Melatonin prevents postovulatory oocyte aging and promotes subsequent embryonic development in the pig

Effects of different concentrations of melatonin on delaying oocyte aging

In our studies, oocytes with a female pronucleus or fragmented oocytes were defined as activated (Fig. 1A). The optimal condition for stimulation is very important to detect the degree of aging in oocytes. Few fresh oocytes and most aged oocytes should be activated under this condition. When we compared different stimulation times (10 μs VS 20 μs), we found that most fresh oocytes were activated under different voltages with 20 μs. However, fresh oocytes showed different degrees of activation under different voltages with 10 μs (Fig. 1B). To detect the optimal voltage for activation, we used 60 V, 70 V, 80 V and 90 V with 10μs to stimulate fresh oocytes and found that fresh oocytes had lower activation under 60 V or 70 V (Fig. 1C). So we used 70V and 10 μs to activate oocytes in the following experiments.

Figure 1. Effects of different stimulus conditions and concentrations of melatonin on oocyte activation. (A) Fresh oocytes were activated by weak stimulus (70V, 10μs, 2 pulses and stained with DAPI to show female pronucleus formation. (B) Oocytes were activated by different combinations of electric pulse times (10 μs and 20 μs) with voltage intensity (20 V, 50 V and 100 V). (C) Oocytes were activated by different voltage intensity treatment (60 V, 70 V, 80 V and 90 V; 10 μs). (D) Oocyte aged for 24 hr or 48 hr in vitro were treated with different concentrations of melatonin and activated by weak stimulus set 70 V, 10 μs, 2 pulses. All graphs show mean ± s.e.m. Abbreviations used in this and all subsequent figures: IVA, in vitro aging. Independent replicates were conducted with a minimum of 25 oocytes/replicate, at least 3 stable replicates were obtained. *P<0.05, **P <0.01. Bar = 100 μm.

It has been reported that 1 mM melatonin could significantly inhibit oocyte aging in the mouse [9]. Here we used 0.1 mM, 1 mM and 2 mM melatonin to treat porcine oocytes for 24 hr or 48 hr. The results showed that 0.1 mM and 1 mM melatonin had no effects on oocytes aged for 24 hr or 48 hr in vitro. However, 2 mM melatonin could significantly decrease the activation percentage of aged oocytes after treatment for either 24 hr or 48 hr (Fig. 1D).

Melatonin maintained normal oocyte morphology during postovulatory oocyte aging

Melatonin could maintain oocyte morphology during postovulatory aging and decrease their spontaneous parthenogenetic activation (Fig. 2A-E). In oocytes aged for 24 hr in vitro, most of the oocytes showed normal morphology with low fragmentation and melatonin had no effects on them. However, the cytoplasm was non-uniform and it was dispersed being dark-pigmented in oocytes aged for 48 hr and more than half of the aged oocytes became fragmented (Fig. 2D, F). If oocytes were treated with 2 mM melatonin for 48 hr, this abnormal morphology could be inhibited effectively and the fragmentation percentage was decreased significantly to a very low percentage (Fig. 2E, F). This phenomenon was rarely presented in IVA 24 hr oocytes, as most of them still exhibited normal morphology (Fig. 2B) with an extremely low fragmentation percentage and there was no difference when they were treated with 2mM melatonin (Fig. 2C).

Figure 2. Effects of oocyte aging and melatonin on oocyte morphology. (A) Fresh oocytes. (B) Oocytes aged for 24 hr. (C) IVA 24 hr oocytes treated with 2mM melatonin. (D) Oocytes aged for 48 hr. (E) IVA 48 hr oocytes treated with 2mM melatonin. (F) Fragmentation percentage in oocytes aged for 48 hr and treated with 2mM melatonin for 48 hr. Oocytes with abnormal morphology were indicated by arrowheads. All graphs show mean ± s.e.m. Abbreviations used in this and all subsequent figures: MT, melatonin. Independent replicates were conducted with a minimum of 30 oocytes/replicate, at least 3 stable replicates were obtained. ***P <0.001. Bar = 100 μm.

Melatonin promoted embryonic development of postovulatory aging oocytes after parthenogenetic activation

In order to determine whether melatonin could maintain the quality of aged oocytes, especially regarding the subsequent embryonic development, parthenogenetic activation was carried out. Approximately 50% fresh oocytes could develop to the blastocyst stage (Fig. 3A, D), while only 10% blastocysts obtained from oocytes aged for 24 hr and none of the oocytes aged for 48 hr could develop to blastocysts after activation (Fig. 3B, D). In the presence of 2 mM melatonin, there were nearly 20% oocytes aged for 24 hr that could develop to blastocysts after parthenogenetic activation (Fig. 3C, D). However, melatonin had no impact on embryonic development of IVA 48hr oocytes and none of the activated oocytes could develop to blastocysts (Fig. 3D). To detect the effects of melatonin on the development of activated oocytes aged for 24 hr, we selected several embryonic development-related genes (SLC2A1, DSC2, DNMT1, DNMT3A, AQP3 and CDH1) and measured their expression. We found that oocyte aging reduced the expression of SLC2A1 in blastocysts from IVA 24 hr oocytes and melatonin could increase its expression significantly (Fig. 3E).

Figure 3. The development of embryos from fresh, aged and melatonin treated oocytes after parthenogenetic activation. (A) Fresh oocytes. (B) Oocytes aged for 24 hr in vitro. (C) IVA 24 hr oocytes treated with 2mM melatonin. (D) Blastocyst formation from fresh oocytes, IVA 24 hr or 48 hr oocytes and oocytes treated with melatonin for 24 hr or 48 hr after strong stimulus (800V, 40μs, 2 pulses). (E) The expression of embryonic development-related genes (SLC2A1, DSC2, DNMT1, DNMT3A, AQP3 and CDH1) in parthenogenetic blastocysts from fresh and aged oocytes. All graphs show mean ± s.e.m. Independent replicates were conducted with a minimum of 25 embryos/replicate, at least 3 stable replicates were obtained. **P <0.01. Bar = 100 μm.

Melatonin alleviated oxidative stress in postovulatory aging oocytes

Oxidative stress is one of the major challenges threatening the developmental potential, thus we used both ROS Assay Kit and single living cell detection system to perform the ROS detection assay. As shown in Fig. 4A-G, the ROS levels of aged oocytes are significantly higher than those of melatonin-treated oocytes either aged for 24 hr or 48 hr, which demonstrated that melatonin reduced oxidative stress during oocyte aging.

Figure 4. Effects of melatonin on ROS levels and total glutathione in aged oocytes. (A) Fresh oocytes. (B) Oocytes aged for 24 hr. (C) IVA 24 hr oocytes treated with 2 mM melatonin. (D) Oocytes aged for 48 hr. (E) IVA 48 hr oocytes treated with 2 mM melatonin. (F) ROS fluorescence pixel intensity in IVA oocytes. (G) ROS level (counts of photons) detection of single cell by optical nanoprobes. (H) Total glutathione (T-GSH) content in fresh and IVA oocytes. (I) The expression of anti-oxidative stress genes (SOD1, GPx4 and CAT) in fresh and aged oocytes. All graphs show mean ± s.e.m. Abbreviations used in this and all subsequent figures: ROS, reactive oxygen species. Independent replicates were conducted with a minimum of 30 oocytes/replicate, at least 3 stable replicates were obtained. **P <0.01, ***P <0.001. Bar = 50 μm.

Because glutathione (GSH) has been shown to be present in various amounts in a diverse variety of cells and because it exerts powerful functions as an antioxidant in defending cells from the damage of oxidative stress, quantification of intracellular GSH was conducted. Surprisingly, the results showed that the level of intracellular GSH decreased significantly in aged oocytes and melatonin had no effects on increasing the level (Fig. 4H). We detected the expression levels of anti-oxidative stress genes (SOD1, GPx4 and CAT) in oocytes aged for 24 hr. Remarkably, oocyte aging decreased the expression of anti-oxidative stress genes; however, melatonin could increase the expression of GPx4 in IVA 24 hr oocytes (Fig. 4I).

Melatonin decreased early apoptosis levels in postovulatory aging oocytes

It had been shown that oocyte aging is accompanied by cellular apoptosis; thus, we next detected early apoptosis levels in aged oocytes by the Annexin V-FITC assay. With this detection, the observation of a green circle indicating the location on the external cellular membrane of the oocyte was defined as Annexin V positive (Fig. 5B), otherwise it would be defined as negative (Fig. 5A). The results showed that only 11% oocytes displayed early apoptosis in fresh oocytes. This percentage increased to 32% and 52% if oocytes were aged for 24 hr or 48 hr in vitro. However, melatonin could inhibit early apoptosis during oocyte aging. The early apoptosis percentage in aged oocytes decreased to 18% and 30% significantly after treatment with melatonin for 24 hr or 48 hr (Fig. 5C).

Figure 5. Effects of melatonin on early apoptosis in aged and melatonin treated oocytes by Annexin-V FITC detection assay. (A) Negative control. (B) Apoptosis positive. (C) Apoptosis positive percentage in fresh, aged and melatonin treated oocytes. (D) The expression of anti-apoptosis genes (BCL2, Sphk1 and BCL-xl) in fresh and aged oocytes. (E) The expression of pro-apoptosis genes (P53, Caspase-3, Bax and BAD) in fresh and aged oocytes. All graphs show mean ± s.e.m. Independent replicates were conducted with a minimum of 25 oocytes/replicate, at least 3 stable replicates were obtained. **P <0.01. Bar = 25μm.

Furthermore, we detected the expression of anti-apoptosis genes (BCL2, Sphk1 and BCL-xl) and pro-apoptosis genes (P53, Caspase-3, Bax and BAD) in oocytes aged or treated with melatonin for 24 hr in vitro. For the anti-apoptosis genes, it was shown that their expressions were decreased in oocytes aged for 24 hr and melatonin could inhibit the decrease of the expression of BCL-xl (Fig. 5D). For the pro-apoptosis genes, oocyte aging induced the increase of Bax and BAD's expression and melatonin could decrease their expression, especially BAD's expression (Fig. 5E). This result suggested that melatonin blocked early apoptosis effectively during oocyte aging by regulating the expression of anti-apoptosis genes or pro-apoptosis genes.

Melatonin retarded the decline of mitochondrial membrane potential ΔΨm in postovulatory aging oocytes

It is well known that mitochondria play a crucial role in maintaining critical metabolic cell functions. So we carried out the mitochondrial membrane potential assay to investigate the mitochondrial membrane potential state during oocyte aging. To detect the mitochondrial membrane potential, we analyzed the ratio of green/red fluorescence. Clearly, fresh oocytes showed the lowest value (Fig. 6A, F, K); the aged oocytes displayed much higher ratios compared to those treated with melatonin for 24hr or 48hr, respectively (Fig. 6B-E, G-J, K), suggesting a significant efficacy of melatonin on keeping mitochondrial membrane potential in postovulatory aging oocytes. In addition, RT-qPCR was performed to quantify the expression levels of mitochondria-related genes (Cytochrom C, SIRT1, Akt2 and Polg2) in oocytes aged for 24 hr. The results showed that oocyte aging decreased the expression of mitochondria-related genes; however, melatonin could increase some genes' expression, like Polg2 (Fig. 6L).

Figure 6. Effects of melatonin on mitochondrial membrane potential in aged and melatonin treated oocytes by JC-1 detection assay. (A) Green-Fresh oocytes. (B) Green-Oocytes aged for 24 hr. (C) Green-IVA 24 hr oocytes treated with 2mM melatonin. (D) Green-Oocytes aged for 48 hr. (E) Green-IVA 48 hr oocytes treated with 2mM melatonin. (F) Red-Fresh oocytes. (G). Red-Oocytes aged for 24 hr. (H) Red-IVA 24 hr oocytes treated with 2 mM melatonin. (I) Red-Oocytes aged for 48 hr. (J) Red-IVA 48 hr oocytes treated with 2 mM melatonin. (K) Fluorescence pixel ratio (green/red) in fresh and aged oocytes. (L) The expression of mitochondria-related genes (Cytochrome C, SIRT1, Akt2 and Polg2) in fresh and aged oocytes. All graphs show mean ± s.e.m. Independent replicates were conducted with a minimum of 25 oocytes/replicate, at least 3 stable replicates were obtained. **P <0.01. Bar = 50 μm.

Preparation of porcine oocytes

Porcine ovaries were obtained from a slaughterhouse and transported to the laboratory while maintained at <34°C. Follicular fluid from 3–6 mm antral follicles was aspirated with an 18-gauge syringe. Cumulus oocyte complexes (COCs) with uniform cytoplasm and several layers of cumulus cells were selected and rinsed three times in washing medium (TCM-199 medium supplemented with 10% porcine follicular fluid (pFF), 5 μg/mL insulin, 10 ng/mL EGF, 0.6 mM cysteine, 0.2 mM pyruvate, 25 μg/mL kanamycin). Approximately 30 COCs per well were cultured in 96 well plates containing TCM-199 medium supplemented with 10% porcine follicular fluid (pFF), 5 μg/mL insulin, 10 ng/mL EGF, 0.6 mM cysteine, 0.2 mM pyruvate, 25 μg/mL kanamycin and 5 IU/mL of each eCG and hCG, covered with mineral oil. The oocytes were matured for 44 hr at 38.5°C, 5% CO₂ in humidified air.

In vitro aging of porcine oocytes

For oocyte aging in vitro, cumulus cells were removed by vortexing for 4 min in 0.1% hyaluronidase (in TLH-PVA [25], TL-Hepes medium supplemented with 0.1% PVA) after porcine COCs maturation for 44 hr. Only oocytes with first polar bodies were used for the experiments. The treated oocytes were then cultured in wells of a 96-well culture plate containing 150 µl of NCSU23 medium (108.7 mM NaCl, 4.8 mM KCl, 1.7 mM CaCl₂.2H₂O, 1.2 mM KH₂PO₄, 1.2 mM MgSO₄.7H₂O, 25.1 mM NaHCO₃, 5.5 mM Glucose, 1.0 mM L-Glutamine, 7.0 mM Taurine, 5.0 mM Hypotaurine, 0.05 mg/mL Gentamicin, 4.0 mg/mL Fatty acid-free BSA) [26] and covered with mineral oil at 38.5°C under 5% CO₂ in humidified air for 24 or 48 hr. For melatonin treatment, 100 mM melatonin stock was diluted to final concentration 2mM. The treated oocytes were cultured in wells containing 150 µl of freshly prepared culture medium described above and covered with mineral oil at 38.5°C under 5% CO2 in humidified air for 24 or 48 hr.

Parthenogenetic activation and in vitro culture of embryos

Fresh or aged oocytes were placed between 0.2-mm-diameter platinum electrodes 1 mm apart in activation medium. Activation was induced with two direct-current (DC) pulses of 1.2 kV/cm for 40 μs on a BTX Elector-Cell Manipulator 200 (BTX, San Diego, CA) according to the experimental design. The medium used for activation was 0.3M mannitol, supplemented with 1.0 mM CaCl₂, 0.1 mM MgCl₂, and 0.5 mM Hepes. The orientation of oocytes and polar bodies was not vertical to platinum wire electrodes during electrical activation. After activation treatment, embryos were washed and transferred into NCSU medium with 5 μg/ml cytochalasin B for 4 hr to inhibit second polar body extrusion, then cultured in 150 µl NCSU medium covered with mineral oil in a 96-well culture plate. The culture environment was 5% CO₂ in air at 38.5°C. Parthenogenetically activated oocytes were evaluated for the percentage of blastocysts on Day 6.

RNA isolation and real-time RT-PCR

Total RNA was isolated from 50 porcine oocytes or blastocysts using RNA prep Pure Micro Kit (TIANGEN, Beijing, China). Enhanced GFP (eGFP) cRNA was transcribed in vitro from pIVT-eGFP [27] and 1 ng was added to each sample prior to RNA isolation as an internal control. Real-time RT-PCR was performed as previously described [28], using cDNA from two oocytes or embryos per reaction. Relative gene expression was calculated using the ΔCt method [29] with eGFP expression for normalization. Primers are listed in Table S1.

Analysis of ROS levels

To identify oxidative stress/intracellular reactive oxygen species (ROS) levels in aged oocytes, a Reactive Oxygen Species Assay Kit was utilized to detect ROS as green fluorescent signals of DCFH diacetate (DCHF-DA; Beyotime Institute of Biotechnology, China). In brief, 20-30 oocytes from each treatment group were incubated at 38.5 °C for 30 min in D-PBS containing 10 μM DCHF-DA. Then, the oocytes were washed 3 times in D-PBS containing 0.1% BSA before mounting on glass slides using D-PBS for microscopy. The fluorescence intensity in each oocyte was measured under a fluorescence microscope (Olympus, Tokyo, Japan) with the same scan settings for each sample. The fluorescent images were saved as graphic files in TIFF format. The normal fluorescence pixel intensities of each oocyte were analyzed using ImageJ software (version 1.50; National Institutes of Health, Bethesda, MD, USA).

To confirm the above results, we also detected ROS levels in single fresh and aged oocytes by employing optical fiber probes with ~10 μm tips (Rayme, Wuxi, China). The distal end of the tip was designed free for transmitting light by careful adjustment of the fiber placement angle in the sputter chamber. The silver coating of the nanoprobes is stable for subsequent fabrication procedures. The counts of photons were constantly detected and calculated during a 150-s period following the tips inserted into an immobilized oocyte in a PBS droplet [30].

Detection of intracellular GSH

The concentrations of total glutathione (T-GSH) were examined with a GSH/GSSG assay kit (Beyotime Institute of Biotechnology, China) based on an enzymatic method according to the manufacturer’s instructions. A total of 30-40 oocytes from each group were mixed with 30 µl of protein scavenger M solution supplied by the kit and vortexed thoroughly for 5 min, then the mixture was frozen at -80°C for 2 min and thawed at 38.5°C repeatedly for three times. The mixture was centrifuged at 10,000 rpm for 10 min at 4°C and put on ice for 5 min. The absorbance was monitored continuously at 412 nm with a microplate spectrophotometer (PerkinElmer, USA) for 25 min, with a reading recorded every 5 min and regarded the readings at the time point 25 min as the final absorbance values.

Annexin V-FITC assay

According to manufacturer’s instructions, Annexin V–fluorescein isothiocyanate (FITC) staining was performed using an Annexin V-FITC Apoptosis Detection kit (Vazyme, Nanjing, China) to detect the externalization of phosphatidylserine (PS) in early apoptotic MII oocytes. Briefly, 20-30 MII oocytes were washed twice in PBS containing 0.01% PVA (w/v), then incubated in 100µl binding buffer containing 5µl of Annexin V-FITC for 10 min at room temperature in the dark, transferred to 100µl 4% paraformaldehyde (PFA) for fixation for 1hr after being washed twice in PBS containing 0.01% PVA (w/v); after fixation and washes twice in PBS containing 0.01% PVA (w/v) again, oocytes were mounted on glass slides using DAPI for DNA fluorescence microscopy. Annexin-V positive oocytes were identified under a confocal system (Zeiss LSM 510 META, Germany) with 450–490 nm (excitation) and 520 nm (emission) filters.

Determination of mitochondrial membrane potential

The mitochondrial membrane potential of the oocytes was evaluated using a mitochondrial membrane potential assay kit with JC-1 (Beyotime Institute of Biotechnology, China). Oocytes were exposed to 10μM JC-1 in 100µl working solution at 38.5°C in 5% CO₂ for 20 min, after which they were washed with washing buffer to remove surface fluorescence, and then mounted on glass slides using D-PBS for microscopy. Laser excitation was set at 488 nm for green and 525 nm for red fluorescence, respectively. The fluorescence intensity in each oocyte was measured under a fluorescence microscope (Olympus, Tokyo, Japan) with the same scan settings for each sample. The fluorescent images were saved as graphic files in TIFF format. The normal fluorescence pixel intensities of each oocyte were analyzed using ImageJ software (version 1.50; National Institutes of Health, Bethesda, MD, USA). The ratio of green to red fluorescence pixels was used to analyze mitochondrial membrane potential.

Data analysis

For each treatment, three replicates were performed. Statistical analyses were carried out by analysis of variance. Differences between control, IVA groups and melatonin treated groups were evaluated by independent-sample t-tests using SPSS 22.0 statistical software. Data are expressed shown as mean ± SEM and P< 0.05 is considered as significant.