Effect of docosahexaenoic acid on in vitro growth of bovine oocytes

Abstract Purpose The present study investigated the effects of docosahexaenoic acid (DHA) on the growth of bovine oocytes. Methods Oocytes and granulosa cell complexes (OGCs) were collected from early antral follicles (0.4‐0.7 mm) on the surface of ovaries harvested from a slaughterhouse. The OGCs were cultured with 0, 1, and 10 μmol/L docosahexanoic acid (DHA) for 16 days. Results Antrum formation of the OGCs and the number of granulosa cells (GCs) surrounding the oocytes were comparable among groups, whereas supplementation of 0.1 μmol/L of DHA significantly improved oocyte growth. Oocytes grown with DHA had a higher fertilization rate, acetylation levels of H4K12, and ATP contents, as well as a lower lipid content compared with those grown without DHA. In addition, GCs surrounding OGCs grown with DHA had low lipid content compared with vehicle counterparts. Furthermore, when GCs were cultured in vitro, DHA increased ATP production, mitochondrial membrane potential, and reduced lipid content and levels of reactive oxygen species. RNA‐seq of GCs revealed that DHA increased CPT1A expression levels and affect genes associated with focal adhesion, oxidative phosphorylation, and PI3K‐AKT etc Conclusion The results suggest that DHA supplementation affects granulosa cell characteristics and supports oocyte growth in vitro.

with the high clinical outcomes after assisted reproductive technology. 2,3 In this context, the beneficial effects of DHA on cow reproduction have been reported. For example, DHA concentration is higher for follicular fluid (FF) of dominant follicles compared with those of subordinate follicles in both cows and heifers. 4 In addition, supplementation of cattle with fish oil or algae containing DHA increased DHA concentration in plasma and improved the number and size of follicles as well as reproduction performance. [5][6][7] Although the causal relationship between good reproductive performance and DHA is unclear, some pioneering studies have reported that supplementation of the in vitro maturation medium of oocytes with DHA improves the developmental competence of oocytes, cleavage of embryos, and the quality of blastocysts in cows and pigs. 8,9 In addition, Elis et al 10 reported that supplementation of the maturation medium of oocytes with DHA decreased lipid content in both oocytes and cumulus cells, and supplementation with the fatty acid receptor FFAR4 agonist TUG-891 also improved oocyte developmental competence. However, in vitro maturation of bovine and porcine oocytes takes only one and two days, respectively. In large mammals such as pigs and cows, oocyte growth takes longer periods, but no study has addressed the effect of DHA on oocyte growth. In the present study, oocytes derived from early antral follicles were cultured for 16 days, and the effect of DHA on in vitro oocyte growth was examined.

| Ethical approval
Collection of ovaries from a slaughterhouse for experimental use was approved by the Committee for the Care and Welfare of Experimental Animals at Tokyo University of Agriculture.

| Medium and chemicals
All chemicals were purchased from Nacalai, unless otherwise stated.

| Measurement of diameter of oocytes grown in vitro
Oocytes were denuded from surrounding cells. The diameter of the ooplasm (horizontal and vertical diameters) was measured using a digital microscope (Keyence). The average of the two values was calculated and reported as the diameter.

| Measurement of the number of GCs consisting a OGCs grown in vitro
GCs were detached from oocytes and dispersed by vigorous pipetting in a cell-dispersion cocktail (Accumax; Innovative Cell Technologies, Inc). The total cell number was calculated using a hemocytometer to obtain the average GC number per OGC.

| Measurement of lipid content in oocytes and surrounding GCs
Granulosa cells and oocytes were separated from each OGC. Lipid content was determined by Nile red staining (Wako), as described previously. 14 Briefly, oocytes were incubated for 10 minutes in PBS containing 10 μg/mL Nile Red. Fluorescence images of the oocytes were captured using a fluorescence microscope (Keyence), and the fluorescence intensity of whole oocytes was measured using ImageJ software (National Institutes of Health, Bethesda, MD, USA). In addition, GCs were enzymatically dispersed as described above and stained with Nile red and Hoechst 33342. GCs were observed under a fluorescence microscope (Leica), and the images were captured to obtain the ratio of the fluorescence intensity of Nile red to that of Hoechst. In addition, the GCs were subjected to flow cytometry using a NovoCyte Flow Cytometer (2000R, ACEA Biosciences, Inc) in FL2 channel (excitation laser 488 nm, emission filter 572/28 nm), followed with data analysis with NovoExpress Software.

| Detection of acetylated H4K12 by fluorescence immunostaining
Oocytes were fixed in 4% paraformaldehyde for 1 day and subjected to immunostaining. Immunostaining was performed as previously reported. 15 The primary and secondary antibodies used for this proce-

| Measurement of ATP in oocytes
At the end of IVG, oocytes were denuded from GCs, and ATP content was determined by measuring the luminescence generated in an ATP-dependent luciferin-luciferase reaction (ATP assay kit; Toyo-Inc), as described previously. 16 Each sample was prepared by adding individual oocytes to 50 μL of distilled water.

| In vitro maturation and fertilization
All OGCs with an antrum cavity were selected, and oocytes with 2-3 GC layers were removed. The oocytes were cultured in IVM for 24 hours, and the frequency of oocytes at metaphase 2 stage, or fertilization rate, followed by IVF, was determined. For IVF, the oocytes were co-incubated with thawed semen from a Japanese black bull for 5 hours, and then the oocytes were subsequently cultured for 13 hours to examine the fertilization rate. The semen was washed with a 45%-60% Percoll solution (Amersham Biosciences) to create a discontinuous gradient for centrifugation (800 g for 10 minutes). The final sperm concentration in IVF medium was 1 × 10 6 cells/mL. To evaluate the fertilization rate, oocytes were denuded from the surrounding GCs, and the oocytes were transferred into aceto-alcohol (ethanol:acetic acid = 3:1) for 3 minutes, and the number of pronuclei was examined under a stereomicroscope (Olympus). Oocytes with two clear pronuclei were determined to be normally fertilized oocytes. Oocytes with over three pronuclei or one pronucleus were determined to be abnormal fertilized oocytes. Maturation and fertilization rates were examined four and five times, respectively.  To determine the ATP content, GCs in each well were frozen and thawed three times with 100 µL water, and water was collected from each well following vigorous pipetting. This water (50 μL) was used to determine the ATP content by measuring the luminescence generated during an ATP-dependent luciferin-luciferase reaction using an ATP assay kit (Toyo-Inc). In addition, the remaining half of the water was used for DNA extraction, and the copy number of nucleic DNA was determined by real-time PCR as described below. Thereafter, the ATP content of 10 000 GCs was calculated.

| Measurement of mitochondrial and nucleic DNA copy number in GCs
Granulosa cells cultured on 96-well plates were lysed in 50 μL of lysis buffer (20 mmol/L Tris, 0.4% proteinase K, 0.9% Nonidet-P40, and 0.9% Tween 20) followed by incubation at 55℃ for 30 minutes and then at 95℃ for 5 minutes. The nuclear DNA (nDNA) and mitochondrial (mtDNA) copy number in GCs were determined using real-time PCR targeting the bovine mitochondrial genome and a single-copy nuclear gene. PCR was performed using a CFX Connect™ real-time PCR detection system (Bio-Rad). The PCR primer set was designed using Primer3Plus (https://w w w.bioin forma tics.nl/cgi-bin/prime r3plu s/prime r3plus.cgi). Primers for nDNA were 5′-ttccactctgcacagtagcg-3′ and 5′-cccttactggttgtggcact-3′, targeting a one-copy sequence of 83 bp (NC_037334.1). The primers for Mt-DNA were 5′-acccttgtacctttgcat-3′ and 5′-tctggtttcgggctcgttag-3′ targeting a mitochondrial genome sequence of 81 bp (NC_006853.1). The PCR conditions were as follows: initial denaturation at 95℃ for 1 minute, followed by 40 cycles at 98℃ for 5 seconds and 60℃ for 10 seconds. A standard curve was generated for each run using 10-fold serial dilutions representing the copy number of the external standard. The external standard was the PCR product of the corresponding gene cloned into a vector using the Zero Blunt TOPO PCR cloning kit (Invitrogen), which was sequenced before use. DNA copy number in the standard was calculated using the concentration of DNA, molecular weight of the vector, and Avogadro's number. The amplification efficiency in all trials was >1.98. Thereafter, mtDNA per nDNA was calculated to obtain the mitochondrial copy number per granulosa cell.

| RNA-seq of GCs cultured with or without DHA
Granulosa cells were collected and cultured with or without DHA for 2 days, as described in Section 2.9 and GCs' RNA was extracted.
RNA extraction was conducted using the RNAqueous ® Kit (Life Technologies), and three batches of RNA were produced using differential ovary series. RNA quality was confirmed using an Agilent

| Statistical analysis
All measurement data are presented as the mean ± standard error of the mean (SEM), and all data were analyzed using the Kolmogorov-Smirnov test followed by Student's t test, and nonparametric data were analyzed using the Mann-Whitney U test. The data among the three groups (DHA, 0, 1, and 10 μmol/L) were analyzed using analysis of variance (ANOVA), followed by Tukey's post hoc test. Maturation and fertilization rate of oocytes were analyzed using Chi square test. Statistical significance was set at P < .05.
Statistical analysis and calculation of correlation coefficients were conducted using Bellcurve for Excel.

| RE SULTS
When OGCs were cultured for 16 days, they formed an antrum-like cavity surrounding the oocytes, and the rate of antrum formation did not differ among groups (vehicle, 1, and 10 μmol/L DHA) (Figure 1).
The number of GCs consisting of OGCs did not differ among the groups, whereas 1 μmol/L DHA significantly increased the diameter of oocytes compared with the vehicle control ( Table 1). Half of the oocytes grown with 1 μmol/L DHA reached metaphase 2 stage, but the value did not significantly differ between groups. Oocytes grown with 1 μmol/L DHA had higher fertilization ability with significantly higher total fertilization rate (73.6%) compared with those cultured with vehicle ( Table 2). Oocytes grown with 1 μmol/L DHA had higher acetylation levels of histone H4K12 and tended to have higher ATP content (P = .08) ( Table 3), whereas lipid content in the oocytes was significantly lower than that in oocytes grown with the vehicle. Supplementation of medium with 1 μmol/L DHA decreased the lipid content in the GCs surrounding oocytes (Figure 2A), and the reduction in lipid content was confirmed by FACS ( Figure 2B).
When GCs were cultured with or without DHA (1 μmol/L or vehicle) for 3 days, DHA increased the ATP content in GCs on both days 2 and 3 ( Figure 3A). Furthermore, DHA increased MMP on day 2 ( Figure 3B) and decreased the lipid content and ROS levels on day 3 ( Figure 3C,D).
To determine the mechanism underlying the effect of DHA on GCs, we cultured GCs with or without DHA for 2 days and conducted RNA-seq. A total of 17 457 genes were detected and 595 genes were found to be differentially expressed. The KEGG pathways enriched by the DEGs (P < .05) were oxidative phosphorylation, focal adhesion, and Ras signaling (Table 4). Regarding oxidative phosphorylation genes, CPT1A was significantly (P = .019) increased, whereas fatty acyl-CoA synthase, CPT2, and carnitine translocase did not differ between the groups. In addition, ten of the 13 mitochondrial genes, except for COX2, ND4L, and ND5, were significantly downregulated, and of the 1139 nuclear genes encoding mitochondrial proteins, 21 genes were significantly downregulated, except for CPT1A. Genes related to mitochondrial biogenesis (PPARGC1a, NRF1, NRF2, and TFAM) did not differ between the DHA and control groups. Using the DNA extracted from the same GCs batch, we examined the mitochondrial DNA copy number in GCs but did not found a difference between the two groups (DHA, 119.6 ± 1.9 vs Vehicle, 120.2 ± 1.4, N.12).

| D ISCUSS I ON
The present study demonstrated that DHA improved oocyte growth and increased their diameters and fertilization ability. Furthermore, DHA improved oocyte quality markers, including the acetylation levels of H4K12. In addition, DHA treatment reduced the lipid content in both oocytes and GCs. When GCs were cultured with DHA, lipid, and ROS levels decreased, concomitant with an increase in ATP content and MMP. RNA-seq showed that DHA affects mitochondrialrelated genes.
The oocyte diameter increases as the follicle develops. 17,18 In cows, oocytes with a small diameter (<120 μm) have low meiotic maturation competence, 18 whereas oocytes with a diameter greater than 120 μm had a high ability to complete nuclear maturation, fertilization, and development to term. 19,20 The quality of oocytes has been evaluated using various indexes. For example, high competent bovine oocytes have higher levels of ATP compared with those in poor quality oocytes, 21  underlying the effect of DHA using microarray could not define signaling pathways. 10 The present RNA-seq showed enrichment in focal adhesion, Ras, mTOR, and PI3K-Akt signaling pathways.
It has been reported that treatment of colon cells with DHA reduced Ras translocalization to the plasma membrane and inhibited GTP-bound Ras at the membrane. 35 In addition, DHA has been shown to protect cells from palmitic acid-induced lipotoxicity through PI3K/AKT and mTOR signaling. 41  induced apoptosis in cancer cells by stimulating AMPK, PI3K-Akt, and mTOR signaling. 42 These reports suggest that mitochondrial function, focal adhesion, Ras, PI3K/Akt, and mTOR are possible mechanisms underlying the effect of DHA on GCs. However, the evidence is not enough to determine possible mechanism of DHA on oocyte growth, and the signaling pathway needs to be elucidated in future studies.
In conclusion, consistent with previous reports on the beneficial effects of DHA on in vivo follicle development, our results indicate that DHA supports the in vitro development of oocytes derived from bovine EAFs.

ACK N OWLED G M ENTS
This study was supported by JSPS KAKENHI (grant number 16K07996).

CO N FLI C T S O F I NTE R E S T
The authors declare no conflicts of interest.

H U M A N/A N I M A L R I G HTS
This article does not contain any studies with human. Animal study: In this study, bovine ovaries were collected from a slaughterhouse.
This study was approved by the ethics committee for animal experiments of the Tokyo University of Agriculture.

CLI N I C A L TR I A L S R EG I S TR ATI O N
This study does not include clinical trials.