Unless otherwise mentioned, the culture media, ascorbic acid, anethole, eugenol and other chemicals used in the present experiments were purchased from Sigma-Aldrich (USA).

All procedures were approved by the Ethics and Animal Use Committee of State University of Ceara, Brazil (Protocol N. 05498214/2019). A total of eight ovarian pairs from eight adult crossbred goats (1–3 years old) were collected at a local slaughterhouse. The surrounding fat tissue and ligaments were removed, and the ovaries were washed in 70% alcohol, followed by two washes in minimum essential medium (MEM) plus HEPES (MEM-HEPES). The ovaries were placed into tubes containing 15 mL of MEM-HEPES, supplemented with penicillin (100 μg/mL) and streptomycin (100 μg/mL) and then transported to the laboratory at 4°C within 1 h.

The basic medium used was α-MEM⁺ (M5650, pH 7.2 −7.4) supplemented with 1.25 mg/mL bovine serum albumin (BSA), ITS (10 μg/mL insulin, 5.5 μg/mL transferrin, 5 ng/mL selenium), 2 mM glutamine, and 2 mM hypoxanthine, which was also named α-MEM⁺. Incubation was carried out at 38.5 °C in 7.5% CO₂ in air for 1 or 7 days. Fresh media were prepared immediately before use and incubated for 1 h prior to use, with 1 mL in each well. The fragments were individually cultured in a 24-well plate containing 1 mL α-MEM⁺ culture medium. The culture medium was replaced with fresh medium every 2 days. Each treatment was replicated five times.

In the laboratory, the ovarian cortex of five ovarian pairs was removed with a sterile scalpel and divided into 14 fragments (3 x 3 x 1 mm). Two fragments were randomly taken and immediately fixed for the non-cultured control treatment as described below. The remaining fragments were randomly distributed (two fragments per treatment) into the six treatments as follows: α-MEM+ (cultured control treatment) either or not supplemented with ascorbic acid 50 μg/mL (AA treatment), anethole 300 μg/mL (ANE 300 treatment), or eugenol at three different concentrations of 10, 20 and 40 μM, which were termed as EUG 10, EUG 20, EUG 40 treatments, respectively. The concentration of ascorbic acid (9), anethole (10), and eugenol (10 μM/mL, 20 μM /mL or 40 μM/mL; 25) were chosen based on previous studies. At the end of each culture period (days 1 and 7), ovarian fragments from each treatment were evaluated for morphology and development (follicular class and growth of follicle and oocyte diameters). In addition, spent culture media at the end of 24 h of culture at day 1 and at day 7 of culture (after medium replacement at day 6) were stored at −80°C for ROS analysis and quantification of thiol groups.

Based on the results of the morphologic analysis, the viability, immunofluorescence detection of histone H3 methylated and calreticulin assays were performed only in the treatments with the concentration of eugenol that maintained follicular morphology and increased follicular activation and diameter. For this, three ovarian pairs collected at a slaughterhouse were prepared in the laboratory as aforementioned and then cut into fragments. One fragment was immediately processed for follicle isolation (fresh control) or fixed, and the remaining fragments were cultured for 7 d in basic culture medium (α-MEM⁺) in the absence or presence AA, ANE 300 or in the eugenol treatment that provided the best outcome, i.e., a significantly higher percentage of morphologically normal follicles at day 7 of culture when compared with the other treatments.

Tissues from all treatments (fresh control or cultured for 1 and 7 days) were fixed in Carnoy's solution for 12 h and were processed for histology. The follicle stage and survival were assessed microscopically on serial sections. Coded anonymized slides were then examined by microscopy (Nikon, Sendai, Japan) at 400 × magnification.

The developmental stages of follicles have been defined as primordial (oocyte surrounded by a few flattened granulosa cells) or developing, i.e., intermediate (oocyte surrounded by flattened and at least one cuboidal granulosa cell), primary (oocyte surrounded by a complete layer of cuboidal granulosa cells), or secondary (oocyte surrounded by two or more complete layers of cuboidal granulosa cells). These follicles were classified as morphologically normal (follicles containing an intact oocyte and granulosa cells well-organized in layers without a pyknotic nucleus) and degenerated follicles (oocyte with a pyknotic nucleus, retracted cytoplasm, or disorganized granulosa cells detached from the basement membrane), as described Silva et al. (26).

Overall, 150 follicles were evaluated for each treatment (30 follicles per treatment in one repetition x 5 repetitions). To calculate follicular activation and growth, only morphologically normal follicles with a visible oocyte nucleus were recorded, and the proportion of primordial and growing follicles was calculated at day 0 and after day 7 of the culture in all treatments. In addition, from the basement membrane, the diameter of each normal oocyte and follicle were measured using a light microscope fitted with an eyepiece micrometer (Zeiss, Cologne, Germany) under 400 × magnification.

Each ovarian pair of ovaries (n = 3) collected at a slaughterhouse was cut into fragments. Two fragments were immediately processed for follicle isolation (non-cultured tissue), and the remaining fragments were cultured for 7 days in basic culture medium (α-MEM+), and ANE 300 or in the treatment group (40 μM/mL eugenol treatment) that provided the best outcome, i.e., a significantly higher percentage of morphologically normal follicles at day 7 of culture when compared with the other treatments. Goat preantral follicles were isolated from ovarian fragments using the mechanical method and stained with trypan blue as previously described (27). The viability of the isolated preantral follicles was assessed by the trypan blue dye exclusion test. The follicles were examined with an inverted microscope (Nikon, Tokyo, Japan) and classified as nonviable or viable if they were positively or negatively stained with trypan blue, respectively.

The ovarian fragments were fixed with 4% paraformaldehyde in PBS (pH 7.2) for 4 h and later dehydrated, diaphanized, and included in paraffin. Briefly, tissue sections (5 μm) were mounted on Superfrost Plus slides (KnittelGlass, Bielefeld, Germany), dewaxed with Citrisolve (Fisher Scientific, Ottawa, Canada), and rehydrated with increasing ethanol concentration. Antigen recovery was performed by incubating tissue sections in 0.01 M sodium citrate buffer (pH 6) for 5 min under a pressure cooker. After cooling to 37°C, the tissue sections were washed in PBS and blocked for 1 h at room temperature using PBS containing 1% (w/v) BSA. After antigenic recovery, the slides were incubated overnight at 4°C with polyclonal primary antibodies H3K4me3 (Abcam ab8580) and calreticulin (Abcam ab2907). The negative control was obtained by omitting the primary antibodies. Subsequently, the slides were incubated with secondary IgG antibody labeled (Alexa Fluor^® 488; Abcam ab150077) for 1 h at room temperature and mounted with Fluoroshield Mounting Medium With DAPI^® (Abcam ab104139). Finally, the slides were viewed using a fluorescence microscope (Nikon, Tokyo, Japan) with 20 × magnification. The images were individually saved using a monochrome digital camera to measure fluorescence intensities (FIs). Exposure gains and rates were consistent across samples. The FIs were quantified only in the limited area of normal (n = 230) preantral follicles using the software Image J 1.4.7 (NIH, Bethesda, MD, US). All evaluations were performed by a single operator.

Total RNA was isolated from frozen-thawed samples (non-cultured and cultured) using the Trizol^® reagent, according to the manufacturer's recommendations (Invitrogen, Carlsbad, CA, USA). Isolated RNA was further purified (PureLink™ RNA Mini Kit; Ambion^®, Carlsbad, CA, USA), treated with DNase I, and RNA samples were quantified with a spectrophotometer. Isolated RNA served as a template to synthesize the complementary DNA using the Superscript™ II RNase H-reverse Transcriptase (Invitrogen, São Paulo, SP, Brazil), followed by quantitative polymerase chain reaction (qPCR) in a final volume of 20 μL containing 1 μL of each complementary DNA, one x Power SYBR Green PCR Master Mix (10 μL), 7.4 μL of ultra-purewater, and 0.5 μL (final concentration) of both sense and antisense primers. Primer pairs (sense and antisense) and sequences for target genes are shown in Table 1. Peptidylprolyl isomerase A (PPIA) gene was selected as the endogenous control for normalizing gene expression among samples. The PCR conditions consisted of an initial step for denaturation and polymerase activation for 15 min at 94°C, followed by 40 cycles of 15 s at 94°C, 30 s at 60°C, and 45 s at 72°C. A final extension was performed for 10 min at 72°C. The specificity of each primer set was tested with melting curves that were carried out between 60°C and 95°C. All amplifications were carried out in a Bio-Rad iQ5 (Bio-Rad, Hercules, CA). Data were analyzed using the efficiency-corrected Delta-Delta-Ct method (28). The fold-change values of the genes of interest were normalized using the geometric mean of the fold-change values of a housekeeping gene.

For the quantification of ROS and thiol groups, 60 μL of each spent culture medium was centrifuged (720 × g, 4°C) for 10 min to eliminate cells and cell debris before quantification. The supernatant was used for the analyses.

The concentration of ROS was assessed through the Amplex Red/HRP (Molecular Probes) method which detects the clotting of fluorescent oxidized substances in the reacting mixture. 10 μL of the medium were incubated with SOD (SOD 100 IU/mL; Sigma-Aldrich, St. Louis, MO, USA), the horseradish peroxidase (PRF 0.5 U/ml; Sigma-Aldrich, St. Louis, MO, USA) and AmplexRed (50 M; Sigma-Aldrich, St. Louis, MO, USA). The fluorescence was measured through a microdishes reader (Victor3, Perkin Elmer) at 595 nm of excitation. The levels of ROS were expressed in micromoles of hydrogen peroxide (H2O2).

For the analysis of thiol groups, 60 μL the supernatant of the centrifuged medium was added mixed with 930 μL of 10 mM Tris-HCl, 0.9% NaCl (w/v), pH 7.4 buffer and methanol with an ultra-turrax homogenizer (Jamke & Kunkel IKA Labortechnik, Germany). The quantification of free thiol groups in the medium was performed using 10 μL of dithiolnitrobenzoic acid (DTNB) in a spectrophotometer (Hitachi U-3300). Thiol groups react with DTNB releasing the disulfide joined to the molecule, leading to the formation of 2-nitro-5-benzoato (NTB-) which ionizes the c in aqueous medium with neutral or alkaline pH. The NTB2- is quantified with spectrophotometer at a wavelength of 412 nm.

Antioxidant activity was measured in 96-well flat-bottom plates using BIOTEK Elisa reader (model ELX 800, software “Gen5 V2.04.11”). For the ABTS method, ABTS⁺ solution (7 mM, 5 mL) was mixed with 88 μL of potassium persulfate (140 mM). The mixture was stirred and kept in the dark at room temperature for 16 h. Then, 1 mL of this solution was added to 99 mL of ethanol. In 96-well plates, 300 μL of ABTS solution and 3 μL of the culture medium (sample) were used per well. For the DPPH analysis, the following solutions were used per well: 180 μL of DPPH (2,2-diphenyl-1-picrylhydrazyl) methanolic solution and 20 μL of the culture medium (sample).

The dilutions of samples and positive standards used in the quantitative microplate evaluations, starting from a stock solution with a concentration of 20 mg/mL were: 200 μg/mL, 100 μg/mL, 50 μg/mL, 25 μg/mL, 12.5 μg/mL, 6.25 μg/mL, 3.12 μg/mL, 1.56 μg/mL, and 0.78 μg/mL.

The absorbance for the ABTS method was measured at 630 nm up to a total of 10 min of incubation. As a negative standard, all solutions except the sample were used. For DPPH, the absorbance was measured at 490 nm for a total of 60 min of incubation.

The results were expressed as a percentage of inhibition and calculated according to the following formula:

Statistical analysis was carried out using Sigma Plot 11 (Systat Software Inc., USA). The normality (Shapiro-Wilk test) and homogeneity of variance (Levene's test) were initially evaluated. All parameters were analyzed with one-way ANOVA followed by Kruskal-Wallis and Mann-Whitney tests, when appropriate. Data are reported as mean (±SEM). Values with P < 0.05 were considered statistically significant.

A total of 2,632 preantral follicles were analyzed by classical histology. Representative images of the follicles after 7 days of culture are shown in Figure 1. The percentages of morphologically normal preantral follicles in each follicular category (primordial, intermediate and primary follicles) in the non-cultured control and after 1 or 7 days of in-vitro culture in all treatments are shown in Table 2. After 1-day culture, a significant reduction in the percentage of normal follicles was observed when compared to the control, except for primordial follicles cultured in the presence of EUG 20 or EUG 40, intermediate follicles cultured in the presence of ANE 300 or EUG 40 and growing follicles cultured in the presence of EUG 40. A significant decrease in the percentage of normal primary follicles was observed only when culture was performed for 1-day in the presence of ANE 300. On day-7 of in-vitro culture, except for the culture in the presence of ANE 300, a significant decrease in the percentage of normal primordial follicles was observed, whereas a decrease in the percentage of normal primary follicles was observed only when culture was performed in the presence of EUG 10. Although in-vitro culture decreased the morphology of growing follicles, the significantly highest percentages of normal growing follicles were observed after culture in the presence of EUG 40 and ANE 300. Moreover, a lower rate of normal primordial follicles was observed at day 7 when compared with day 1 when culture was performed in the presence of EUG, regardless of the tested concentration.

The proportions of morphologically normal primordial (Figure 2A) and growing follicles (intermediate + primary—Figure 2B) as well as degenerated primordial (Figure 2C) and growing (Figure 2D) follicles were determined in non-cultured ovarian tissue or in tissues cultured for 1 or 7 days. Regardless of the cultured treatment, from day 1 to day 7, there was a significant reduction in the percentage of normal primordial follicles, with a concomitant increase in the percentage of normally growing follicles. On days 1 and 7 of in-vitro culture, a decrease in the percentage of normal preantral follicles was observed in all treatments, except on day 1, when tissue was cultured in the presence of EUG 40. On day 7, medium supplementation with EUG 40 resulted in a decreased rate of normal primordial follicles together with the highest percentages of normally developing follicles.

The follicular and oocyte diameters in the non-cultured control treatment and after 1 or 7 days of culture are shown in Tables 3, 4, respectively. When compared to the non-cultured control, smaller follicular diameters were obtained when cultured in the presence of MEM alone (primordial—day 7, intermediate—days 1 and 7) or supplemented with AA (primordial and primary—day 7, and intermediate—days 1 and 7), ANE 300 and EUG 10 (primordial, intermediate and primary—day 7), EUG 20 (primordial, intermediate and primary—days 1 and 7) and EUG 40 (primordial—day 1). Moreover, the intermediate follicles from the EUG 40 presented a greater diameter than the non-cultured control group (Table 3). Regarding the oocyte diameter, regardless of the follicular category and the day of culture, the lowest diameters were obtained in groups grown in vitro in the absence or presence of antioxidants, except for the EUG 40 treatment. In this treatment, regardless of the follicular category, oocyte diameter was significantly higher than in the non-cultured control (Table 4). In general, in the tested treatments, a reduction in follicular and oocyte diameter was observed from day 1 to day 7 only in primordial and intermediate follicles. However, the EUG 40 treatment was the only treatment in which both follicular and oocyte diameters were increased significantly throughout the culture, regardless of the follicular category (Tables 3, 4). When compared to MEM, after 7 days of culture, except for the primordial follicle category in AA treatment, the addition of AA, ANE and EUG (regardless of the concentration) resulted in primordial, intermediate and primary follicles with significantly greater diameters. Concerning oocyte diameter, the addition of AA (only for intermediate follicle), ANE (primordial and intermediate follicles), EUG 10 and 20 (only for intermediate follicles) and EUG 40 (primordial, intermediate and primary follicles) significantly increased oocyte diameter after 7 days of culture. When the tested antioxidants were compared, except for the oocyte diameter of primary follicles, greater follicular and oocyte diameters were observed after 7 days of culture when EUG at 40 μM was added to the culture medium. Considering primary follicles, after 7 days of culture, a greater oocyte diameter was observed in the EUG 40 treatment when compared to the AA treatment (P < 0.05).

In the present study, the follicular and oocyte diameters were compared among the follicular categories, i.e., primordial, intermediate and primary follicles within the same treatment and day of culture (Table 3). Except for the MEM treatment, when similar (P < 0.05) follicular (day 7) and oocyte (days 1 and 7) diameters were observed between primordial and intermediate follicles, a significant and progressive increase in follicular and oocyte diameter was observed from the primordial to the primary follicle category both in the non-cultured control and in the cultured treated groups, regardless of the day of culture.

Follicles were classified as viable (Figure 1H) or non-viable (Figure 1I) if they were negatively or positively stained with trypan blue, respectively. After 7 days of culture, the percentages of viable follicles were similar among MEM (68.6%) ANE 300 (75.0%), AA (65.7%), and EUG 40 (83.9%) treatments. However, lower rates of viable follicles were observed in MEM and AA treatments than in the non-cultured control (88.2%) (Figure 3).

The quantification of H₂O₂ production and thiol groups was performed after 7 days of culture (Figure 4). In all treatments, H₂O₂ production was similar, except when EUG 40 was added to the medium, which resulted in a decrease in H₂O₂ production. No differences were observed when comparing thiol levels.

The neutralizing capacity against ABTS and DPPH on day 7 is shown in Figure 5. All tested antioxidants, except ANE 300, were able to significantly decrease the levels of ABTS and DPPH when compared to the control. The highest capacity to neutralize free radicals was observed with EUG 40, whereas AA was similar to EUG 20 and 40.

Representative staining images of H3K4me3 (Figures 6A–F) and calreticulin (Figures 6G–L) follicles after 7 days of culture are shown. The labeling intensity of H3K4me3 after 7 days of in-vitro culture is shown in Figure 6M. All cultured tissues presented a similar intensity of H3K4me3 labeling, regardless of the follicular class, except when the ovarian tissue was cultured in the presence of EUG 40, resulting in a significant decrease of H3K4me3 labeling. All ovarian tissues cultured in the presence of antioxidants, especially EUG 40, presented a significant decrease in H3K4me3 labeling.

Figure 6N shows the labeling intensity for calreticulin after 7 days of in-vitro culture. Only culturing of primordial and growing follicles in the presence of EUG 40 resulted in a labeling intensity similar to that observed in the non-cultured tissue. When compared to cultured control tissues, both EUG 40 and ANE 300 significantly increased the labeling intensity of calreticulin in growing follicles. Such effect was observed in all follicular categories when EUG 40 was added to the culture medium.

Figure 7 depicts the mRNA expression of the markers CAT (7A), ERP29 (7B), KDM1AX1 (7C) and KDM3A (7D). The mRNA expression of CAT was not affected by the treatments, whereas the mRNA relative expression of ERP29 and KDM3A was significantly increased when the culture medium was supplemented with EUG 40. The mRNA relative expression of KDM1AX1 was significantly increased when the culture medium was supplemented with EUG 40 or ANE 300.