Anti-Mullerian hormone (AMH) protects ovarian follicle loss by downregulating granulosa cell function in in vitro and in vivo models.

Purpose AMH inhibits hormone production in luteinized granulosa cells (GCs) and stalls ovarian follicle development in vitro and in vivo. We sought to confirm AMH’s mechanism of action through SMAD activation and investigate AMH inhibition of follicle development and function, in vitro and in vivo.

A primary culture of GCs isolated from follicular fluid was used, and cells were treated with recombinant AMH (rAMH) or placebo for 24 h. For the mouse model, 18-weeks old C57BL female mice were either euthanized at the beginning or treated with rAMH or normal saline for 3 weeks. Primordial (PDF), primary follicle (PRF), secondary (SEF), and tertiary follicles (TEF) were calculated. Real-time RT-PCR and ELISA were performed to quantify GC gene expression and protein translation of human SMAD 1, 5, and 8, FSH-R and mouse FSH-R, inhibin B, caspase 3, Ki67, BMP15, GDF9, and the epigenetic regulators miRNAa and b.

In vitro, rAMH-treated GC showed activation of the SMAD 1, 5 and downregulation of SMAD 8, with greater magnitude at increasing rAMH doses (p < 0.04) and consequential control of downstream regulators. In vivo, the rAMH-treated mice showed increased SEFs and decreased PRFs while PDFs, TEFs, were unchanged compared with baseline. Compared with Placebo, the rAMH group showed increased PDFs, while PRFs, and TEFs were significantly decreased, and SEFs were unchanged.

AMH caused SMAD activation in a dose-dependent manner, with downstream downregulation of cell function and replication, also through activation of miRNAs. These mechanisms were confirmed by the in vivo findings with ultimate downregulation of follicular development and preservation of the ovarian follicle number. Counteracting follicular depletion, AMH could be used to protect the ovarian follicle reservoir.

AMH, AMH-R2, Ovary, Ovarian follicle, Fertility preservation

Ovarian aging is expressed by follicular depletion and is quantified by decreasing Anti-Mullerian hormone (AMH) and increasing follicular stimulating hormone (FSH) [1]. The mechanisms of ovarian aging are multiple and only partially understood, but AMH is considered a key regulator of this process. In fact, in naturally aging women, AMH was shown to maintain the follicle pool within the ovaries by inhibiting the activation of the primordial follicles and diminishing the number of growing follicles [2]. Additionally, women with polycystic ovary syndrome (PCOS) have higher AMH levels and go into menopause at a later time than women without PCOS [3]. AMH is a member of the transforming growth factor-beta gene family. Its expression is restricted in the male to the Sertoli cells of fetal and postnatal testis, and in the female to the granulosa cells (GC) of the ovary. Its production starts at the stage of primary follicles, soon after the columnar differentiation of the flat GC of primordial follicles [4]. AMH exerts a paracrine inhibitory function in the activation and development of primordial follicles into developing follicles in animal models. In the human, AMH production is maximum in secondary follicles and early antral tertiary follicles, and gradually decreases in antral tertiary follicles greater than 8 mm [5]. Figure 1 depicts the process of folliculogenesis and the effects of age-related follicle number and AMH production decline. In engineered AMH-null mice, the ovaries contain almost three-fold greater number of growing secondary follicles and a lower number of primordial follicles [6]. In fact, in the absence of AMH, primordial follicles are recruited at a faster pace [7]. Further, in granulosa cell cultures, AMH attenuates the FSH (follicle-stimulating hormone)-dependent increase in aromatase activity and LH (luteinizing hormone) receptor expression, thus inhibiting FSH-dependent follicular growth [8]. Thus, in animal models AMH appears to downregulate two important steps of follicular development: follicle recruitment and cyclic selection for dominance [4, 6, 8]. Its action is mediated by two receptors, where AMH-R2 is the first binding AMH and AMH-R1 is secondarily activated. Activation of SMAD 1, 5, and modulation of SMAD 8 follows AMH-R2 and AMH-R1 binding, producing downstream gene expression and protein translation [9]. Over the past decade, our group has gathered undisputable evidence that supports the key role of AMH as a regulator of ovarian cortex function [10–12]. In fact, recombinant AMH (rAMH) reduced tissue expression of AMH itself, inhibin B, AMH-R2, FSH-R (FSH receptor), LH-R (LH receptor), and IGF1-R1 (insulin-like growth factor 1-receptor 1) in human ovarian cortex [10]. Additionally, rAMH also significantly reduced expression of AMH, AMH-R2, FSH-R, and inhibin B in human luteinized GC and remarkably reduced cell proliferation and apoptosis [11]. In an in vivo mouse model, rAMH inhibited follicle pre-activation and burn out after xenotransplantation of human ovarian cortex into nude mice, downregulating ovarian hormone production [12]. These findings were later confirmed by other investigators where rAMH was found to protect ovarian follicle reserve during chemotherapy treatment [13, 14]. Also, rAMH was described to preserve ovarian follicles from depletion while used as contraception during chemotherapy treatment in mice [15]. Cell function and differentiation is regulated also by epigenetic regulators such as micro-RNAs (miRNA), which inhibit translation of multiple RNAs within specific cell-types. MiRNA181 is a cluster of miRNAs that was identified to elicit different effects within different human cell types where it controls many processes including differentiation, proliferation, inflammation, and metabolism [16]. In fact, higher miRNA181a-b levels enhanced differentiation and proliferation in human embryonic chondroblasts [17], adipocytes [18], vascular [19], and myocardial cells [20], while in granulocytes and macrophages, they seem to inhibit the same functions [21]. Additionally, mouse studies showed a critical role of miR- 181 in regulating growth and development, where knock-out mice for each of the miR- 181 clusters showed a reduction in size, body weight, and viability [22]. Presently, it is not known whether AMH controls these epigenetic regulators, and it would be interesting to find a relationship between hormonal and post-transcription regulation of ovarian follicle function. The objective of these studies was to assess a dose-dependent effect of AMH in the activation of the SMAD cascade in luteinized GC in vitro, with consequent downregulation of cellular function also through activation of the epigenetic regulators, miRNA181 s. In addition, we sought to confirm AMH’s effects in vivo and assess its efficacy in preserving the ovarian follicle reserve in the mouse, in vivo.

The different studies here described were approved by an internal ethics review board at IVF Michigan Rochester Hills and Flint, PC, in 2017 and by the University of Tennessee Health Science Center (UTHSC) Institutional Animal Care and Use Committee (IACUC) in 2015, protocol No. 14–066.0. Mechanism of action and dose-related effects of AMH on GC To confirm that AMH acts through activation of the SMAD signaling pathway, we measured SMAD 1, 5, and 8 in a primary culture of luteinized GC created back in 2018 and cryopreserved. The study was originally approved by an internal ethics review board at IVF Michigan Rochester Hills and Flint, PC, in 2017. To create the primary GC culture, GCs were isolated from follicular aspirates and centrifuged at 900 × g after blood contaminants were removed by Histopaque® 1077 (Sigma-Aldrich, St. Louis, MO, USA) as previously reported [11]. Following further washing, the cell stock was resuspended in the GC culture medium and frozen for future use. After thawing, GC was seeded in well culture plates at a density of 100,000 cells/well in medium. Following an overnight incubation (37 °C, 5% CO2), cells were treated with recombinant AMH 4.0 ng/ml (R&D Systems, Inc., Minneapolis, MN, USA; rAMH group), or phosphate-buffered saline (PBS, control group), for 24 h. The rAMH 4.0 ng/ml dose was chosen to resemble the average human plasma AMH concentration. Total RNA was extracted from all cells and subjected to real-time RT-PCR to quantify SMAD 1, 5, and 8 gene expression. Additionally, total protein was extracted and subjected to ELISA to quantify SMAD 5 and 8/9 proteins in GC cultured in increasing rAMH doses of 0.4 ng/ml and 0.6 ng/ml. ELISA kit for SMAD 1 was not commercially available. AMH stalls ovarian follicle development and oocyte function in a mouse model To investigate whether AMH can inhibit follicular development in vivo, we used 18 18-week-old C57BL female mice that were assigned to three treatments: baseline (euthanized just prior to the experiment); rAMH (recombinant AMH, 1.8 µg/day); and placebo group (normal saline), via intraperitoneal pumps. The AMH dose was chosen based on our group’s previous study where a daily dose of 1.23 µg/day produced effective results with the same administration method [12]. The study was approved by the UTHSC Institutional Animal Care and Use Committee (IACUC) in 2015, protocol No. 14–066.0, which assured animal welfare. Mice were housed in the Animal Center at UTHSC under a 12-h light/dark cycle with food and water ad libitum. Mice were euthanized 3 weeks after pump placement, and the ovaries were explanted for histological analysis. Primordial (PDF), primary (PRF), secondary (SEF), and tertiary follicles (TEF) were counted in the ovarian specimens. In addition, PCR expression of Ki67 (indicator of cell proliferation), caspase 3 (indicator of apoptosis), inhibin B, BMP15, and GDF9 were measured. After dissection, a portion of the ovarian specimen was fixed in formalin, embedded in paraffin, and stained with hematoxylin and eosin (H&E) and with immune stains. The H&E-stained slides were used to evaluate the follicle number, while the immune-stained slides were used for examination of follicular function and development. Five-µm-thick sections were serially cut, and every fifth section analyzed for follicular counts. At least three ovaries from different animals were obtained from each group, and at least four sections were analyzed in each ovary. The computerized program used for image analysis, Spectrum (Version 10.2.2.2314; by Aperio, Vista, CA, USA), allowed us to concomitantly examine four tissue Sects. 25 µm apart (every fifth section) and to count all the follicles in each section without redundancy accurately. For follicle counting, the investigators were blinded to the treatment groups. One oocyte surrounded by a monolayer of flat or cubic GC enclosed by a basement membrane identified a primordial (PDF) and a primary follicle (PRF), respectively. Secondary follicles (SEF) were identified with one oocyte surrounded by multiple cubic GC, and tertiary follicles (TEF) by fluid accumulation amid the GC enclosed by a basement membrane. The total number of PDF, PRF, SEF, and TEF, and corpora lutea in each section was divided by the section area to calculate the number of follicles/mm3. Real-time RT-PCR analysis We utilized real-time RT-PCR to determine tissue mRNA levels for all markers under investigation in each experiment, adhering to the following protocol. Total RNA was extracted from tissues with the RNeasy Mini Kit (Qiagen). Frozen tissues were first homogenized in lysis buffer provided in the kit and run through a QIA tissue shredder column (Qiagen) followed by RNA extraction according to the protocol provided by the manufacturer. A 20-μL cDNA reaction volume using an equal amount of RNA (up to 1 µg) was prepared using the SuperScript VILO MasterMix Kit (Life Technologies, Grand Island, NY), as described by the manufacturer’s protocol. Real-time RT-PCR was performed with the Express SYBR GreenER qPCR SuperMix RT-PCR kit (Life Technologies) and a Cepheid 1.2f Detection System (Cepheid, Sunnyvale, CA). Each 25-μL reaction included 12.5 µL of 2 × QuantiTect SYBR Green RT-PCR master mix, cDNA template, and 0.2 µM each of target-specific primer were selected with the aid of the software program, Beacon Designer (Premier Biosoft, Palo Alto, CA). Human oligonucleotide primers that amplify variable portions of the protein coding regions were established and verified, described in Table 1, as previously described [10–12]. Standards with known concentrations were designed specifically for β-actin (79 base pairs (bp) and all the markers under investigation in each experiment using the Beacon Designer software, allowing for construction of a standard curve using a tenfold dilution series. An individual standard for each gene of interest provides a method for absolute quantification of the gene in interest. The PCR reaction conditions were programmed as follows: an initial cycle was performed at 95 °C for 60 s. Next, there were 35 cycles of denaturation at 95 °C for 15 s, annealing time, and temperatures, followed by a final cycle at 72 °C for 30 s to allow completion of product synthesis. A melting curve analysis was performed to demonstrate the specificity of the PCR product as a single peak. A control, which contains all the reaction components except for the template, was included in all experiments. All PCR analyses were executed in triplicates. Table 1. | Gene | Sense (3′− 5′) | Antisense (5′− 3′) | Amplicon length (bp) | Annealing temperature (°C) | Annealing time (sec) | |---|---|---|---|---|---| | B-actin | ATGACTTAGTTGCGTTACAC | AATAAAGCCATGCCAATCTC | 79 | 58 | 10 | | AMH | GTGCTGCTGCTGAAGATG | CTCCGACAGGCTGATGAG | 102 | 62 | 10 | | AMH-R2 | CCAGAAGCACGGCTGACAG | TGGAAAGGGGTGGCTCTCT | 81 | 62 | 10 | | Ki67 | TCCTTTGGTGGGCACCTAAGACCTG | TGATGGTTGAGGCTGTTCCTTGATG | 156 | 55 | 30 | | SMAD 1 | CTGCATGTGTATTCGTGAGTTCGC | AACCCAGTCAGCACAAAGACA | 76 | 72 | 63 | | SMAD 5 | CAGGAGTTTGCTCAGCTTCTGG | GGTGCTGGTTACATCCTGCCG | 141 | 72 | 62 | | SMAD 8/9 | AGTGGCCAACCTGTAGATGC | TGGGGCTCCTCGTAACAAAC | 92 | 63 | 72 | | GDF9 | TCGCATTACTACCGTTGAA | CACACATTTGACAGCAGAG | 91 | 53 | 10 | | BMP 15 | TCAAGCCAAACACAAACAGC | AGCCACAATCCAGTCATTCC | 92 | 54 | 10 | | FSH-R | GTCCACAACACCCATCCAAGG | GGGCTAAATGACTTAGAGGGACAA | 97 | 66 | 10 | | miRNA181a1 | GTGAACATTCAACGCTGTCGG | CATAGGGTACAATCAACGGTCG | 70 | 56 | 72 | | miRNA181a2 | GGAACATTCAACGCTGTCGG | TAAGGACCCCAAGGTACAGT | 74 | 57 | 72 | | miRNA181b1 | AACATTCATTGCTGTCGGTGGG | GGGGCCACAGTTGCATTCATTG | 71 | 62 | 72 | | miRNA181b2 | CATTCATTGCTGTCGGTGGG | TTTGGTCCGCAGTTTGCATT | 70 | 56 | 72 | ELISA To validate the gene expression study results, we performed ELISA on GC and mouse ovary lysates to quantify protein concentration of FSH receptor (FSH-R), SMAD 5, SMAD 8/9, caspase 3, Ki67, and the two oocyte-derived hormones that regulate GC’s mitotic proliferation and differentiation, BMP15 and GDF9 [23, 24]. A specific antibody for the target protein was pre-coated onto a microplate in sandwich ELISA kits. Standards and samples were added into the wells and the immobilized antibody binds any target protein present. After washing away any unbound substances, an enzyme-linked polyclonal antibody specific for the target protein was added followed by the addition of a substrate solution and color was developed in proportion to the amount of target protein bound. The color development was stopped, and the intensity of color was measured at 450 nm. For human FSH-R we utilized sandwich Colorimetric ELISA (ThermoFisher Scientific, catalog number: EH202RB) as described in the manufacturer protocol. For SMAD 5 and SMAD 8/9, we utilized sandwich enzyme immunoassay for in vitro quantitative measurement (MYBioSource, SMAD 5, MBS9714237, and SMAD 8/9, MBS9714235), as described in the manufacturer protocol. For mouse FSH-R, we utilized sandwich Colorimetric ELISA purchased from Bioassay technology laboratory, as described in the manufacturer protocol. For Ki67, we utilized sandwich Colorimetric ELISA (MYBioSource, catalog number: MBS1601117) as described in the manufacturer protocol. For mouse BMP15, we utilized sandwich enzyme immunoassay for in vitro quantitative measurement (ABclonal Technology, catalog number: RK08181) as described in the manufacturer protocol. For GDF9, we utilized sandwich enzyme immunoassay for in vitro quantitative measurement (Elabscience, catalog number: E-EL-M0601) as described in the manufacturer protocol. Statistical analysis All our variables were continuous, however, because of the skewed distribution, our results were summarized as mean ± standard deviation and median and quartiles (Q1, Q3). Kruskal–Wallis Test was used to compare the groups, where indicated. For blood serum levels, we used the same test to evaluate the changes between the placebo and rAMH groups of mice at the various time-points. Comparisons between groups were made using SPSS (IBM SPSS Statistics v29, Somers, NY 10589, USA). A p-value < 0.05 indicated a statistically significant difference.

Mechanism of action and AMH Because AMH acts by activating SMAD 1, 5, and modulating SMAD 8, we measured gene expression of FSH-R, SMAD 1, 5, 8, miRNA181a1, miRNA181b1, miRNA181a2, and miRNA181b2, in a primary culture of luteinized GC (GC) after adding rAMH 4.0 ng/ml (rAMH group), or phosphate-buffered saline (PBS, control group), for 24 h. Table 2 shows the results of real-time PCR analysis. As expected, expression of FSH-R was significantly reduced by rAMH, compared to the control group (p ≤ 0.030), consistent with our previous findings [8]. In AMH-treated cells, we also observed activation of the SMAD 1, 5, and 8 cascade (collective p < 0.024) and a differential down-regulation of miRNA181a1 (p < 0.004) and miRNA181b1 (p < 0.005) and up-regulation of miRNA181a2 (ns) and miRNA181b2 (p < 0.039) (Table 2). Table 2. | Marker | Untreated (± SD) | rAMH 0.4 ng/ml (± SD) | p-value* | |---|---|---|---| | FSH-R (Fg/μg RNA) | 66.17 ± 3.45 | 51.96 ± 8.49 | 0.030 | | SMAD 1 (Fg/μg RNA) | 1.27 ± 0.25 | 2.96 ± 0.48 | 0.024 | | SMAD 5 (Pg/μg RNA) | 377.48 ± 3.71 | 505.62 ± 17.25 | 0.016 | | SMAD 8 (Fg/μg RNA) | 676.39 ± 31.34 | 432.54 ± 19.41 | 0.022 | | miRNA181a1 (Fg/μg RNA) | 24.68 ± 1.20 | 19.04 ± 0.74 | 0.004 | | miRNA181a2 (Fg/μg RNA) | 510.43 ± 15.93 | 527.68 ± 14.86 | ns | | miRNA181b1 (Fg/μg RNA) | 155.34 ± 3.33 | 129.59 ± 5.14 | 0.005 | | miRNA181b2 (Fg/μg RNA) | 422.04 ± 23.25 | 465.26 ± 11.64 | 0.039 | * = Kruskall-Wallis’s test for comparison of medians between untreated and rAMH, groups To corroborate the gene expression results with protein translation, we evaluated a dose-dependent GC response to AMH, assessing protein levels of FSH-R and SMAD 5, and 8/9. These markers were measured in GC cultured in increasing rAMH doses of 0.4 ng/ml and 0.6 ng/ml. As shown in Table 3, increasing the rAMH dose, FSH-R and SMAD 5 translation was significantly enhanced. Differently from the real-time RT-PCR results, also SMAD 8/9 tissue protein synthesis was stimulated by rAMH. Table 3. | Marker | Untreated (± SD) | rAMH 0.4 ng/ml (± SD) | rAMH 0.6 ng/ml (± SD) | p-value* | |---|---|---|---|---| | FSH-R (Pg/μg) | 54.98 ± 1.71 | 33.24 ± 0.32 | 26.44 ± 0.64 | 0.009 | | SMAD 5 (Pg/μg) | 27.18 ± 7.19 | 74.21 ± 0.21 | 142.07 ± 3.84 | 0.003 | | SMAD 8/9 (Pg/μg) | 10.42 ± 6.80 | 49.49 ± 3.56 | n/a** | n/a | * = Kruskall-Wallis’s test for comparison of medians between rAMH 0.4 and 0.6 ng/ml groups ** = not enough protein quantity to perform triplicate tests AMH stalls ovarian follicle development and oocyte function in a mouse model rAMH (recombinant AMH, 1.8 µg/day) and normal saline (placebo group) were continuously administered for 3 weeks via intraperitoneal pumps to 18-weeks-old (or mature) C57BL female mice, equivalent to 3 menstrual cycles. After euthanasia, we measured the number of ovarian follicles in all their developmental stages (from primordial to primary, secondary, and tertiary follicles), in addition to measuring PCR expression of Ki67 (stimulates cell proliferation), caspase 3 (apoptosis), and inhibin B. Table 4 reports the individual groups’ data. Compared with the placebo group (21 weeks’ age mice), the rAMH-treated groups showed higher PDFs, while PRFs and TEFs were significantly lower, and SEFs were unchanged. Figure 2 provides a graphical representation of the ovarian follicles’ distribution in the three groups and shows that rAMH treatment preserved PDF, SEF, and TEF, at baseline levels (mice euthanized at the beginning of the experiment, at 18 weeks’ age), while it caused a decrease in PRF due to the diminished PDF activation. RAMH treatment similarly caused decreased ovarian cortex expression of Ki67, caspase 3, and inhibin B expression. In addition, rAMH maintained baseline expression of the two oocyte-derived growth factors BMP15 and GDF9, which further confirms GC quiescence. To validate the gene expression study results, we performed ELISA on mouse ovary lysates to quantify protein concentration of the FSH receptor (FSH-R), Ki67, BMP15, and GDF9, as reported in Table 4. Compared with the baseline and placebo groups, AMHR2BP and rAMH administration caused a significant decrease in concentration of all the parameters under investigation, confirming the real-time RT-PCR results and the mechanisms by which the ovary inhibits its own cellular function through the activation of AMH-R2. Table 4. | Variable | Baseline median (Q1, Q3) | Placebo group median (Q1, Q3) | rAMH group median (Q1, Q3) | p-value* | |---|---|---|---|---| | PDF/mm3 | 1195 (1191, 1248) | 614 (538, 705) | 1182 (1095, 1248) | 0.011 | | PRF/mm3 | 788 (670, 817) | 1165 (1152, 1176) | 514 (497, 548) | 0.016 | | SEF/mm3 | 970 (968, 1061) | 1613 (1528, 1664) | 1144 (1119, 1300) | ns | | TEF/mm3 | 583 (579, 606) | 1082 (1076, 1119) | 871 (808, 890) | 0.045 | | Corpora Lutea/mm3 | 6.0 (5.0, 6.0) | 4.0 (2.0, 5.0) | 7.5 (6.3, 8.0) | 0.004 | | Real-Time RT-PCR | |||| | Inhibin B (pg/µg RNA) | 34.53 (34.53, 34.54) | 48.41 (47.36, 49.47) | 4.07 (3.86, 4.27) | 0.001 | | Ki67 (pg/µg RNA) | 22.84 (20.49, 25.21) | 69.26 (67.40, 71.12) | 8.05 (7.20, 8.89) | 0.005 | | Caspase 3 (pg/µg RNA) | 1.31 (1.27, 1.36) | 3.10 (3.00, 3.20) | 0.46 (0.43, 0.50) | 0.004 | | BMP15 (fg/µg RNA) | 206.9 (153.8, 976.6) | 4007.3 (2875.7, 5573.0) | 116.1 (101.9, 800.3) | 0.005 | | GDF9 (fg/µg RNA) | 49.1 (28.3, 61.1) | 387.4 (355.2, 419.9) | 21.5 (12.8, 33.1) | 0.013 | | ELISA | |||| | FSH-R (ng/µg protein) | 36.6 (31.0, 43.1) | 39.0 (32.0, 40.5) | 14.6 (13.9, 16.2) | < 0.001 | | Ki67 (ng/µg protein) | 149.1 (131.7, 164.7) | 148.6 (134.8, 154.3) | 68.0 (65.2, 78.8) | 0.001 | | BMP15 (ng/µg protein) | 22.6 (20.1, 30.8) | 36.0 (33.9, 47.7) | 6.3 (4.0, 11.2) | < 0.001 | | GDF9 (ng/µg protein) | 17.4 (13.7, 25.0) | 19.5 (16.0, 26.3) | 3.2 (2.9, 6.7) | < 0.001 | * = Kruskall-Wallis’s test for comparison of medians between baseline, placebo, and rAMH groups

The experiments reported in this study confirm AMH as a key regulator of ovarian follicle function and longevity. In vitro, rAMH showed activation in gene expression of SMAD 1, 5, downregulation of SMAD 8, and differential miRNA181 regulation in GC. Additionally, we established a dose-dependent regulation of FSH-R and SMAD 5 translation in response to AMH. An interpretation of the conflicting decrease in SMAD 8 gene expression and an increase in SMAD 8/9 protein synthesis by ELISA after administration of rAMH is elusive. However, because of similar FSH-R and SMAD 5 trends, we believe this discrepancy was due to the unavailability of an individual SMAD 8 ELISA kit and that the increased SMAD 8/9 protein synthesis was principally due to an upregulation of SMAD 9. The significance of SMAD 9 upregulation by AMH remains unknown; however, acting as an effector of BMP1 signaling pathway, we speculate SMAD 9 should be involved in transcription regulation [25]. The presence of miRNA181 in GC has never been reported before and the current study results on AMH’s regulation of miRNA181a and b are novel. Because AMH has been shown to bind only to the AMH-R2 and 1 complex and because in the adult female this is found on ovarian epithelial cells as well as other reproductive organs, such as the ductal epithelium of the mammary gland and the endometrium, we assume that miRNA181a and b portray their effects also on these reproductive organs. However, the effects of AMH on up- and down-regulating miRNA181a and b are currently unidentified. Based on the effects reported in other cell types [17–21] and the robust histology results in the mouse, we speculate that the observed differential down-regulation of miRNA181a1 and b1 and up-regulation of miRNA181a2 and b2 in this study further controls the regulation of GC differentiation, metabolism, and replication through post-transcriptional alterations. This mechanism would further strengthen the inhibitory effects of AMH on the ovarian cortex demonstrated in this and our previous studies [10–12]. In vivo, we established AMH’s hindering of cell proliferation as a function of an inhibited response to hormonal stimuli and a diminished hormonal production, confirming our previous studies [10–12]. In fact, while indirectly controlling pituitary stimulation of follicular growth by modulating GC’s FSH-R expression, AMH also exerts direct inhibition of cellular replication (demonstrated by Ki67 reduction) and prolongs follicle lifespan (demonstrated by caspase 3 reduction). AMH’s reduction of BMP15 and GDF9 validated its inhibition of granulosa and theca cell function, as well as of oocyte differentiation and evolution thus precluding its cytoplasmic and nuclear maturation [23, 24]. The fact that AMH and AMHR2BP can regulate BMP15 and GDF9 sheds a light on physiological mechanisms that govern the granulosa cell-oocyte interaction during follicular development. By minimizing the progression of follicular growth and development, and decreasing hormone production, cell replication and apoptosis, rAMH treatment preserved the ovarian follicle number and thus total ovarian reserve. In fact, rAMH treatment “locked” in time the number of PDF, and the larger-size follicles SEF and TEF by decreasing activation and proliferation, while it decreased PRF by inhibiting PDF activation. Combining our group’s previous reports on AMH function [10–12] with other investigators’ results of AMH protecting the ovary from chemotherapy damage by decreasing primordial follicle activation and “burnout” [13–15], the current results confirm AMH’s downregulation of ovarian cortex functions through dwindling of cell activities and proliferation, which ultimately enhance cellular and follicular steadiness and longevity. The strengths of the performed studies lie on the rigorous testing with consistent results in both in vivo and in vitro models as well as the evaluation of both gene activation and protein synthesis. An important limitation is that we have not assessed ovarian cortex or granulosa cell function after stopping AMH administration, blocking AMH effects with AMH-antibodies, or administering an AMH-R2 antagonist, such as the timing to return of normal function in human GC as well as mouse ovarian cortex. In this manuscript, we presented studies that demonstrate the mechanism of action of AMH. We also showed the efficacy of AMH on two levels: in vivo (in mice) and in vitro (in human luteinized GC), with two assessments—gene activation, translation, and protein synthesis. Through these studies, we confirmed that AMH inhibits replication, apoptosis, and hormonal production of GC in vitro. We also established that AMH prevents ovarian follicle loss and the progression of primordial to antral follicles in mice in vivo, which ultimately preserves the follicular reservoir for the length of administration. Previous results in primates seem to contradict our results in human luteinized GC, where AMH was found to promote folliculogenesis in vitro, while inhibiting antral follicle growth and the selection of the dominant follicle, in vivo [26]. However, in the in vitro portion by Xu et al., AMH effects were studied only on individually cultured secondary and small antral macaque follicles, thus preventing to draw definitive conclusions on the overall AMH’s effects. Together with previous studies, the current findings establish AMH as an interesting therapeutic choice for protection from physiologic (such as aging) and pathologic conditions (such as gonadotoxic therapy, endometriosis, and inflammation) that cause infertility and primary ovarian insufficiency by activation of primordial follicles and consequent follicular depletion [27–29]. Therefore, AMH could be classified as an innovative medical therapy where currently there is undisputable necessity. Author contribution LD and GMS conceived and designed research for the reported studies. LD and GMS conducted in vivo and in vitro experiments. GMS contributed all reagents and analytical tools. LD and MCM analyzed data and produced all tables and figures. LD, MCM, MPD, and GMS wrote the manuscript. All authors read and approved the manuscript. Funding This study was supported by an institutional grant (E07 - 3225–001) from the University of Tennessee Health Science Center, Memphis, TN (2016–2020), and by IRE Conceiving Future Families Philanthropy funding from Baylor College of Medicine-Texas Children’s Hospital, Houston, TX (2022–2024). Data availability The data described in this manuscript is kept in a private database and can be made available to the reviewers. Declarations Ethical approval The different studies here described were approved by an internal ethics review board at IVF Michigan Rochester Hills and Flint, PC, in 2017 and by the University of Tennessee Health Science Center (UTHSC) Institutional Animal Care and Use Committee (IACUC) in 2015. Conflict of interest The authors declare no competing interests. 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- 1.Gold EB. The timing of the age at which natural menopause occurs. Obstet Gynecol Clin North Am. 2011;38(3):425–40. 10.1016/j.ogc.2011.05.002. [DOI] [PMC free article] [PubMed] [Google Scholar] - 2.Pankhurst MW. A putative role for anti-Müllerian hormone (AMH) in optimising ovarian reserve expenditure. J Endocrinol. 2017;233(1):R1–13. 10.1530/JOE-16-0522. [DOI] [PubMed] [Google Scholar] - 3.Minooee S, Ramezani Tehrani F, Rahmati M, Ali Mansournia M, Azizi F. Prediction of age at menopause in women with polycystic ovary syndrome. Climacteric. 2018;21(1):29–34. [DOI] [PubMed] [Google Scholar] - 4.Durlinger ALL, Kramer P, Karels B, et al. Control of primordial follicle recruitment by anti-Mullerian hormone in the mouse ovary. Endocrinol. 1999;140:5789–96. [DOI] [PubMed] [Google Scholar] - 5.Jeppesen JV, Anderson RA, Kelsey TW, Christiansen SL, Kristensen SG, Jayaprakasan K, Raine-Fenning N, Campbell BK, Yding AC. Which follicles make the most anti-Müllerian hormone in humans? Evidence for an abrupt decline in AMH production at the time of follicle selection. Mol Hum Reprod. 2013;19(8):519–27. [DOI] [PubMed] [Google Scholar] - 6.Durlinger AL, Gruijters MJ, Kramer P, Karels B, Kumar TR, Matzuk MM, et al. Anti-Müllerian hormone attenuates the effects of FSH on follicle development in the mouse ovary. Endocrinology. 2001;142:4891–9. [DOI] [PubMed] [Google Scholar] - 7.Visser JA, Durlinger AL, Peters IJ, van den Heuvel ER, Rose UM, Kramer P, de Jong FH, Themmen AP. Increased oocyte degeneration and follicular atresia during the estrous cycle in anti-Müllerian hormone null mice. Endocrinology. 2007;148(5):2301–8. [DOI] [PubMed] [Google Scholar] - 8.Pellat L, Rice S, Dilaver N, Heshri A, Galea R, Brincat M, Brown K, Simpson ER, Mason HD. Anti-Müllerian hormone reduces follicle sensitivity to follicle-stimulating hormone in human granulosa cells. Fertil Steril. 2011;96:1246–51. [DOI] [PubMed] [Google Scholar] - 9.Josso N, di Clemente N, Gouedard L. Anti-Mullerian hormone and its receptors. Mol Cell Endocrinol. 2001;179:25–32. [DOI] [PubMed] [Google Scholar] - 10.Detti L, Fletcher NM, Saed GM, Peregrin-Alvarez I, Uhlmann RA. Anti-Müllerian Hormone (AMH) may stall ovarian cortex function through modulation of hormone receptors other than the AMH receptor. Reprod Sci. 2018;25(8):1218–23. 10.1177/1933719117737850. [DOI] [PubMed] [Google Scholar] - 11.Detti L, Abuzeid MI, Peregrin-Alvarez I, Christiansen ME, Malekzadeh P, Sledge J, Saed GM. Recombinant anti-Müllerian hormone (rAMH) for stalling in vitro granulosa cell replication. Reprod Sci. 2020;27:1873–8. [DOI] [PubMed] [Google Scholar] - 12.Detti L, Fletcher NM, Saed GM, Sweatman T, Uhlmann RA, Pappo A, Peregrin-Alvarez I. Anti-Mullerian Hormone (AMH) for prevention of ovarian follicle depletion after pre-pubertal ovarian cortex xenotransplantation. J Assist Reprod Genet. 2018;35:1831–41. 10.1007/s10815-018-1260-z. [DOI] [PMC free article] [PubMed] [Google Scholar] - 13.Roness H, Spector I, Leichtmann-Bardoogo Y, Savino AM, Dereh-Haim S, Meirow D. Pharmacological administration of recombinant AMH rescues ovarian reserve and preserves fertility in a mouse model of chemotherapy, without interfering with anti-tumoral effects. J Assist Reprod Gen. 2019;36:1793–803. [DOI] [PMC free article] [PubMed] [Google Scholar] - 14.Sonigo C, Beau I, Grynberg M, Binart N. AMH prevents primordial ovarian follicle loss and fertility alteration in cyclophosphamide-treated mice. FASEB J. 2019;33(1):1278–87. 10.1096/fj.201801089R. [DOI] [PubMed] [Google Scholar] - 15.Kano M, Sosulski AE, Zhang L, Saatcioglu HD, Wang D, Nagykery N, et al. AMH/MIS as a contraceptive that protects the ovarian reserve during chemotherapy. Proc Natl Acad Sci USA. 2017;114:E1688–97. [DOI] [PMC free article] [PubMed] [Google Scholar] - 16.Bell-Hensley A, Das S, McAlinden A. The miR-181 family: wide-ranging pathophysiological effects on cell fate and function. J Cell Physiol. 2023;238(4):698–713. [DOI] [PMC free article] [PubMed] [Google Scholar] - 17.Lv S, Xu J, Chen L, Wu H, Feng W, Zheng Y, Li P, Zhang H, Zhang L, Chi G, Li Y. MicroRNA-27b targets CBFB to inhibit differentiation of human bone marrow mesenchymal stem cells into hypertrophic chondrocytes. Stem Cell Res Ther. 2020;11(1):392. [DOI] [PMC free article] [PubMed] [Google Scholar] - 18.Virtue AT, McCright SJ, Wright JM, Jimenez MT, Mowel WK, Kotzin JJ, Joannas L, Basavappa MG, Spencer SP, Clark ML, Eisennagel SH, Williams A, Levy M, Manne S, Henrickson SE, Wherry EJ, Thaiss CA, Elinav E, Henao-Mejia J. The gut microbiota regulates white adipose tissue inflammation and obesity via a family of microRNAs. Sci Transl Med. 2019;11(496):eaav1892. [DOI] [PMC free article] [PubMed] [Google Scholar] - 19.Su Y, Yuan J, Zhang F, Lei Q, Zhang T, Li K, Guo J, Hong Y, Bu G, Lv X, Liang S, Ou J, Zhou J, Luo B, Shang J. MicroRNA-181a-5p and microRNA-181a-3p cooperatively restrict vascular inflammation and atherosclerosis. Cell Death Dis. 2019;10(5):365. [DOI] [PMC free article] [PubMed] [Google Scholar] - 20.Das S, Kohr M, Dunkerly-Eyring B, Lee DI, Bedja D, Kent OA, Leung AK, Henao-Mejia J, Flavell RA, Steenbergen C. (a). Divergent effects of miR-181 family members on myocardial function through protective cytosolic and detrimental mitochondrial microRNA Targets. J Am Heart Assoc 2017; 6(3). [DOI] [PMC free article] [PubMed] - 21.Su R, Lin HS, Zhang XH, Yin XL, Ning HM, Liu B, Zhai PF, Gong JN, Shen C, Song L, Chen J, Wang F, Zhao HL, Ma YN, Yu J, Zhang JW. MiR-181 family: regulators of myeloid differentiation and acute myeloid leukemia as well as potential therapeutic targets. Oncogene. 2015;34(25):3226–39. [DOI] [PubMed] [Google Scholar] - 22.Williams A, Henao-Mejia J, Harman CC, Flavell RA. miR-181 and metabolic regulation in the immune system. Cold Spring Harb Symp Quant Biol. 2013;78:223–30. [DOI] [PubMed] [Google Scholar] - 23.Otsuka F, McTavish K, Shimasaki S. Integral role of GDF-9 and BMP-15 in ovarian function. Mol Reprod Dev. 2011;78(1):9–21. [DOI] [PMC free article] [PubMed] [Google Scholar] - 24.Sanfins A, Rodrigues P, Albertini DF. GDF-9 and BMP-15 direct the follicle symphony. J Assist Reprod Genet. 2018;35(10):1741–50. [DOI] [PMC free article] [PubMed] [Google Scholar] - 25.NIH National Library of Medicine. Smad9 SMAD family member 9. Gene ID: 55994, updated on 9-May-2024. https://www.ncbi.nlm.nih.gov/gene/55994#:~:text=findings%20suggest%20that%20Smad9%20is,Wnt%2Fbeta%2Dcatenin%20pathways. Accessed 11 May 11 2024. - 26.Xu J, Bishop CV, Lawson MS, Park BS, Xu F. Anti-Mullerian hormone promotes pre-antral follicle growth, but inhibits antral follicle maturation and dominant follicle selection in primates. Hum Reprod. 2016;31:1522–30. [DOI] [PMC free article] [PubMed] [Google Scholar] - 27.Kushnir VA, Seifer DB, Barad DH, Sen A, Gleicher N. Potential therapeutic applications of human anti-Müllerian hormone (AMH) analogues in reproductive medicine. J Assist Reprod Genet. 2017;34(9):1105–13. 10.1007/s10815-017-0977-4. [DOI] [PMC free article] [PubMed] [Google Scholar] - 28.Detti L. Website Newsletter. International Society for Fertility Preservation. Anti-Müllerian hormone (AMH) and its multiple purposes in fertility preservation. January 2019; ISFP website - 29.Rodgers RJ, Abbott JA, Walters KA, Ledger WL. Translational physiology of Anti-Müllerian hormone: clinical applications in female fertility preservation and cancer treatment. Front Endocrinol. 2021;12:689532. 10.3389/fendo.2021.689532. eCollection 2021. [DOI] [PMC free article] [PubMed] Associated Data This section collects any data citations, data availability statements, or supplementary materials included in this article. Data Availability Statement The data described in this manuscript is kept in a private database and can be made available to the reviewers.