EF exposure design

A 10 kV/m EF intensity was selected based on previous studies demonstrating that fields of this magnitude can induce biologically relevant cellular and tissue responses without causing detectable thermal injury or necrosis (Ozcan et al. 2025; Akın et al. 2025). This exposure level approaches the range in which electromechanical perturbations of cellular structures may occur, enabling investigation of inflammatory signaling pathways, vascular reactivity, and extracellular matrix remodeling while minimizing the risk of irreversible electrothermal damage (Zhao et al. 2006; Okatan et al. 2018; Li et al. 2023). Despite accumulating evidence indicating that electric fields can influence cellular processes, comprehensive histopathological and molecular investigations of EF exposure in reproductive tissues remain limited (Kirson et al. 2004, 2007).

The selected EF intensity also falls within the upper range of environmental and occupational electric field exposures reported in proximity to high-voltage power systems and certain biomedical applications, including electrotherapy and pulsed-field technologies, which may transiently generate comparable local field strengths (WHO 2007; Yarmush et al. 2014). Therefore, the applied field strength represents a biologically effective and translationally relevant exposure scenario for evaluating potential tissue responses, particularly in hormonally responsive and highly vascularized reproductive organs such as the ovary and uterus.

EF exposure was performed using a custom-designed high-voltage parallel-plate system developed to produce a controlled and stable EF environment (Fig. 1). The apparatus consisted of two galvanized stainless-steel plates (50 cm × 100 cm, 1 mm thick, rounded corners) positioned horizontally at a fixed separation distance of 0.55 m and connected to a custom-built direct current (DC) high-voltage power supply (maximum 5.5 kVdc). Electrical connections were positioned centrally on each plate to minimize local distortion of the field distribution, and the plates were electrically isolated from the ground plane using on-conductive supports.

Fig. 1The alternative text for this image may have been generated using AI.

Schematic representation of potential cellular and tissue-level effects of high-intensity electric field exposure on the female reproductive tract. EF electric field, TNF-α tumor necrosis factor-alpha, VEGF vascular endothelial growth factor

Animal cages (26 cm × 43 cm × 15 cm, nonconductive material) were placed centrally between the plates. Cages were spaced approximately 50 cm apart to standardize cage positioning within the exposure region and to reduce potential inter-cage perturbation of the electric field (Fig. 2). Animals were not physically restrained and were allowed to move freely within the cages during exposure to minimize stress-related neuroendocrine and inflammatory confounders. Animal behavior was continuously monitored to prevent excessive contact with conductive components of the experimental setup.

Fig. 2The alternative text for this image may have been generated using AI.

Schematic diagram of the experimental apparatus and electrical configuration used to generate a controlled high-intensity electric field for in vivo exposure of female rats. The setup ensures stable EF application across the central exposure region while allowing unrestrained animal movement

Comprehensive spatial mapping of the EF distribution across the entire cage volume was not performed. Instead, EF intensity was verified at representative positions within the central exposure region using a calibrated electric field meter under surrogate loaded conditions designed to approximate the dielectric presence of animals. Measurements were obtained at three predefined points approximately 10–15 cm above the cage floor, corresponding to the typical vertical position of the animals’ bodies during exposure.

The measured EF values ranged between 9.6 and 10.4 kV/m (mean ± standard deviation (SD): 10.0 ± 0.3 kV/m), confirming that the nominal exposure intensity was maintained within a narrow range around the target value in the central exposure region. These measurements were performed using the same geometric configuration as that employed in the experiments (parallel plates with centrally positioned cages and unrestrained animals). Because the animals were free to move within the cages during exposure, the reported measurements should be interpreted as nominal field values characterizing the central exposure zone rather than a fully resolved spatial field distribution.

In addition, the electrode dimensions were substantially larger than the cage footprint, a configuration commonly used in parallel-plate exposure systems to minimize edge-related field distortions and to stabilize the electric field in the central exposure region. Under such conditions, the central zone between the electrodes is expected to exhibit relatively stable field characteristics compared with peripheral regions. Consequently, animals positioned within the middle portion of the apparatus were exposed to comparable nominal EF conditions during the experimental sessions.

Animals and experimental design

All experimental procedures were conducted in compliance with the Animal Research: Reporting of In Vivo Experiments (ARRIVE) 2.0 guidelines and internationally accepted ethical standards for animal research. The study protocol was approved by the Suleyman Demirel University Animal Experiments Local Ethics Committee (approval no. 18.09.2025–09/622). All procedures adhered to the principles of the 3Rs (replacement, reduction, and refinement) to minimize animal use and suffering. Animals were monitored daily for signs of distress, including reduced mobility, abnormal posture, respiratory difficulty, or body weight loss exceeding 20%. No adverse effects were observed throughout the study.

Sample size was determined using G*Power software (version 3.1.9.7) with the parameters α = 0.05, power (1 – β) = 0.90, and an effect size of 0.32, yielding a required sample size of eight animals per group. A total of forty adult female Wistar albino rats (12–14 weeks old; body weight 250–300 g) were obtained from the Experimental Animal Research Center of Suleyman Demirel University.

All female animals were age-matched and maintained under controlled environmental conditions to minimize physiological variability. Rats were housed in individually ventilated cages at a temperature of 22 °C ± 2 °C and relative humidity of 55–65%, under a 12-h light/12-h dark cycle, with ad libitum access to standard pellet chow and water. Before the initiation of the experimental procedures, all animals were acclimatized to the housing conditions for 1 week.

Animals were randomly allocated to the experimental groups to reduce potential allocation bias. Estrous cycle synchronization was not performed before EF exposure. However, all animals were selected within a similar age range and reproductive maturity stage to reduce potential variability. Because the primary aim of the study was to evaluate general in vivo tissue responses to short-term electric field exposure, random group allocation under standardized housing conditions was considered an appropriate approach to minimize potential cycle-related confounding effects.

Following acclimatization, animals were randomly assigned to five experimental groups (n = 8 per group) according to EF exposure duration: group I, control (0 min); group II, 1 min; group III, 5 min; group IV, 15 min; and group V, 30 min. Rats in the exposure groups were subjected to a single session of a nominal electric field intensity 10 kV/m generated by a custom-designed parallel-plate EF exposure system (Fig. 1). The field intensity was verified at predefined measurement points within the exposure setup. During exposure, animals were placed individually in a nonconductive cage positioned within the electrode system and were allowed to move freely without physical restraint. Control animals were handled under identical conditions but without activation of the electric field.

All animals remained within the experimental setup for a total duration of 30 min to ensure consistent environmental exposure across groups. In the control group, the device remained inactive for of the entire 30-min period. In the exposure groups, the electric field was activated for the designated exposure durations (1, 5, 15, or 30 min), after which the device was switched off while the animals remained in the setup for the remainder of the 30-min period.

After completion of the exposure protocol, all animals were monitored for vital signs, behavioral changes, and general well-being before euthanasia and tissue collection for histological and immunohistochemical analyses. Each animal underwent a single exposed session, after which the experimental protocol was completed.

Histopathological examinations

Euthanasia was performed by decapitation under deep anesthesia induced by an intraperitoneal injection of xylazine (10 mg/kg; Xylazinbio 2%, Bioveta, Ivanovice na Hané, Czech Republic) and ketamine (90 mg/kg; Ketalar, Pfizer, İstanbul, Türkiye), in accordance with institutional and internationally accepted animal welfare guidelines. Adequate anesthetic depth was confirmed by the absence of pedal withdrawal and corneal reflexes before decapitation.

A comprehensive necropsy was subsequently performed, and tissues of the female reproductive system, including the ovaries, uterus, and uterine tubes, were carefully excised. Tissue samples were immediately fixed in 10% neutral buffered formalin and processed using a fully automated tissue processor (Leica ASP300S, Leica Microsystems, Wetzlar, Germany). Following routine dehydration and clearing procedures, the samples were embedded in paraffin wax and sectioned at a thickness of 5 μm using a rotary microtome (Leica RM2155, Leica Microsystems, Wetzlar, Germany).

All sections were stained with hematoxylin and eosin (H&E) (Sigma-Aldrich, cat. no. HT110132) and examined under a light microscope. Histological evaluations were independently performed by two experienced histopathologists affiliated with separate external institutions, both of whom were blinded to the experimental group allocations to minimize observational bias. Histopathological alterations in the ovary, uterus, and uterine tubes were assessed using a semiquantitative scoring system designed to provide standardized and reproducible evaluation of tissue injury. The evaluated parameters included vascular changes (hyperemia and hemorrhage), interstitial edema, inflammatory cell infiltration, and structural tissue damage (including cellular degeneration and epithelial loss). These features were defined according to established histopathological criteria. Each parameter was graded on a four-point ordinal scale based on lesion extent and severity and distribution (0 = absent, 1 = mild, 2 = moderate, 3 = severe), as summarized in Table 1.

Table 1 Parameters to scoring and characterize EF-induced tissue damage for each organ (ovary, uterus and uterine tube)Semi-quantitative scoring system

Histopathological alterations in the ovary, uterus, and uterine tube tissues were assessed using a semi-quantitative scoring system by two board-certified histopathologists who were independently blinded to the experimental group assignments. For each specimen, an overall tissue damage score was calculated by summing the scores of all evaluated histopathological parameters, ensuring standardized and reproducible assessment of tissue injury.

For each section, at least five randomly selected microscopic fields were examined at ×400 magnification using a light microscope (Olympus CX43, Tokyo, Japan). All scoring was performed manually, and the results were independently verified by a second blinded observer to confirm interobserver consistency. This systematic and blinded approach enabled robust comparison of tissue injury severity among experimental groups and facilitated correlation with immunohistochemical marker expression.

Immunohistochemical examinations

Immunohistochemical staining was performed on selected sections of the female reproductive system to evaluate the expression of Osteonectin/SPARC (SPARC antibody, #DF6503; Affinity Bioscience, Canada), vascular endothelial growth factor (VEGF; VEGFA antibody, #AF5131; Affinity Bioscience, Canada), and tumor necrosis factor-alpha (TNF-α; anti-TNF-α recombinant antibody [RM1005], ab307164; Abcam, Cambridge, UK). After deparaffinization, rehydration, and heat-induced antigen retrieval, tissue sections were incubated with the respective primary antibodies at a dilution of 1:100 for 60 min at room temperature.

Immunodetection was carried out using a micro-polymer-based secondary antibody system (Mouse and Rabbit Specific HRP/DAB IHC Detection Kit; ab236466, Abcam, Cambridge, UK) according to the manufacturer’s instructions. This approach eliminates the need for a streptavidin–biotin complex and reduces background staining associated with endogenous biotin. Following incubation with the HRP-conjugated secondary reagent, immunoreactivity was visualized using diaminobenzidine (DAB) as the chromogen. Sections were then counterstained with Harris hematoxylin (Sigma-Aldrich, cat. no. HHS32), rinsed thoroughly under running tap water, and dehydrated through graded of ethanol series, cleared in xylene and coverslipped using Entellan mounting medium (catalogue no. 107961, Sigma-Aldrich, USA). Prepared slides were subsequently examined under a light microscope for semiquantitative analysis of marker expression.

Image analysis and quantification

Appropriate positive and negative controls were included for each immunohistochemical marker. Osteosarcoma tissue served as a positive control for osteonectin, VEGF, and TNF-α immunoreactivity, whereas negative controls were prepared by omitting the primary antibody. All histological and immunohistochemical evaluations were performed in a blinded manner.

Sections were examined using a light microscope (Olympus CX43, Tokyo, Japan), and images were captured with a calibrated digital camera (Olympus DP27) using Plan-Apochromat objectives at ×10 and ×40 magnifications under standardized optical settings.

Immunohistochemically stained sections were evaluated through a combination of manual microscopic assessment and digital image analysis. For each marker (osteonectin, TNF-α, and VEGF), the percentage of positively stained cells was determined by counting 100 cells in five randomly selected high-power fields (×400 magnification) per tissue section. Quantitative analysis was performed using ImageJ software (version 1.53a; National Institutes of Health, Bethesda, MD, USA), applying color deconvolution and thresholding algorithms to isolate DAB-positive signals. Immunoreactivity was expressed as the ratio of positively stained area to total field area.

All images were analyzed under identical acquisition and processing parameters to ensure comparability. Measurements were independently performed by two investigators blinded to group allocation, and the mean values were used for statistical analysis. Interobserver reliability was assessed using intraclass correlation coefficients (ICCs), and discrepancies exceeding 10% were jointly re-evaluated. This methodology provided reproducible and objective quantification of immunohistochemical marker expression across experimental groups.

Statistical analysis

Histopathological alterations were assessed using a semi-quantitative ordinal scoring system (0–3), were treated as nonparametric data. Intergroup comparisons were performed using the Kruskal–Wallis test, followed by Dunn’s multiple comparison post hoc test when a significant overall difference was detected.

Immunohistochemical (IHC) data, expressed as the percentage of positively stained cells, were treated as continuous variables. Normality of distribution was assessed using the Shapiro–Wilk test (p > 0.05), and group comparisons were subsequently conducted using one-way analysis of variance (ANOVA). When a significant main effect was detected, Tukey’s post hoc test was applied for multiple pairwise comparisons.

All statistical analyses were performed using GraphPad Prism software (GraphPad Software Inc., San Diego, CA, USA), and statistical significance was defined as p < 0.05.