hair follicles and associated sebaceous glands displayed normal morphology. The dermis was composed of dense connective tissue with interlaced collagen fibers, a delicate capillary network, and predominantly spindle-shaped fibrocytes, consistent with normal skin architecture. In the other groups (SHAM, G1, and G2), the lesion area was easily identified through the following histopathological features: (i) thickening of the epidermal tissue; (ii) absent or underdeveloped skin appendages (e.g., hair follicles and sebaceous glands); (iii) mild hyperplasia of the granular layer; and (iv) greater cellularity of the normal papillary/reticular dermis compared to normal skin (Figure 6D,G,J).

In the SHAM group (Figure 6D–F), the connective tissue of the reticular dermis exhibited a mild residual lymphohistiocytic inflammatory infiltrate. Most of the stromal cells exhibited stellate morphology, with broad eosinophilic cytoplasm with precise boundaries (young fibroblasts/myofibroblasts) and nuclei with chromatin of variable density. The collagen fibers were thick and coarse and disposed in an irregular interlaced arrangement.

In groups G1 (Figure 6G–I) and G2 (Figure 6J–L), the inflammatory infiltrate was inconspicuous. However, in group G1, the cellularity was sparser, and the fibroblasts had a more mature appearance (fusiform morphology, with dark and elongated nuclei). On the other hand, group G2 presented more intense cellularity and consisted of voluminous and stellate fibroblasts, especially in the papillary dermis.

The quantitative analysis of the mean epidermal thickness is presented in Figure 7. The mean epidermal thickness in GC (37.3 ± 7.7 µm) was significantly lower than in the other groups (p < 0.001). Furthermore, the SHAM group exhibited the highest mean epidermal thickness (58.1 ± 10.6 µm), significantly thicker than in G1 (51.6 ± 8.4 µm; p < 0.01) and G2 (53.2 ± 9.3 µm; p < 0.05). However, no statistically significant difference was observed between groups G1 and G2 (p > 0.05).

Analysis of histological sections stained with Masson’s trichrome reveals that in all groups analyzed, intense collagen deposition was observed in the scar area after 21 days (Figure 8). Overall, the fibers were thick and coarse and exhibited a predominantly interlaced architectural arrangement. Collagen deposition was less dense in the papillary dermis than in the reticular dermis and was particularly loose around skin appendages (sebaceous glands and hair follicles). Nevertheless, the SHAM and G2 groups exhibited a predominance of stellate cells with more conspicuous cytoplasm, consistent with immature fibroblasts. In contrast, stromal cells in the G1 group exhibited a fusiform, elongated morphology with scant cytoplasm, indicating a more mature fibroblast population, similar to that observed in the GC group.

Collagen deposition was also analyzed in histological sections stained with picrosirius red, observed under polarized light (Figure 9). In all groups evaluated, two distinct birefringence patterns were observed: (i) shorter and thinner fibers, with greenish or greenish-yellow birefringence, interpreted as type III collagen, and (ii) longer and thicker fibers, with golden-yellow or orange birefringence, understood as type I collagen.

The GC group exhibited a collagen deposition pattern characterized by a dense, interlaced network of type I and type III collagen fibers. The birefringent fibrous matrix contained small, irregular non-birefringent areas corresponding to interfibrillar spaces, vascular structures, cutaneous appendages, or hypodermic components such as adipose tissue, skeletal muscle fibers, and peripheral nerves. Collagen deposition was homogeneously distributed throughout the papillary and reticular dermis, with a marked reduction in the hypodermic region.

In contrast, the SHAM, G1, and G2 groups displayed a distinct collagen organization, with thinner and more delicate fibers arranged predominantly in parallel, particularly within the papillary dermis. These groups also exhibited more evident interfibrillar spaces. Although type I collagen predominated in all groups, type III collagen was more abundant in the G1 and G2 groups, showing a pattern more similar to the GC group, than in the SHAM group. The quantitative analysis of the collagen deposition density in the four experimental groups is presented in Figure 10. The mean collagen deposition density in the GC group (56.6 ± 14.5%) was significantly higher than in the groups: SHAM (41.5 ± 15.2%; p < 0.05), G1 (38.4 ± 12.6%; p < 0.01) and G2 (43.5 ± 9.7%; p < 0.05). However, no significant difference was observed between the collagen density of the last three groups (p: 0.05).

4. Discussion

The present study investigated the effects of four weeks of resistance training on markers of tissue damage and oxidative stress in rats subjected to skin injury at different time points. The results showed that RT can enhance antioxidant capacity and, regardless of the time point at which the injury occurred, was effective in promoting wound healing. To assess tissue damage, markers of muscle (CK and LDH) and liver (ALT and AST) injury were used. Resistance training is known to cause microtrauma in skeletal muscle, increasing serum concentrations of CK and LDH. In addition, the enzymes ALT and AST are recognized as reliable markers of necrosis or hepatocellular injury [19].

In our study, a significant increase in CK and LDH concentrations was observed in groups G1 and G2 compared to the control (GC) and SHAM groups. These results indicate that, regardless of the time point of skin injury, resistance training induced muscle damage. Increased serum concentrations of these enzymes are widely associated with exercise-induced muscle microinjuries [20]. This phenomenon is related to muscle hypertrophy, which results from the mechanical stress generated by training. This process can cause structural changes in the sarcomere and the disruption of cytoskeletal proteins, leading to the leakage of cytosolic enzymes, such as CK and LDH, into the plasma [21,22]. High-intensity training can increase serum CK and LDH values, using them as a parameter to assess the intensity of physical effort and muscle injuries [23,24,25]. A study conducted by [26] also identified an increase in CK and LDH levels, indicating that RT is capable of inducing muscle injuries, corroborating the findings of this study.

In addition, increased concentrations of ALT and AST were observed in groups G1 and G2 compared to groups GC and SHAM. AST and ALT are enzymes that, when elevated in plasma, indicate peripheral changes resulting from exercise [27]. Thus, the data found in this study may indicate that pyramid resistance training induces peripheral metabolic changes in rats subjected to skin injuries at different time points. These findings corroborate the clinical study proposed by [28], in which he investigated the pattern of injuries during intense exercise in a hot and humid environment, specifically in the liver, in a model of exercise-induced heatstroke in rats, finding an increase in serum levels of these hepatic biomarkers.

Furthermore, significant differences were evidenced in plasma, muscle, and liver MDA, showing an increase in groups G1 and G2 when compared to groups GC and SHAM. MDA is the final product of lipid peroxidation and is used as a marker of oxidative stress [29]. This also demonstrates a state of oxidative stress during resistance training, which can result in oxidative damage and reduced performance [30].

The sources of ROS generation during physical exercise depend on the type, duration, and intensity of the exercise. However, most of the ROS produced is predominantly by the contraction of skeletal muscles during exercise [31]. In a clinical study proposed by [32] in a weightlifting training session, elevated values of this marker were also evidenced. Intense physical exercise increases lipid peroxidation, as well as the body’s antioxidant defenses [32,33].

In addition, our studies identified an increase in SH concentrations in plasma and liver and muscle tissues in groups G1 and G2 when compared to group GC, demonstrating an increase in antioxidant activity after resistance training. SH is present in the Cys residue of glutathione (GSH) and is responsible for its strong electron donation feature [34]. The molecules contained in the sulfhydryl side chain group act as antioxidants, stabilizing free radicals by receiving their unpaired electron; they are also called thiols [35].

Plasma uric acid concentrations were elevated in groups G1 and G2 when compared to the control and SHAM groups, demonstrating that regardless of the time of injury, resistance training promotes more excellent antioxidant activity. Uric acid is considered an important element of the antioxidant status of plasma. This, in turn, acts through the formation of stable coordination complexes with iron ions and prevents the Fenton reaction, reducing ROS and stabilizing other plasma antioxidants, such as thiols and ascorbic acid [36,37]. Uric acid is a final product of the xanthine oxidase reaction generated by reactive oxygen species during physical exercise. Thus, the increase in acid concentrations may be a physiological or adaptive response to exercise as a way of accelerating antioxidant mechanisms [38].

The data found in our study corroborate what was presented by [39], who found that eight weeks of intense training with rats in a swimming model increased uric acid concentrations when compared to moderate continuous training. In this way, providing an elevating action as a response to the body’s antioxidant defense against resistance training in a situation of skin injury.

Resistance exercise induces a transient increase in oxidative stress, which acts as a redox signal capable of activating adaptive pathways involved in tissue repair. This phenomenon, frequently described within the context of hormesis, promotes the activation of mechanisms that favor cutaneous healing and improve the quality of scar tissue [40]. Among these responses, key features include the modulation of pathways associated with angiogenesis [41], increased myokine secretion, and the release of exosomes with paracrine functions [42], in addition to the attenuation of the local inflammatory response, evidenced by the reduction in pro-inflammatory cytokines such as TNF-α, KC, and MCP-1. Together, these adaptations contribute to a tissue environment more favorable for repair, potentially accelerating wound closure, especially in conditions where the healing process is compromised.

Regarding the data on the wound closure process, as previously mentioned, the healing process is closely related to the inflammatory process, and its resolution is important for the outcome of the wound, which would be its closure [41]. However, this inflammation must not be too expressive since, when it is too prolonged, the healing process can be slow and may become a chronic wound [43].

In the present study, it was observed that the SHAM group presented inflammatory infiltrate, in addition to the presence of coarse collagen fibers and irregular arrangement. Meanwhile, the G1 and G2 groups did not present inflammatory infiltrate. They showed more mature and organized cellularity, which may indicate that the lower presence of inflammatory infiltrate was necessary for healing to occur more efficiently.

As the healing process progresses, the emphasis is on the deposition of collagen in the wound, and the scar remodeling process involves successive stages of synthesis, degradation, and orientation of collagen fibrils [44]. In the present study, the groups that performed physical exercise, G1 and G2, showed collagen deposition similar to the SHAM group, with no difference.

However, the type of collagen deposited during wound healing is a critical factor, as optimal scar quality depends on its similarity to healthy tissue [45]. Although type I collagen predominated in all groups, type III collagen was more abundant in the G1 and G2 groups, showing a pattern closer to that of the GC group. This finding suggests that the physically active groups developed scar tissue more similar to healthy skin, potentially conferring greater tissue elasticity [45]. In summary, the findings of the present study are consistent with the literature regarding exercise-induced muscle injury and oxidative stress, as reflected by increased CK, LDH, and MDA levels. Additionally, physical exercise was associated with elevated antioxidant markers, including thiol (-SH) content and uric acid concentration. Furthermore, histological analysis showed that the G1 and G2 groups developed scar tissue more closely resembling healthy skin, suggesting that physical exercise promoted improved scar quality regardless of the timing of wound induction.

5. Conclusions

In conclusion, resistance training with progressive loads induced muscle and liver stress and increased oxidative markers, such as MDA, irrespective of the timing of skin injury, while simultaneously enhancing antioxidant defenses, including thiol groups and uric acid. Additionally, physical exercise improved scar quality, yielding tissue more similar to healthy skin. Although limited to a single training model and injury method, these findings highlight the potential of resistance training to modulate redox balance and support wound healing, warranting further investigation using alternative protocols and biomarkers.