Anti-Müllerian hormone treatment enhances oocyte quality, embryonic development and live birth rate

Abstract

The anti-Müllerian hormone (AMH) produced by the granulosa cells of growing follicles is critical for folliculogenesis and is clinically used as a diagnostic and prognostic marker of female fertility. Previous studies report that AMH-pretreatment in mice creates a pool of quiescent follicles that are released following superovulation, resulting in an increased number of ovulated oocytes. However, the quality and developmental competency of oocytes derived from AMH-induced accumulated follicles as well as the effect of AMH treatment on live birth are not known. This study reports that AMH priming positively affects oocyte maturation and early embryonic development culminating in higher number of live births. Our results show that AMH treatment results in good-quality oocytes with greater developmental competence that enhances embryonic development resulting in blastocysts with higher gene expression. The transcriptomic analysis of oocytes from AMH-primed mice compared with those of control mice reveal that AMH upregulates a large number of genes and pathways associated with oocyte quality and embryonic development. Mitochondrial function is the most affected pathway by AMH priming, which is supported by more abundant active mitochondria, mitochondrial DNA content and adenosine triphosphate levels in oocytes and embryos isolated from AMH-primed animals compared with control animals. These studies for the first time provide an insight into the overall impact of AMH on female fertility and highlight the critical knowledge necessary to develop AMH as a therapeutic option to improve female fertility.

Anti-Müllerian hormone treatment increases number of ovulated oocytes and, positively affects oocyte quality and early embryonic development resulting in higher number of live births.

Introduction

In mammals, proper follicular development is critical for female fertility and is regulated by gonadotropic hormones and local paracrine/autocrine factors in a stage-dependent manner. The anti-Müllerian hormone (AMH) is a particular intraovarian peptide that is dynamically produced by developing ovarian follicles and plays a central role in the regulation of folliculogenesis in adult female mammals [1]. In recent years, AMH has received considerable attention, as it is extensively used as a diagnostic and prognostic marker for fertility as well as for pathophysiological conditions in women [2, 3]. For example, in women with polycystic ovary syndrome (PCOS), AMH levels are significantly higher and studies have shown that high AMH levels negatively affect neuroendocrine function [4]. Moreover, prenatal AMH treatment in mice can cause PCOS-like phenotype in the offspring [5]. Despite widespread use of AMH as a clinical biomarker, the mechanistic understanding of AMH actions and its influence in regulating female fertility is limited.

The AMH or Müllerian inhibiting substance is a member of the transforming growth factor-β super family with well-established roles in reproductive organ differentiation and ovarian follicular development [6]. In females, AMH is produced by the ovarian granulosa cells (GCs) of small growing follicles and AMH levels are tightly regulated throughout folliculogenesis [7, 8]. Studies in an AMH knockout mouse model [1, 9–11] have established that AMH controls the activation of primordial follicles and prevents primordial follicles from entering the pool of growing follicles thereby maintaining the ovarian reserve (follicle pool) throughout reproductive life [1, 10, 12, 13]. Interestingly, several in vitro studies have shown that AMH also inhibits follicle stimulating hormone (FSH)-stimulated follicle growth by decreasing the sensitivity of ovarian follicles to FSH [8, 10]. AMH inhibits FSH-induced processes like proliferation, aromatase activity, and luteinizing hormone receptor expression across species [14, 15]. These in vivo and in vitro studies have established that, in addition to inhibiting primordial follicle activation, AMH also inhibits FSH-stimulated follicle growth.

Previously [16], we reported that AMH administration in mice not only inhibits primordial follicle recruitment, but it also leads to more preantral but fewer antral follicles. However, the effect of AMH on the quality and developmental competency of oocytes as well as the in vivo effect of long-term AMH treatment on live birth are not known, a knowledge that is essential for understanding the overall impact of AMH on female fertility. In this study, we determine for the first time, the effects of AMH priming on oocyte quality and competence, early embryonic development, and fertility. Our results show that in vivo AMH treatment not only increases the number of ovulated oocytes but also has a positive effect on oocyte maturation and early embryonic development that result in more live births. Our data also offer new insights into the mechanisms of AMH action, which provide an overall picture of the influence of AMH on female fertility.

Materials and methods

Animal studies

Mouse studies were performed in accordance with the guidelines for the care and use of laboratory animals and were approved by the Institutional Animal Care and Use Committee at Michigan State University under the approval number PROTO202000156. For studies involving AMH treatment, 21-day-old prepubertal C57BL/6 J mice (Jackson Laboratory) were subjected to daily intraperitoneal (IP) injection of AMH (300 ng; recombinant mouse Mullerian-inhibiting factor (Amh) expressed in Escherichia coli, MBS968365, MyBiosource; n = 40 mice) or vehicle (n = 44 mice) for 21 days [16]. The concentration of AMH used was based on our previously published studies [16]. Following the treatment, animals were divided into five groups: Group 1 for oocyte collection (n = 10 mice/treatment); Group 2 for embryo/cleavage study (n = 7 mice/treatment); Group 3 for blastocyst staining (n = 14 mice/control and n = 13 mice/AMH treatment), Group 4 for mitochondrial studies (n = 5 mice/treatment), and Group 5 for fertility test (n = 5 mice/treatment). The experimental details are shown in Figure 1A.

Figure 1.

AMH priming prior to superovulation enhances the number of ovulated MII oocytes and their development to two-celled embryo and blastocyst stages. Prepubertal mice were treated with AMH (300 ng; n = 40 animals) or vehicle/control (n = 44 animals) for 21 days followed by 7–12 days of no treatment. Thereafter these animals were divided into groups for various experiments. All the groups were subjected to a superovulation regime (10 U pregnant mare serum gonadotropin (PMSG); 10 U human chorionic gonadotropin (hCG)). (A) The number of superovulated MII oocytes (n = 10 mice/treatment, P = 0.0321 using the t-test) isolated from AMH-primed and vehicle-treated (control) mice. (B) Percentage of the of superovulated MII oocytes isolated from AMH-primed and vehicle-treated (control) mice that were fertilized following in vitro fertilization (n = 80 oocytes isolated from 3 mice/treatment). (C) Percentage of fertilized oocytes that developed to the two-celled embryo stage after 24 h. (D) One-celled fertilized embryos collected from AMH-primed and vehicle-treated mice following mating with male mice (in vivo fertilization) (n = 7 animals/treatment). (E, F) Percentage of one-celled embryos reaching the two-celled stage (E) and then to the blastocyst stage (F). Data are represented as mean ± SEM (^*P < 0.05).

Superovulation, oocyte metaphase II (MII)/one-celled embryo collection, and in vitro fertilization

For AMH priming and superovulation, mice were treated with daily IP injection of AMH (300 ng) or vehicle for 21 days followed by 7–12 days of no treatment. Thereafter, these animals were subjected to a superovulation regime involving a single IP injection of 10 U of pregnant mare serum gonadotropin (Sigma, St. Louis, MO) followed 48 h later by 10 U of human chorionic gonadotropin (Sigma), as described previously [16–18]. Groups 2, 3, and 4 were mated with fertile males and 16–18 h from the appearance of vaginal plugs, cumulus–oocyte complexes (COCs) and one-celled embryo were surgically isolated from the ampulla of oviducts. Mature oocytes (MII stage) and one-celled embryo were denuded by digestion in M2 media containing 0.3% hyaluronidase (Sigma-Aldrich). The number of oocytes (MII) was determined by the presence of polar bodies and counted and either snap-frozen for gene expression (RNA-seq and real-time polymerase chain reaction (RT-PCR)) studies or subjected to in vitro fertilization (IVF).

For IVF, 10 μl of capacitated (incubated at 37°C for 1 h) sperm suspension (10 × 10⁶) isolated from the epididymis of C57BL/6J mice was introduced into a 500 μl drop containing three COCs. Sperm and oocytes were incubated together for 6 h, and then the oocytes were washed and cultured in a 20 μl drop of KSOM+AA medium (MR-121-D, Millipore, USA) under mineral oil at 37°C in a humidified chamber (5% CO₂) [19] and the two-celled stage was determined after 24 h.

RNA isolation and RNA-seq

Total RNA isolation, library construction, and RNA sequencing services were provided by Novogene. Details of RNA isolation, quality, and bioinformatic analysis of the RNA-seq data are provided in the Supplemental Material S3. For RNA-seq studies, 30–40 oocytes collected from AMH-primed and vehicle-treated (control) mice (n = 3/treatment) were used. Briefly, raw reads of the fastq format were processed through in-house Perl scripts and clean reads were mapped to the reference Mus musculus GRCm38 genome. Unique gene hit counts were calculated by the feature Counts from the Subread package v1.5.0-p3. Using the DESeq2 R package (1.20.0), the differentially expressed genes (DEGs) were identified. The Wald test was used to generate P-values and the Benjamini–Hochberg test for adjusted P-value. Genes with adjusted P-values < 0.05 and absolute log2 fold changes >1 were called as DEGs for each comparison. A heat map was constructed using rlog-transformed values obtained from RNA-seq data followed by z-normalization. A list of all the DEGs is in the Supplemental Material S1 and S2. Gene Ontology (GO) enrichment analysis of DEGs was done by the clusterProfiler R package, in which gene length bias was corrected. The enrichment of GO terms was tested using Fisher’s exact test to identify general biological processes, cellular compartments, and molecular functions affected by the significant DEGs. Pathways with adjusted P-values <0.05 and Q-values <0.05 were considered significant. The RNA-seq data are available in the Gene Expression Omnibus GSE195720 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE195720).

RNA extraction and quantitative real time PCR

Total RNA was isolated by the E.Z.N.A Total RNA micro kit (Omega) according to the manufacturer’s instructions. 1 μg of RNA was used for all the quantitative RT-PCR (qRT-PCR) reactions. Levels of Uqcrq, Cox7b, Tfam, Rps13, Sox18, Bmp2, Hif3a, and Il17rb were analyzed by the ∆∆Ct method using Taqman gene expression assay primers. Each target gene was normalized to Gapdh. The list of all the Taqman gene expression assay primers are provided in the Supplemental Information S3.

Immunocytochemistry and confocal microscopy

One-celled embryos were cultured for 4.5 days in KSOM+AA medium (MR-121-D, Millipore, USA) to the blastocyst stage, fixed in 3.7% paraformaldehyde, permeabilized with 0.1% (v/v) Triton X100 [20] and incubated with sex-determining region Y-box 2 (SOX2) or NANOG and caudal-type homeodomain (CDX2) primary antibodies overnight at 4°C and then labeled with the secondary antibodies, Alexa Fluor 594 anti-goat immunoglobulin (Ig) G (for Sox2), Alexa Fluor 594 anti-rabbit IgG (H + L) (for Nanog) and Alexa Fluor 488 anti-mouse IgG (for CDX2) for 1 h at room temperature. Thereafter, blastocysts were stained with 4′, 6-diamidino-2-phenylindole (DAPI) (1:10) and viewed using an Olympus FV1000 confocal laser scanning microscope (CARV; Atto Bioscience). Data were analyzed by the FluoView Viewer 3.0 software (Molecular Devices). Alexa 488 was excited using a 488 nm laser line, and fluorescence was measured using a 500–525 nm bandpass filter. Alexa 594 was excited using a 559 nm diode laser, and fluorescence was measured using a 570–650 nm bandpass filter. Details of all the antibodies and the concentrations used are provided in the Supplemental Information S3. For SOX2 staining, n = 14 (control) and 13 (AMH) mice and for NANOG staining n = 9 (control) and 10 (AMH) mice (1 blastocyst/animal) were used. The number (and percentage) of SOX2-, NANOG-, and CDX2-positive cells were determined by counting the z-stacks for blastocysts in the control and AMH-treated groups. The images presented are single optical sections (representing the middle region of the embryos) from the confocal z-series [20].

Assessment of mitochondrial distribution and mitochondrial membrane potential

Mitochondrial distribution was evaluated using MitoTracker Red (200 nM; M7512, ThermoFisher), which stains mitochondria of live cells. Mitochondrial membrane potential was specifically determined in single live oocytes and embryos using tetramethylrhodamine (TMRM) (75 nM), a low toxicity fluorescent dye tetramethyl rhodamine methyl ester perchlorate (T668, ThermoFisher) [21, 22], that accumulates only in active mitochondria with intact membrane potential. MitoTacker Red and TMRM were excited using a 559 nm diode laser, and fluorescence was measured using a 570–650 nm bandpass filter in an Olympus FV1000 confocal laser scanning microscope (CARV; Atto Bioscience). Data were analyzed by the FluoView Viewer 3.0 software (Molecular Devices). To allow comparison of staining intensity following confocal microscopy, the auto gain functions were switched off and the same gain and photomultiplier settings were used for all oocytes and embryos. The acquired images were processed and quantified by the ImageJ software. For each image, mean fluorescence intensity per oocyte and embryo were calculated and expressed as mean ± SEM of relative intensity (au) for control and treatment. For oocytes, n = 10 oocytes from 4 animals/treatment and for embryos n = 4 embryos from 4 mice/treatment were used.

Adenosine triphosphate quantification

Denuded oocytes (10 oocytes pooled/mice, n = 5 mice) isolated from gonadotropin-stimulated mice were collected in 50 μl of filtered ultrapure water and stored at −80°C until use. Adenosine triphosphate (ATP) levels were determined by the ATPlite™ luminescence ATP detection assay system (Perkin Elmer Life and Analytical Sciences B.V., Groningen, Netherlands, cat #6016943) according to the manufacturer’s instructions and previously published study [22] using a luminometer (Spectra Max M5, Molecular devices). Oocyte ATP values were then converted to weight (pg) using the molarity calculator by GraphPad as described previously [23] and normalized to the total number of oocytes.

Quantification of mitochondrial DNA copy number

Mitochondrial DNA (mtDNA) content was calculated using qRT-PCR as described previously [22]. Oocytes (five oocytes pooled/animal, n = 5 mice) were lysed using mammalian cell lysis buffer (Abcam) (with 200 μg/ml proteinase K and 0.1 mM ethylenediaminetetraacetic acid (EDTA), pH 8.0) and samples were used directly for PCR analysis. The mtDNA content was calculated using qRT-PCR by measuring the threshold cycle ratio (ΔCt) of a mitochondrial encoded gene B6 (B6F- 5′AACCTGGCACTGAGTCACCA-3′ and B6R 5′-GGGTCTGAGTGTATATATCATGAAGAGAAT-3′) versus a nuclear gene NDUFV1 (NDUFV1F 5′- CTTCCCCACTGGC CTCAAG-3′ and NDUFV1R 5′-CCAAAACCCAGTGATCCA GC-3′). Data were expressed as mtDNA/nuclear DNA (nDNA) as well as mtDNA copy number [24]. All measurements were performed in triplicate.

Fertility test

Mice pretreated with AMH or vehicle (n = 5 mice) for 21 days were mated with 6–10-week-old fertile male mice and the number of pups was recorded.

Statistical analysis

Statistical analysis was performed by GraphPad Prism version 9 (GraphPad Software Inc, California, USA). Statistical comparisons were done by two-tailed paired or unpaired t-test with Welch’s correction (for comparing two groups). Data are presented as mean ± SEM and a two-way analysis of variance (ANOVA) followed by Tukey’s post hoc analysis (for comparing multiple groups and variables) was carried out. Results with P ≤ 0.05 were considered significant.

Results

AMH priming enhances the number of ovulated MII oocytes and their development to the two-celled and blastocyst stage

In accordance with our previous studies [16], results (Figure 1B) show that AMH priming followed by superovulation increases the number of ovulated MII oocytes compared to vehicle-treated control animals (n = 10 mice/treatment, P = 0.0324). The percentage of superovulated oocytes that did not reach the MII stage was 19.34% for the control and 24.48% for AMH treatment. This difference is primarily due to more total retrieved oocytes obtained from the AMH-treated animals. To determine the developmental competence of these oocytes, ovulated MII oocytes were subjected to IVF and cultured to the two-celled stage. Results (Figure 1C) show that out of a total of 80 oocytes isolated from three AMH-primed animals, 48 oocytes (60%) were fertilized, (determined by the presence of pronuclei) compared with 38 out of 73 oocytes (52.1%) collected from three vehicle-treated animals. Furthermore, the percentage of fertilized oocytes that developed to the two-celled embryo after 24 h of in vitro insemination (Figure 1D) was significantly higher in the AMH-primed cohort (35 out of 48; 72.9 ± 1.28%) compared with the control (20 out of 38; 52.6 ± 5.30%). The percentage of oocytes that reached the two-celled stage was 27.39% for the control and 43.73% for AMH.

To negate any confounding effect of IVF, AMH- and vehicle-treated animals (n = 7 mice) following superovulation were mated with males and one-celled stage embryos were surgically isolated from the ampulla of oviducts. The number of one-celled embryos (19.4 ± 2.3 versus 13.42 ± 1.04, P = 0.0366) isolated from AMH-primed mice subjected to in vivo fertilization was significantly higher compared with that from vehicle-treated animals (Figure 1E). Moreover, when these isolated one-celled embryos were cultured, the percentage of one-celled embryos reaching the two-celled stage (86.1 versus 62.74%) and blastocyst stage (76.9 versus 69.47%) was also significantly higher in the AMH-treated group compared with the control animals (Figure 1F and G). The number of one-celled embryos collected after in vivo fertilization that reached blastocyst stage were 43.63% for control and 68.98% for AMH. This shows that in vivo AMH priming positively affects oocyte competence and early embryonic development.

AMH pretreatment increases the number of pluripotent cells in blastocysts

To gain further insights into the effects of AMH priming on embryonic development, we determined the pluripotency of the inner cell mass of blastocysts as a marker of embryo quality. SOX2 is a well-established transcription factor and a pluripotent marker associated with the pluripotent lineage of the early mouse embryo [25]. Similarly, NANOG is another transcription factor also expressed in the inner cell mass and participates in maintaining pluripotency [26]. Here we used SOX2- and NANOG-positive cells as a marker of embryonic quality. Results show that in general, blastocysts developed from fertilized one-celled embryos isolated from AMH-primed mice have significantly numerous cells along with SOX2-positive (Figure 2A, P = 0.0064) and NANOG-positive (Figure 2B, P = 0.0331)nuclei as well as CDX2-positive cells (marker of trophectoderm, P = 0.0001 for both SOX2 and NANOG). This shows that embryos developed from AMH-primed mice are of better quality compared with the vehicle-treated control mice.

Figure 2.

AMH pretreatment increases the number of pluripotent cells in blastocysts. (A, B) Representative immunofluorescence images from confocal microscopy depicting SOX2-positive (A), NANOG-positive (B) nuclei and CDX2-positive cells in blastocysts developed from fertilized one-celled embryos (from in vivo fertilization) isolated from AMH-primed and vehicle-treated (control) animals (Scale bars: 20 μm). The number (and percentage) of SOX2-positive (P = 0.0064 using two-way ANOVA), NANOG-positive (P = 0.0331 using two-way ANOVA), and CDX2-positive (P = 0.0001 for both SOX2 and NANOG using two-way ANOVA) cells were determined by counting the z-stacks for blastocysts in the control and AMH-treated groups. The images presented are single optical sections (representing the middle region of the embryos) from the confocal z-series. For SOX2 staining n = 14 and 13 mice and for NANOG staining n = 9 and 10 mice (1 blastocyst/animal) for control and AMH treatment, respectively were used.

To determine whether the observed positive effects of AMH treatment on the number of ovulated oocytes and their development to the blastocyst stage manifest in increased live births, we performed the fertility test of AMH- and vehicle-treated animals. Our results show that AMH-primed mice had significantly more pups compared with the vehicle-treated control animals (Table 1).

Table 1.

Fertility test

Treatment	n	Number of pups
Control	5	5.6 ± 0.75
AMH	5	7.8 ± 0.2*

Effect of AMH on MII oocyte transcriptome

To seek an understanding of the observed effects of AMH at the molecular level, we hypothesized that AMH priming positively affects oocyte quality that forms the basis of the enhanced oocyte competence and improved embryonic development leading to increased fertility. To confirm oocyte quality, we performed RNA-seq analysis in MII oocytes isolated from AMH-primed and vehicle-treated animals. DEseq2 analysis identified a total of 2760 annotated differentially expressed ENSEMBL genes, out of which, 2576 were upregulated and 184 were downregulated (adjusted P-value <0.05 by the Benjamini–Hochberg method and log2 fold changes >1). Hierarchical clustering of top 100 significant DEGs in oocytes from the control versus AMH-treated mice are shown in Figure 3A whereas the global transcriptional change across the two groups compared (control versus AMH) is represented by a volcano plot in Figure 3B. The complete list of significant DEGs is presented in the Supplemental Dataset. Based on established functions in the oocytes, eight DEGs (six upregulated and two downregulated)—Uqcrq, Cox7b, and Tfam (involved in electron transport chain [27, 28]); Rps13 (involved in cytoplasmic protein translation [29]); Sox18 (a pluripotent gene [30]), Bmp2 (critical for embryo development [31]), Hif3a (hypoxia-inducible factor, causes oocyte dormancy [32]), and Il17rb (inflammation marker, reduces oocyte quality [33]) were selected to verify the gene expression dataset. Figure 3C shows AMH priming significantly increases mRNA abundance of Uqcrq, Cox7b, Tfam, Rps13, Sox18, and Bmp2 whereas it decreases Hif3a and Il17rb mRNA levels in oocytes, which is in accordance with the RNA-seq dataset.

Figure 3.

Effect of AMH on MII oocyte transcriptome. (A) Heatmap of top 100 significant DEGs sorted by adjusted P-value by plotting their log2-transformed expression values in MII oocytes isolated from control (C1–C3) and AMH treated (T1–T3) animals. (B) Volcano plot representing the global transcriptional change in oocytes isolated from control versus AMH-primed mice. Each data point in the scatter plot represents a gene. Genes with an adjusted P ≤ 0.05 and a log2 fold change ≥1 are indicated by red dots and represent upregulated genes. Genes with an adjusted P ≤ 0.05 and a log2 fold change ≤−1 are indicated by green dots and represent downregulated genes. (C) Relative expression of Uqcrq (ubiquinol–cytochrome C reductase complex III subunit VII), Cox7b (cytochrome C oxidase subunit 7B), Tfam (transcription factor A, mitochondrial), Rps13 (ribosomal protein S13), Sox18 (SRY-box transcription factor 18), Bmp2 (bone morphogenetic factor 2), Hif3a (hypoxia-inducible factor 3a), and Il17rb (interleukin 17 receptor B) mRNA levels by quantitative PCR in oocytes isolated from AMH-primed and vehicle-treated (control) mice. Data are represented as mean ± SEM (n = 4 samples; each sample contained 50 oocytes pooled from 2 mice and a total of 8 mice/treatment) and normalized to Gapdh (P = 0.0102, two-way ANOVA).

GO enrichment analysis (Figure 4) of the DEGs reveal mitochondrial pathways to be highly enriched in all three components (biological processes, molecular pathways, and cellular components) analyzed. These enriched pathways include genes like Ndufs6, Uqcrq, Cox7c, Uqcrh, Park7, Coq9, Uqcc3, and Sdhd responsible for ATP synthesis in the electron transport chain; genes like Atp5l, Ndufa11, Ndufs6, Ndufb4, Atp5j2, and Cox7c encoding proteins of the inner mitochondrial membrane regulating ATP synthesis; and genes like Polg2, Dalrd3, Cars2, Ttll3, Alox15, Ddt, Ppia, Hpgds, and Ppil3 involved in the regulation of mtDNA accumulation during oocyte growth and increase in mtDNA copy number. Moreover, genes involved in lipid oxidation (Acox2, Alox15, Alox12e, Fabp3, Alox5, Appl2, and Mlycd) and fatty acid beta-oxidation (Acox2, Mlycd, Mtln, Hsd17b10, Gcdh, Acaa2, Hadha, Eci1, Etfb, and Hadh) that promote relocalization of lipids during oocyte maturation improving embryo development as well as genes involved in cytoskeletal changes (Dctn2 and Tubb5) important in the growth, maturation, and fertilization of oocytes are upregulated in oocytes isolated from AMH-primed compared with control animals. These upregulated pathways/genes reveal that AMH affects a wide range of molecular and biological processes critical for oocyte development and thus has a far-reaching influence in female fertility.

Figure 4.

Gene ontology (GO) enrichment analysis. GO terms of significantly enriched pathways with adjusted P-values <0.05 in the DEG sets. The numbers on each bar represent the number of DEGs in the corresponding pathways. Important mitochondrial pathways have been selected and their corresponding genes have been used to show their relative fold change. MF, molecular function; CC, cellular component; BP, biological pathways; DEGs, differentially expressed genes.

AMH enhances mitochondrial functions in mouse oocytes

The primary role of mitochondria is to produce energy, in the form of ATP, to fuel cellular processes, with approximately 95% of ATP being generated by the oxidative phosphorylation pathway [34]. Mammalian oocytes have limited capacity of glycolysis, which makes oocytes rely heavily on mitochondria for energy-intensive processes like growth, maturation, fertilization, and subsequent embryo development [35]. Given that our RNA-seq analysis highlights several genes involved in the mitochondrial pathways to be highly enriched, we investigated mitochondrial distribution and function in oocytes isolated from AMH-primed and vehicle-treated control mice. In oocytes and two-celled embryo isolated from control and AMH-treated mice, there was no difference in the distribution of mitochondria based on the homogeneous distribution of Mitotracker staining (Figure 5A and B) throughout the cytoplasm. Interestingly, although there was no difference in the intensity of Mitotracker staining in oocytes (Figure 5A), the mtDNA copy number (Figure 5E, P = 0.0045) and mtDNA/nDNA ratio were significantly higher in oocytes isolated from AMH-primed versus vehicle-treated mice. In contrast, two-celled stage embryos (Figure 5B, P = 0.010) derived from oocytes isolated from AMH-primed animals show higher level of staining, signifying more mitochondria than in the control. Moreover, mitochondrial membrane potential measured by TMRM staining in both oocytes (Figure 5C, P = 0.0001) and two-celled embryos (Figure 5D, P = 0.010) was significantly higher in AMH-primed versus vehicle-treated animals. This increased mitochondrial membrane potential represents abundant active mitochondria which is further supported by high ATP levels (27.94 ± 1.36 versus 19.31 ± 0.85 pg/oocyte, P = 0.0012) in oocytes isolated from AMH-primed compared to the control mice (Figure 5G).

Figure 5.

AMH priming has a positive effect on mitochondrial function in mouse oocytes. (A, B) MitoTracker red staining representing mitochondrial distribution in (A) oocytes (n = 10 oocytes from 4 different animals/treatment; Scale bars: 50 μm; P = ns using t-test) and (B) two-celled embryo (n = 4 embryos from 4 different mice/treatment; Scale bars: 20 μm; P = 0.010 using the t-test) isolated from AMH-primed and vehicle-treated (control) mice. (C, D) TMRM staining representing mitochondrial membrane potential in (C) oocytes (n = 10 oocytes from 4 different animals/treatment; Scale bars: 50 μm; P = 0.0001 using the t-test) and (D) two-celled embryo (n = 4 embryos from 4 different mice/treatment; Scale bars: 20 μm; P = 0.010 using the t-test) isolated from AMH-primed and vehicle-treated (control) mice. Tables represent quantification of fluorescence intensity per oocyte and embryo. Data are represented as mean ± SEM of relative intensity (au) and ^*P < 0.05 was considered significant. (E, F) Mitochondrial DNA (mtDNA) content represented as the ratio of mitochondrial to mtDNA/nuclear DNA (E) and (F) mtDNA copy number in oocytes isolated from AMH-primed versus vehicle-treated (control) mice. Data are displayed as mean ± SEM (n = 5 mice and 5 oocytes pooled/animal, P = 0.0045 using the t-test). (G) ATP levels in oocytes isolated from AMH-primed compared with control mice. Data are displayed as mean ± SEM (n = 5 mice; 10 oocytes pooled/mice, P = 0.0012 using the t-test).

Discussion

AMH has long been considered a biomarker for ovarian function. Only recently, the perception that AMH can be used as a therapeutic option and a target to improve ovarian function and female fertility is being contemplated. This hypothesis is based on studies in mouse models that show AMH treatment creates a quiescent ovarian environment [36] and, AMH pretreatment prior to superovulation can increase the number of ovulated oocytes [16]. Moreover, clinical data show a positive correlation between AMH levels and the number of oocytes retrieved during an IVF cycle [37–40]. However, the knowledge of how this AMH-induced quiescent ovarian environment affects oocyte quality, early embryonic development, and live birth rate are unknown. This study for the first time shows that in vivo AMH treatment in mice has no negative effect on oocyte and early embryonic development. In fact, more oocytes and embryos obtained following superovulation from AMH-induced quiescent ovary are of better quality, a characteristic further reflected by more pups born from AMH-primed versus unprimed animals.

Oocyte quality can be defined as its ability to be fertilized, mature, and give rise to normal offspring [41]. Whether serum AMH level is a predictor of oocyte quality is debatable [42]. A number of clinical studies have reported a positive correlation between serum AMH levels and oocyte quality, embryo development, and establishment of clinical pregnancy [37, 38, 43–45]. In contrast, others have shown that AMH has no role in predicting oocyte quality and IVF outcome [46]. However, this study highlights that exogenous AMH treatment has no toxic or deleterious effects on ovarian function and fertility. On the contrary, we find that in vivo AMH treatment not only increases oocyte yield but these oocytes also have higher competence with respect to their ability to become fertilized, form two-celled-stage embryos and develop to blastocysts. Moreover, given that these blastocysts have higher level of SOX2- and NANOG-positive cells, which are markers of pluripotency in inner cell mass [25, 26], we propose that these blastocysts are potentially of better quality. These observations are in accordance with in vitro studies that have previously reported that AMH supplementation enhanced oocyte competence/quality and improved embryonic development in nonhuman primates [47], mice [19], and humans [48]. Further, the fact that AMH-primed animals give birth to more pups supports our hypothesis that exogenous AMH treatment in general has a positive effect on overall female fertility.

At the molecular level, as per our knowledge, this is the first study that shows the genes and pathways that are affected in the MII oocyte with exogenous AMH treatment. The fate of an embryo is primarily dependent on the quality of the oocyte from which it is derived [49–51]. At the very early stages of embryogenesis, there is no transcription before embryonic genome activation, and so the first steps of embryogenesis are exclusively dependent on the maternally inherited genes from the oocyte [49–51]. Thus, there have been several studies correlating oocyte gene expression as a measure of oocyte quality with successful embryonic development and implantation [52–54]. For example, mitochondrial function and mtDNA content are major markers of oocyte quality [55] and have been associated with reproductive aging [56]. Moreover, embryo development is also strongly correlated with the activity of mitochondria in oocytes [57]. Mitochondrial number also increases dramatically throughout oogenesis and folliculogenesis such that mature oocytes can contain 100 000 mitochondria and 50 000–1 500 000 copies of the mitochondrial genome [58]. The fact that a large number of genes associated with mitochondrial function are upregulated in oocytes isolated from AMH-primed animals along with the increased number of active mitochondria, higher ATP level and mtDNA content in these oocytes provide the basis for the improved quality of oocytes observed in these animals. Additionally, genes involved in pluripotency, lipid metabolism, and cytoskeleton dynamics are also upregulated in oocytes isolated from AMH-primed animals. Therefore, this study provides a molecular insight into how AMH can influence oocyte quality and competence, the underlying mechanism of which remains to be elucidated. The effect of AMH may vary with age, in a follicle-type-specific and cell-specific manner. For example, a previous study [59] focused toward understanding how AMH inhibits GC activation and differentiation thereby halting follicle growth to preserve ovarian reserve had used single-cell RNA sequencing of whole neonatal mouse ovaries treated with AMH at birth and analyzed at postnatal day 6. This study [59] reported significant changes in the transcription signatures in GCs related to GC proliferation and differentiation following AMH treatment. However, there was only a modest change (only three DEGs) in the oocyte transcriptome. The primary reason proposed for this modest effect of AMH on oocyte transcriptome is the lack of AMH receptor 2 (AMHR2) expression in the oocyte [59]. However, these studies [59] were done in neonatal ovaries that primarily contained primordial, primary, and secondary follicles. In contrast, studies [19] in COCs and MII oocytes isolated from 6–8-week-old mice following superovulation have reported that AMHR2 is expressed in the oocytes and cumulus cells before and after in vitro maturation. Moreover, AMHR2 is also expressed in oocytes isolated from bovines [60], rats [61], and humans [48, 62]. Therefore, AMHR2 expression in oocytes may vary with respect to follicle type and among species. Further investigation is needed to elucidate whether the effects of AMH on oocyte transcriptome is direct, through AMHR2, or indirect, through its effect on GCs. The importance of bidirectional signaling between the oocyte–GC in ensuring oocyte competence and development is well established [63] and GCs have been shown to modulate the transcriptional activity of the oocyte genome [64]. Therefore, the indirect effects of AMH on oocytes through its actions in GCs also cannot be discounted. For example, previous studies [36] have shown that in GCs [16], AMH treatment blocks FSH actions by inhibiting FSH-induced cAMP and aromatase and lowering estradiol levels [16]. The lower estradiol levels cause an increase in FSH levels in the AMH-treated animals. It is possible that some of the observed effects on oocytes may be attributed to this AMH-induced FSH increase. Studies have reported that FSH can promote oocyte growth [65] as well as mouse oocyte meiotic resumption [66].

In summary, this study shows the effect of AMH on oocyte quality and early embryonic development. Here we report that in vivo AMH treatment of mice not only increases the number of ovulated oocytes but also has a positive effect on oocyte maturation and early embryonic development with more live births. This phenotype is further supported by the fact that AMH priming upregulates the expression of genes in oocytes associated with oocyte quality and competence.

Data availability statement

The RNA-seq data underlying this article are available in the Gene Expression Omnibus GSE195720 at https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE195720.

Conflict of interest

A.S. is a co-owner of a US patent related to therapeutic use of AMH for regulation of fertility. The remaining authors have nothing to disclose.