Prophylactic Effects of Radiofrequency Electromagnetic Field on Pulmonary Ischemia‐Reperfusion via HIF‐1α/eNOS Pathway and BCL2/BAX Signaling

ABSTRACT

This study aimed to investigate the vascular effects of a radiofrequency electromagnetic field (RF‐EMF) applied in the lung ischemia and reperfusion (IR) model on the hypoxia‐inducible factor 1 alpha (Hif‐1α)/endothelial nitric oxide synthase (eNOS) pathway and B cell lymphoma 2 (BCL2)/BCL‐2 associated X protein (BAX) signaling. Forty male Wistar rats were randomly divided into four groups, each consisting of 10 rats: Sham, IR, IR + RF‐EMF, and RF‐EMF. IR was applied to rats by 60 min of clamping hilus of left lungs and 60 min of reperfusion. Rats were kept in the RF‐EMF unit for 60 min with or without activation. After sacrification, lung tissues were excised for histopathological, immunohistochemical, biochemical, and genetic analyses. IR injury led to increased damage‐related emphysematous findings, significant hyperemia, and increased septal tissue thickness, as observed histopathologically, and immunoexpression levels of tumor necrosis factor‐alpha and caspase‐3. In addition, it was noted that the biochemical parameters total oxidant status, oxidative stress index, and genetic parameters Hif 1 α, eNOS, BAX increased, and BCL2 decreased due to IR damage. In the IR‐RF‐EMF group, improvement has been detected in all parameters. RF‐EMF applied in the IR model exerts antioxidant, antiapoptotic, and anti‐inflammatory effects on lung tissue damage through the Hif‐1α/eNOS pathway and BCL‐2/BAX signaling. The use of RF‐EMF in IR damage is promising, as models that examine the long‐term effects of RF‐EMF at different frequencies.

1. Introduction

Ischemia‐reperfusion (IR) injury is linked to several clinical situations, including significant trauma, vascular obstruction, shock, surgery, and infarction of the heart and brain [1]. It is described as a decrease in tissue blood flow or organ for some time and subsequent re‐blood flow, which results in both local and systemic distant organ damage through the rapid formation of free oxygen radicals and inflammatory mediators in the distal area of the obstruction area [2]. Hypoxia‐induced inflammation and oxidative stress secondary to obstruction in lung tissue can cause acute damage to the distal parenchymal tissue and limit lung expansion capacity due to tissue loss [3].

In addition, prolonged IR injury‐induced cell damage can lead to apoptosis, autophagy, and necrosis. There is an activation of some intracellular pathways in this mechanism of cellular damage, such as the vascularization‐related AKT/hypoxia‐inducible factor‐1 alpha (Hif‐1α)/endothelial nitric oxide synthase (eNOS) pathway and mitochondrial stress‐related B‐cell lymphoma‐2 (BCL‐2)/BCL2 Associated X Protein (BAX) signaling during ischemia and hypoxia‐induced inflammation and apoptosis [3, 4]. In particular, the nuclear factor kappa beta (NF‐κβ) signaling pathway increases Hif‐1α synthesis, and Hif‐1α is degraded by proteolysis in the presence of normoxic conditions, which provides the synthesis of hypoxia‐related factors from DNA under hypoxic conditions [4, 5, 6, 7] (Figure 1).

Figure 1.

The pathophysiological mechanism of IR‐induced lung damage and the potential effects of RF‐EMF.

It is known that the radiofrequency electromagnetic field (RF‐EMF) method increases nitric oxide (NO) synthesis by enhancing eNOS synthesis in the endothelial layer of the vascular structures in the areas where it is applied [8]. Asci et al. proved that the 30‐min RF‐EMF method that they used in an experimental wound model increased wound healing by increasing eNOS enzyme levels [9].

This study aimed to investigate the vascular effects and effects of RF‐EMF applied in the lung IR model on Hif‐1α/eNOS pathway and BCL2/BAX signaling.

2. Materials and Methods

2.1. Ethical Approval

The protocols for animal research were followed in all of the experimental studies we conducted for this study, Animal Research: Reporting in Vivo Experiments (ARRIVE) 2.0 at all stages of the experiment. The experimental protocol was approved by the local animal experiments ethics committee of Suleyman Demirel University with the number 26.01.2023/01‐123, and the experiments were carried out by this protocol. This study was supported by the Scientific Research Projects Coordination Unit of Suleyman Demirel University with project code TSG‐2023‐9010.

2.2. Reagents

Xylazin Bio 2% (Bioveta, Czech Republic) and Keta‐Control (Doğa İlaç, Turkey) were used to induce sedation and anesthesia.

2.3. RF‐EMF Setup

The circuit was fed with 12 Vdc to generate RF‐EMF and obtain radiations with two 27.12‐MHz RF antennas with an aimed electric field value of 10 V/m. An 0.8 W RF output was obtained from the 27.12‐MHz PCB antenna. Experiments were conducted by feeding and tuning circuits to achieve an aimed electric field intensity of 10 V/m in each cage [10].

The scientific literature emphasizes that such exposure mechanisms should be isolated from the external environment. The tests were conducted in a Faraday cage, an electromagnetically separated environment, where pre‐study measurements were taken. The operating frequency's shielding efficacy is expressed in terms of (80 dB). Figure 2 shows the RF‐EMF measuring setup.

Figure 2.

RF‐EMF application setup. RF‐EMF, radiofrequency electromagnetic field.

2.4. Surgical Procedure

The lung IR model was implemented as previously employed in other studies [11]. Briefly, after a 12‐h fasting period, experimental animals underwent intraperitoneal (ip) anesthesia with 90 mg/kg ketamine and 10 mg/kg xylazine. After a vertical incision was made in the neck, the trachea was exposed and tracheotomy was performed. A 16 G branula was used for tracheal intubation, and ventilation was provided with a mechanical ventilator. Following ventilation, the thoracic region was shaved, and a left thoracotomy incision was made from the sternum to the lower end of the scapula through the left 5th intercostal space. After identifying the left lung hilum and distal trachea, non‐traumatic vascular clamping was performed using a vascular clamp for 60 min, followed by 60 min of reperfusion.

2.5. Experimental Design

In the experimental study, 40 male Wistar Albino rats weighing between 300 and 350 grams were used. The rats were kept between 21°C and 22°C and subjected to 12 h of light and 12 h of darkness. An ad libitum feeding regimen was implemented. Euro type 4 cages were used, each housing ten rats. Forty rats were divided into four groups:

1‐Sham: Rats were kept for 60 min without activating the RF‐EMF unit. Afterward, a thoracotomy was performed, but an IR model was not created. Hilus was visualized.

2‐IR group: Rats were kept for 60 min without activating the RF‐EMF unit. Then, after the left thoracotomy, non‐traumatic vascular clamping was applied to the hilus, followed by 60 min of ischemia and then 60 min of reperfusion [12].

3‐RF‐EMF group: Rats were kept in the unit for 60 min by activating the RF‐EMF unit. Afterward, a thoracotomy was performed, but an IR model was not created [13, 14].

4‐IR + RF‐EMF group: Rats were kept in the RF‐EMF unit for 60 min. Subsequently, following left thoracotomy, a non‐traumatic vascular clamp was placed on the hilus, and 60 min of ischemia followed by 60 min of reperfusion were applied.

The induction of the IR model within the unit began during the last 10 min of the 60‐min RF‐EMF application, and ischemia was initiated exactly at the end of the 60‐min RF‐EMF application. Therefore, no time loss was incurred between the RF‐EMF application and ischemia. By placing each animal in the unit at different times to conduct the model, the standardization of the procedure was ensured.

Animals were killed upon confirmation of reperfusion. Surgical exsanguination was performed by collecting blood from the inferior vena cava through an abdominal incision. Blood samples and lung tissue samples were collected after euthanasia. Half of the lung tissue was preserved in 10% buffered formalin under conditions suitable for histopathological and immunohistochemical analysis. The remaining lung tissue samples were stored at −20°C for biochemical analysis and −80°C for genetic analysis.

2.6. Histopathological Evaluation

All lung tissue samples were taken in a routine histological tissue processing procedure using a fully automated tissue processing device (Leica ASP300S; Leica Microsystem, Nussloch, Germany), and then all samples were embedded in paraffin wax. After chilling the paraffin blocks, 5 µm sections were taken using a rotary microtome (Leica RM 2155; Leica Microsystem, Nussloch, Germany) from paraffin blocks. Hematoxylin‐Eosin (HE) staining and a coverslip were applied and examined under a light microscope. Histopathological lesions were scored on a scale of 0 to 3 based on their severity of hyperemia, edema, inflammatory cell infiltration, and epithelial cell loss.

2.7. Immunohistochemical Examination

Furthermore, two series of sections scraped from each paraffin block and drawn on poly‐l‐lysine‐coated slides were stained immunohistochemically for the expression of TNF‐α and Cas‐3 by the manufacturer's instructions using recombinant anti‐caspase‐3 antibody [EPR18297] (ab184787) and recombinant anti‐TNF alpha antibody [RM1005] (ab307164). Each primary antibody was diluted to 1/100. Sections were incubated with primary antibodies overnight. Then, streptavidin‐alkaline phosphatase conjugate and a biotinylated secondary antibody were used for immunohistochemistry. As a secondary antibody, we used the Mouse and Rabbit Specific HRP/AEC (ABC) Detection IHC Kit (ab93705). The chromogen employed was aminoethyl carbazole (AEC). Main and secondary antibodies were provided for all samples by Abcam (Cambridge, UK). An antigen dilution solution was used as a negative control rather than a primary antibody. A specialized pathologist from another university conducted each test on the blinded samples. The percentage of cells that were immunostained positively for each marker in 10 separate fields on each slide for all groups was calculated at an objective magnification of ×40. The output of the image analyzer was counted using the ImageJ program (National Institutes of Health, Bethesda, MD, version 1.48). The photos were separated into color channels and cropped, and any artifacts were eliminated before counting. After being chosen using a selection tool, cells inside the regions of interest were counted using the software's counting tool. Red color was used to identify positive staining and only cells with strong red staining were considered positive. Microphotos were taken using the Database Manual Cell Sens Life Science Imaging Software System (Olympus Co., Tokyo, Japan).

2.8. Biochemical Analysis

Lung tissue specimens were homogenized using an Ultra Turrax Janke & Kunkel homogenizer (IKA Werke, Germany) for oxidant‐antioxidant analysis. Utilizing commercial kits obtained from Rel Assay Diagnostics (Gaziantep, Turkey), spectrophotometric measurements of TAS and TOS were performed using the Beckman Coulter AU 5800 autoanalyzer (Beckman Coulter, USA). Calculation of the Oxidative Stress Index (OSI) was accomplished utilizing the formula OSI = [(TOS/TAS) × 100]. TAS and TOS analyses have been performed according to Erel's protocols [15]. Protein concentrations were measured using an autoanalyzer (Beckman Coulter, USA). The outcomes were quantified in units per gram of protein [16].

2.9. ReverseTranscription‐Polymerase Chain Reaction (RT‐qPCR)

Total RNA was obtained using the RNA isolation kit (Nepenthe, Turkey). The purity and quality of RNAs were measured by nanodrop (Shimadzu Ltd. Kyoto, Japan). cDNA was synthesized using 1 µg of RNA (Atlas Biotechnology, Turkey). Specific mRNA primer sequences were determined using the NCBI website (Table 1). Gene expression levels were determined using a real‐time PCR instrument (Biorad CFX Connect, California, USA) with 2X SYBR green master mix (Nepenthe, Turkey). The reaction mixture was prepared according to the manufacturer's instructions. For normalization, the GAPDH gene was used as housekeeping. Relative mRNA levels were calculated using the $2^{-∆∆C_{\mathrm{t}}}$ [15].

Table 1.

Primary sequences, product size, and accession numbers of genes.

Genes	Primary sequence	Product size	Accession number
GAPDH (HouseKeeping)	F: AGTGCCAGCCTCGTCTCATA	248 bp	NM_017008.4
	R: GATGGTGATGGGTTTCCCGT
Hif‐1α	F: GCAACTAGGAACCCGAACCA	251 bp	NM_024359.2
	R: TCGACGTTCGGAACTCATCC
eNOS	F: GGTTGACCAAGGCAAACCAC	247 bp	NM_021838.2
	R: CCTAATACCACAGCCGGAGG
BAX	F: CACGTCTGCGGGGAGTCAC	419 bp	NM_017059.2
	R: TAGAAAAGGGCAACCACCCG
BCL‐2	F: CATCTCATGCCAAGGGGGAA	284 bp	NM_016993.2
	R: TATCCCACTCGTAGCCCCTC

Abbreviations: BAX, BCL‐2 associated X protein; BCL‐2, B‐cell lymphoma 2; eNOS, endothelial nitric oxide synthase; F, forward; GAPDH, glyceraldehyde‐3‐phosphate dehydrogenase; Hif‐1α, hypoxia‐inducible factor 1 subunit alpha; R, reverse.

2.10. Statistical Analyses

For statistical analysis, immunohistochemical and genetic scores, tissue TAS, TOS, and OSI levels were compared between the groups. For the comparison between the groups, a one‐way ANOVA post hoc Tukey test with the GraphPad Prism program was used for statistical analysis, and p < 0.05 was considered significant.

3. Results

3.1. Histopathological Findings

Histopathology of the lungs of the sham and RF‐EMF groups revealed normal tissue arrangement. The IR group displayed emphysema, marked hyperemia, increased septal tissue thickness, and inflammatory cell infiltration. The IR + RF‐EMF group's adverse effects were lessened by RF‐EMF treatment (Figure 3).

Figure 3.

Histopathological appearance of three rats in each group and statistical analysis of lungs between the groups. (A) Normal lung histology in the sham group. (B) Increased septal tissue thickness (arrowheads) and inflammatory reaction (arrow) in lungs in the IR group. (C) Decreased inflammatory reaction and septal tissue thickness in the IR + RF‐EMF group, (D) Normal tissue architecture in the RF‐EMF group, HE, scale bars = 50 µm. IR, ischemia and reperfusion; RF‐EMF, radiofrequency electromagnetic field. Data expressed mean ± standard deviation (SD). One‐way ANOVA post hoc LSD tests were used.

3.2. Immunohistochemical Findings

Slides from the sham group's immunohistochemical staining revealed either no or very little Cas‐3 and TNF‐α expression. Cas‐3 and TNF‐α expression significantly enhanced in the IR group compared with the sham group (p ≤ 0.001; respectively). After RF‐EMF treatment, the expression of TNF‐α and Cas‐3 were also downregulated significantly (p ≤ 0.001, respectively). Using markers, the IR group displayed expressions similar to those of the control group. Inflammatory cells, alveolar macrophages, and alveolar epithelial cells all often exhibited expressions. The results of a statistical analysis of immunohistochemistry expressions are shown in Figure 4.

Figure 4.

Immunohistochemical appearance of Cas‐3 (upper row) and TNF‐ α (below row) expressions and the statistical analysis between the groups (A) Negative expression in sham group. (B) A marked increase in expression (arrows) in the IR group. (C) Decreased expression (arrow) in IR + RF‐EMF group. (D) No immunoreaction in the RF‐EMF group. Streptavidin biotin peroxidase method, scale bars = 50 µm. Data expressed mean ± standard deviation (SD). One‐way ANOVA post hoc LSD tests were used. IR, ischemia and reperfusion; RF‐EMF, radiofrequency electromagnetic field. Data expressed mean ± standard deviation (SD). One‐way ANOVA post hoc LSD tests were used, *p < 0.05, **p ≤ 0.01 ***p ≤ 0.001.

3.3. Biochemical Examination

TOS and OSI levels were found to be increased in the IR group of this study, and TAS levels were found to be significantly decreased compared with the SHAM and only RF‐EMF applied group (p ≤ 0.001 for all). The TOS and OSI levels significantly decreased (p = 0.004 and p ≤ 0.001, respectively). TAS levels decreased (p ≤ 0.001) in the IR + RF‐EMF applied group compared to the IR and IR + RF‐EMF groups. In the RF‐EMF group, TOS levels increased significantly compared with the SHAM group (p = 0.046) and RF‐EMF group (p = 0.024). Also, a significant increase was observed in IR + RF‐EMF compared to the RF‐EMF group (p = 0.020) (Figure 5).

Figure 5.

Graphs showing TOS, TAS, and OSI levels and statistical analysis of lungs Data presented as the mean ± standard deviation. The oxidative stress markers were compared between groups using one‐way ANOVA (post hoc LSD test). IR, ischemia and reperfusion; OSI, oxidative stress index; RF‐EMF, radiofrequency electromagnetic field; TAS, total antioxidant status; TOS, total oxidant status. Data expressed mean ± standard deviation (SD). One‐way ANOVA post hoc LSD tests were used, *p < 0.05, **p ≤ 0.01 ***p ≤ 0.001.

3.4. Genetic Examination

Hif‐1α and eNOS expression was significantly increased in the IR group compared with the sham and RF‐EMF groups and significantly decreased in the IR + RF‐EMF group compared with the IR group (p < 0.001 for all).

BAX expression was significantly increased in the IR group compared with the sham and RF‐EMF groups (p = 0.001 for both) and significantly decreased in the IR + RF‐EMF group compared with the IR group (p = 0.001). Conversely, BCL‐2 expression was found to be significantly reduced in the IR group compared with the sham and RF‐EMF (p = 0.001 for both) groups and significantly increased in the IR + RF‐EMF group compared with the IR group (p = 0.004) (Figure 6).

Figure 6.

Graphs showing mRNA relative fold changes of BCL‐2, BAX, Hif‐1α, eNOS genes Values are presented as means ± SD. Statistical analysis was performed with one‐way ANOVA and post hoc LSD test. BAX, BCL‐2 associated X protein, BCL‐2, B‐cell lymphoma 2; eNOS, Endothelial nitric oxide synthase; Hif‐1α, Hypoxia‐inducible factor‐1 alpha; IR, Ischemia and Reperfusion; RF‐EMF, radiofrequency electromagnetic field. Data expressed mean ± standard deviation (SD). One‐way ANOVA post hoc LSD tests were used, *p < 0.05, **p ≤ 0.01 ***p ≤ 0.001.

4. Discussion

IR is defined as a decrease in blood flow to a tissue or organ for a period of time and subsequent re‐blood flow. Many studies have used different methods to prevent organ damage due to IR. The functional consequences of depriving tissue of blood flow, (i.e., oxygen), have been known for years. Recently, it has been shown that reperfusion, the restoration of blood flow following ischemia, may make ischemic organs more prone to cellular necrosis, thereby restricting functional return. Ultimately, prolonged IR‐induced cell damage can lead to apoptosis, autophagy, and necrosis [1, 2, 11]. The inflammatory response following ischemia and reperfusion in the lungs is shaped by a series of events, including oxygen deprivation, production of free radicals, cellular damage, endothelial dysfunction, activation of inflammatory cells, and subsequent edema. These processes can affect lung function and lead to serious long‐term complications [2, 11]. In this study, significant histopathological findings were observed in all rats in the ischemia‐reperfusion‐induced groups. The findings were found to be consistent with previous studies.

There is not enough data regarding the mechanisms through which RF‐EMF exerts its effect on hypoxia‐related damage in the lungs. Some intracellular mechanisms are activated, such as the Hif‐1α pathway, which plays a role in damage caused by hypoxia and the resulting inflammation and apoptosis, and BCL2/BAX, essential indicators of mitochondrial damage [3]. While increased Hif‐1α is degraded by proteolysis in the presence of normoxic conditions; it provides the synthesis of hypoxia‐related factors from DNA under hypoxic conditions [4, 5, 6]. Again, eNOS is one of the enzymes that shows its properties, such as regulating vascular tone and inflammatory response through this pathway [7]. By enhancing levels of interleukin 6 (IL‐6) levels, increased Hif‐1α affects eNOS‐mediated permeability and vascular tone [5, 8] In this study, the significant decrease in eNOS and Hif‐1α levels in the IR + RF‐EMF group compared with the IR group suggests that RF‐EMF exerts its preventive effect on lung tissue damage in acute injury. When looking at the literature, publications show that a 30‐min RF application increases eNOS activity without IL‐6 increasing levels. Still, in these studies, the application of RF is repeated and for a more extended period daily. In this study, the 60‐min RF‐EMF application was planned to be performed simultaneously. In addition, the suppression of eNOS activity in this study, which increased under normal conditions of the inflammatory state, gives the impression that it may be secondary to the decrease in IL‐6 levels, which causes the activation of the pathway. It is thought that reducing.

IL6 levels, an indicator of acute inflammation, by other anti‐inflammatory mechanisms may cause this situation. This can be interpreted as the RF‐EMF method exhibiting vasodilatory and other anti‐inflammatory effects.

In histopathological examinations, the significant decrease in hyperemia, septal tissue thickness, and inflammatory cell infiltration detected in the IR + RF‐EMF group, also supports the stated anti‐inflammatory activity. In the damage model used in this study, regressing the acute inflammatory response in lung tissue and secondary septal thickening is very important in terms of preserving lung functions and minimizing damage.

TNF‐α and Cas‐3, whose expressions are examined immunohistochemically, are pleiotropic cytokines that affect inflammation and apoptosis processes and play a key role in various lung pathologies [17]. TNF‐α initiates the apoptotic process, while Cas‐3 initiates apoptosis by taking part in the last common part of the extrinsic (death ligand) and intrinsic (mitochondrial) pathways of the apoptotic process. In this study, Cas‐3 and TNF‐α expressions were found to be decreased in the immunohistochemical staining in lung tissues in the group receiving RF‐EMF treatment, and it was shown that RF‐EMF had effects on reducing lung damage caused by ischemia‐reperfusion related apoptosis and inflammation. The fact that the RF‐EMF method can reduce TNF‐α levels gives the impression that it can be used for treatment purposes in many other acute events.

Apoptosis can occur via both the extrinsic pathway and the mitochondrial internal pathway. In both pathways, cell death occurs with Cas‐3 activation, and in the mitochondrial pathway, Cas‐3 activation is carried out by caspase‐9 stimulated by cytochrome‐c, which is released by increased mitochondrial membrane permeability as a result of damage. This mitochondrial pathway is increased by BAX, which disrupts mitochondrial membrane stabilization by cytochrome‐c activating caspases, and is inhibited by the BCL‐2 protein family, which suppresses BAX gene expression [17, 18]. Recent studies have shown that many aspects of mitochondrial biology, including mitochondrial dynamics, are critical determinants of the occurrence and progression of lung disease. In this model, the increase in BCL‐2 expression and the decrease in BAX expression in the RF‐EMF group suggest that it affects apoptosis especially through the mitochondrial pathway. We think that especially the high‐density vascular structure of lung tissue and the large number of mitochondria contained in lung cells strengthen these effects. In addition, when we look at the findings, the RF‐EMF method applied in the treatment can also show its effect through this pathway.

As a result of the above genetic analyses, it can be said that the effects of RF‐EMF on HIF‐1α/eNOS and BCL2/BAX pathways develop secondary to reducing inflammation or oxidative stress. It seems likely that RF‐EMF, which can normally activate eNOS, does not cause this in the 60‐min application here or that it decreases during this period. The reason for this decrease may be secondary to either the depletion of the precursor substance in synthesis, l‐Arginine, or other anti‐inflammatory mechanisms such as the suppression of TNF‐α it creates. It is also known that reduced damage may cause lower Hif‐1α levels and that reduced TNF‐α expressions may prevent apoptosis, and it is necessary to focus on this anti‐TNF‐α behavior of RF‐EMF in subsequent studies.

The anti‐inflammatory and antiapoptotic effects of RF‐EMF result in positive effects in terms of oxidant stress and antioxidant enzyme activities. In the study conducted by Moskowits et al. examining the therapeutic effects of levetiracetam and magnetic field in experimental spinal cord injury, combined levetiracetam, and magnetic field therapy may be effective in correcting damage‐induced changes in lipid structure in spinal cord trauma, preventing oxidative stress, regulating lipid composition, that is, preventing secondary damage. It has been shown that it may be a promising treatment alternative with further research [18, 19, 20, 21]. It has been shown that RF‐EMF and low‐frequency magnetic fields can increase oxidative stress. Signs of oxidative stress have been observed in the brain, testes, liver and kidneys, especially in rats and mice [20]. The application of electromagnetic fields in wound healing is also the application of PRFE at a carrier frequency of 27.12 MHz. Such applications have effects on inflammation and wound healing [21]. In this study, the decrease in TOS and OSI levels detected in the IR + RF‐EMF group in the biochemical analysis and the significant increase in TAS levels compared with the sham group also support the antioxidant activity of RF‐EMF. The fact that these results are parallel to the inflammatory response can be interpreted as preventing oxidative stress in regressing damage and thus reducing inflammation.

As a result, the RF‐EMF applied in the IR model exerts its antioxidant, antiapoptotic, and anti‐inflammatory effects on lung tissue damage through the Hif‐1α/eNOS signaling pathway and BCL2‐BAX signaling. The use of RF‐EMF in IR damage will be promising with models that examine the long‐term effects of RF‐EMF at different frequencies. Despite these demonstrated positive effects of RF‐EMF, its effectiveness should be evaluated in more clinical and experimental studies before entering routine clinical practice in patients with IR injury.

Author Contributions

All authors contributed to the study's design, sample collection, analysis, and data interpretation.

Ethics Statement

The experimental protocol was approved by the local animal experiments ethics committee of Suleyman Demirel University with the number 26.01.2023/01‐123.

Conflicts of Interest

Dr. Akin and the co‐authors have no conflicts of interest to declare in association with this study.