Role of cellular prion protein in mouse granulosa cells and its effects on ovarian function in knockout mice

Introduction

The cellular prion protein (PrPc) has been studied for its diverse roles across various biological systems, such as providing neuroprotection through the regulation of copper and NMDA receptors, as well as maintaining stem cell health by influencing Wnt/β-catenin signaling pathways (1–6), with particular emphasis on unraveling its mechanisms of action within the nervous system (7,8). Notably, PrPc has been revealed to interact with amyloid-β peptides in Alzheimer's disease and α-synuclein in Parkinson's disease (9,10). Furthermore, PrPc can modulate neuroendocrine signaling pathways, especially pathways that are associated with the regulation of gonadotropin, revealing a possible role in reproductive physiology (11,12). PrPc is involved in oocyte maturation (13). Furthermore, in ovariectomized ewes, the expression of PrPc in ovine uteroplacental tissues was shown to increase when the ewes were treated with estrogen and during the early stage of pregnancy (14). However, the roles of PrPc, including how it regulates ovarian reserve maintenance and the dynamics of follicular recruitment, have not been fully elucidated.

Recent advances in reproductive endocrinology demonstrate the need to clarify the non-steroidogenic mechanisms that govern follicular homeostasis, with a particular focus on mechanisms involving anti-Müllerian hormone (AMH)-mediated signaling (15–17). As AMH levels are closely linked to ovarian reserve and are recognized as the most sensitive clinical biomarker for diminished ovarian reserve (DOR) syndromes, it is crucial to clarify the regulatory mechanisms involved (18,19). We hypothesized that PrPc may act as a novel type of modulator for AMH-dependent follicular recruitment, potentially through mechanisms that are different from the regulation of classical steroid hormones. To test this hypothesis, two complementary models were used, the in vitro mouse ovarian granulosa cells (mGCs) with prion protein gene (PRNP) knockdown and overexpression (OE) and the in vivo PRNP knockout (KO) and wild-type (WT) models. The specific effects of PrPc on AMH secretion were compared with those on progesterone (P4) and estradiol (E2) production. The present study aimed to analyzed the effects of maintaining the follicular pool through histomorphometric analysis and reproductive longevity via litter size tracking, while also examining neuroendocrine adaptations in the HPG axis using vaginal cytology, serum FSH quantification, and ovarian reserve evaluation through follicle counting.

Materials and methods

Cell culture

mGCs were procured from Haixing Biosciences Co., Ltd. (cat. no. ORCM050). The cells were grown in high-glucose DMEM (cat. no. 12,100; Beijing Solarbio Science & Technology Co., Ltd.), which included 100 U/ml penicillin and 100 µg/ml streptomycin as standard components, with 10% heat-inactivated FBS (cat. no. FSS500; Shanghai ExCell Biology, Inc.). mGCs were maintained at 37°C in an atmosphere with 5% CO2, and the culture medium was replaced every 2 days. To preserve physiological relevance, cell passage was restricted to early generations (P2-P3), thereby minimizing cellular dedifferentiation. Additionally, follicle stimulating hormone receptor (FSHR) expression levels were periodically confirmed by immunofluorescence, ≥80% of cells exhibited positive FSHR staining (Fig. S1A).

Immunofluorescence

Cell coverslips were prepared by seeding 1×105 cells in 1 ml of medium and incubating them overnight at 37°C with 5% CO2 until they reached 70–80% confluency. After removing the medium, the cells were washed with ice-cold PBS and fixed using 4% paraformaldehyde (PFA) for 30 min at room temperature. Following additional washes with PBS, the cells were permeabilized with 0.5% Triton™ X-100 for 15 min and then blocked with 10% goat serum (cat. no. S9070; Beijing Solarbio Science & Technology Co., Ltd.) for 60 min at room temperature. The coverslips were then incubated overnight at 4°C with a rabbit anti-FSHR antibody diluted to 1:200 (cat. no. 22665-1-AP; Proteintech Group, Inc.). After washing, the cells were stained with an Alexa Fluor® 488-conjugated secondary antibody at a dilution of 1:500 (cat. no. SA00006-2; Proteintech Group, Inc.) for 2 h at room temperature. Finally, the coverslips were mounted on slides using DAPI antifade medium (cat. no. S2110; Beijing Solarbio Science & Technology Co., Ltd.), and the samples were imaged for analysis using an inverted platform (Primovert LED; ZEISS, Germany).

Transfection

Recombinant lentiviruses, rLV-short hairpin (sh)RNA-Puro-mPRNP and rLV-mPRNP−3flag-ZsGreen-Puro were procured from Haixing Biosciences Co., Ltd. Empty vectors were used as negative controls (NCs). The target sequences of the PRNP shRNAs were as follows: mPRNP sh-1, 5′-GGACAACCTCATGGTGGTAGT-3′; mPRNP sh-2, 5′-GCGTCAATATCACCATCAAGC-3′; and mPRNP sh-3, 5′-GCCTATTACGACGGGAGAAGA-3′. To achieve stable silencing and OE of the PRNP gene, the cells were seeded into 6-well plates at a density of 2.5×105 cells per well, and cultured overnight until they reached a confluence of 40–50%. On the subsequent day, lentiviral transfection of the cells was performed following the protocol supplied by the manufacturer. Lentiviral vectors that encode EGFP reporters were generated using a third-generation system in 293T cells (cat. no. CL-0005; Wuhan Pricella Biotechnology Co., Ltd.). The cells were transfected at 70% confluence with 3 µg pLVX-shRNA2-Puro-mPrnp or pLVX-mPrnp-3flag-ZsGreen-Puro transfer plasmid, 2 µg of psPAX2, and 1 µg of pMD2.G in a ratio of 3:2:1 using PEI in Opti-MEM. After 48 h of incubation at 37°C with 5% CO2, fluorescence screening indicated a packaging efficiency of over 85%. To obtain stably transfected mGCs, lentiviral transduction was performed at the optimized multiplicity of infection (MOI) of 80, followed by selection with 3 µg/ml of puromycin for 72 h (cat. no. PS1224; Beijing Puxitang Biotechnology Co., Ltd.), and 1 µg/ml thereafter for culture maintenance. After 72 h of transduction, the infected mGCs were collected for subsequent experiments. Stable PrPc-knockdown mGCs were established through lentiviral shRNA transduction. Following this, these cells were infected with a lentivirus designed to overexpress PrPc, allowing for both the knockdown and overexpression of PrPc within the same cellular environment.

Reverse transcription quantitative PCR (RT-qPCR)

Total RNA was extracted from mGCs following the manufacturer's instructions (cat. no. UE-MN-MS-RNA-10; Suzhou UElandy Biotechnology Co., Ltd.). Quantity and purity of RNA were determined using a NanoDrop™ 2000 spectrophotometer (Thermo Scientific). The RNA concentrations were measured at A260, while the purity was evaluated by calculating the A260/A280 ratios. Subsequently, RT was performed following the manufacturer's instructions for the RevertAid™ Master Mix (cat. no. M1632; Thermo Fisher Scientific, Inc.). qPCR was carried out on a CFX Connect fluorescent Real-Time PCR System (Bio-Rad Laboratories, Inc.), using the Universal SYBR Green qPCR Supermix (cat. no. S2024L; US Everbright, Inc.). The thermocycling conditions were as follows: 95°C for 5 min, followed by 45 cycles of 95°C for 5 sec and 60°C for 30 sec. The PRNP primer sequences (cat. no. MQP094390; GeneCopoeia, Inc.) were as follows: PRNP forward (F), 5′-GGCCCATGATCCATTTTGGC-3′ and reverse (R), 5′-TGCTGTACTGATCCACTGGC-3′. Additionally, the following primer sequences were used for β-actin (Wuhan Servicebio Technology Co., Ltd.): β-actin F, 5′-GACTTTGTACATTGTTTTG-3′ and R, 5′-TGCACTTTTATTGGTCTCA-3′, which was selected as the internal control for normalization. The 2−ΔΔCq method was employed for quantification of the data (20). qPCR was selected for its superior sensitivity in detecting transcript-level changes (detection threshold ≤10 copies/µl) compared with conventional PCR (21,22), with SYBR Green chemistry providing cost-effective quantification of PRNP expression dynamics.

Western blotting (WB)

mGCs and ovarian tissue were lysed on ice for 30 min in RIPA buffer supplemented with protease inhibitors (cat. no. R0010; Beijing Solarbio Science & Technology Co., Ltd.). The total protein content was determined using a BCA Protein Quantification Kit (cat. no. C0050; TargetMol Chemicals Inc.). Following protein quantification, 25 µg total protein/lane in each sample were separated via 10% SWE Rapid High Resolution Running Buffer (cat. no. G2081-1L; Wuhan Servicebio Technology Co., Ltd.). Subsequently, the separated proteins were transferred onto PVDF membranes (cat. no. IPVH00010; MilliporeSigma).

To block non-specific binding, the PVDF membranes were incubated in a solution of 5% non-fat dried milk (cat. no. S10191; Beijing Puxitang Biotechnology Co., Ltd.) in Tris-buffered saline containing 0.1% Tween 20 (TBST; cat. no. T1082; Beijing Solarbio Science & Technology Co., Ltd.) for 2 h at room temperature. After blocking, the membranes were incubated overnight at 4°C with the following primary antibodies: Anti-CD230 (Prion; cat. no. 808001; BioLegend, Inc.) and anti-β-actin at a 1:5,000 dilution (cat. no 20536-1-AP; Proteintech Group, Inc.).

After the overnight incubation, the membranes were washed three times with TBST and incubated for 2 h at room temperature with the following secondary antibodies: HRP-conjugated goat anti-rabbit IgG at a 1:1,000 dilution (SA00001-2; Proteintech Group, Inc.) and HRP-conjugated goat anti-mouse IgG at a 1:10,000 dilution (SA00001-1; Proteintech Group, Inc.).

Finally, an enhanced chemiluminescence detection reagent (cat. no. S6009L; Suzhou UE Landi Biotechnology Co., Ltd.) was used to visualize the immunoreactive bands on ChemiDoc MP (Bio-Rad Laboratories, Inc.). Densitometry of the WB bands was performed using ImageJ 1.48V software (National Institutes of Health).

Apoptosis assay

Following the instructions of the Annexin V/PI detection Kit (cat. no. Y6026S; Suzhou UE Landy Biotechnology Co., Ltd.), 1×105 mGCs were digested with 0.25% trypsinization solution (without EDTA)(cat. no. T1350; Beijing Solarbio Science & Technology Co., Ltd.), then enzymatically neutralized and resuspended in cold staining buffer, then resuspended in a staining buffer, which was prepared by adding 5 µl each Annexin V storage solution and 5 µl PI storage solution. Subsequently, the cells were incubated in the dark at 4°C for 20 min and analyzed using a NovoCyte D3000 flow cytometer (Agilent Technologies, Inc.) Flow cytometry data were analyzed using De Novo FCS Express™ 6 software (Agilent Technologies, Inc.). Annexin V/PI dual staining was employed to differentiate early apoptotic (Annexin V+/PI−) from late apoptotic/necrotic (Annexin V+/PI+) populations, enabling stage-specific analysis of mGC death pathways, the total percentage of apoptotic cells reflects the overall measurement of both early and late apoptotic populations.

Cell cycle assay

For cell cycle analysis, 5×105 mGCs were harvested by trypsinization at 72 h post-infection. The cells were then washed with cold PBS and fixed in 70% ice-cold ethanol at 4°C overnight. Next, mGCs were incubated with RNase A and PI in a 0.5 ml reaction mixture (cat. no. C6031S; Suzhou UE Landy Biotechnology Co., Ltd.) at 37°C for 30 min in a dark chamber. Cell cycle distribution was measured using a NovoCyte D3000 flow cytometer and analyzed with De Novo FCS Express™ 6 software (Agilent Technologies, Inc.).

ELISA for hormone measurements

mGC culture medium containing 10% FBS and mouse serum collected by centrifugation at 3,000 × g for 5 min at 4°C were used to measure the concentrations of E2 (cat. no. SEKM-0286; Beijing Solarbio Science & Technology Co., Ltd.), AMH (cat. no SEKM-0310; Beijing Solarbio Science & Technology Co., Ltd.), P4 (cat. no. SEKSM-0002; Beijing Solarbio Science & Technology Co., Ltd.), FSH (cat. no. KE1425; ImmunoWay Biotechnology Company) and luteinizing hormone (LH; cat. no. KE1421; ImmunoWay Biotechnology Company), following the manufacturer's instructions.

Animals

A total of 24 mice were used, comprising FVB-PRNP-/- and FVB wild-type (WT) mice, with each group consisting of 12 females. At the start of the experiment, the mice were 7 weeks old, and body weight was of 19.1–22.9 g (mean ± SD: 20.82±1.13 g). PRNP KO mice with an FVB genetic background were provided by Professor Wen-Quan Zou (Jiangxi Academy of Clinical Medical Sciences of the First Affiliated Hospital of Nanchang University) and Professor Li Cui (Department of Neurology, First Hospital of Jilin University). FVB WT mice, sourced from GemPharmatech Co. Ltd., were utilized as controls in the present study. All experimental protocols were approved by the Institutional Animal Care and Use Committee (IACUC) of the First Affiliated Hospital of Nanchang University (approval no. CDYFY-IACUC-202310QR030; Nanchang, China) in accordance with the National Institutes of Health Guidelines for animal welfare. To maintain optimal health and experimental integrity, both the KO and WT mice were housed in individually ventilated cages under specific pathogen-free (SPF) conditions, maintaining a temperature of 22±1°C and humidity levels of 55±10%. They were kept on a 12:12-h light/dark cycle. The animals had unrestricted access to autoclaved standard rodent chow and reverse-osmosis purified water. These carefully controlled housing conditions minimized the potential influence of external pathogens on the experimental outcomes, ensuring that any observed differences between the KO and WT groups could be more accurately attributed to the genetic modifications rather than environmental factors. Breeding pairs were formed by housing WT and KO female mice (n=6 per group) with proven-fertility WT males at 10 weeks of age under specific pathogen-free conditions. After confirming the presence of copulatory plugs, which indicated successful mating and was designated as Gestational Day 0, the dams were individually housed in ventilated cages. Inhalant anesthesia was induced with 5% isoflurane (cat. no. R510-22-16; RWD Life Science Co., Ltd.) in oxygen at 1 l/min flow rate, and maintained for 5 min until the pedal withdrawal reflex ceased and the respiratory rate stabilized (40–60 breaths/min). Cervical dislocation was performed with simultaneous isoflurane overdose to ensure humane euthanasia. Death validation included confirmation of apnea for >3 min and absence of the corneal reflex. Terminal whole blood samples (~0.3 ml) were obtained via retro-orbital venous plexus puncture within 2 min post-mortem to ensure coagulation integrity. Ovarian tissues were dissected, fixed in 4% paraformaldehyde at 4°C for 24–48 h, then processed for paraffin embedding and hematoxylin & eosin (H&E) staining.

H&E staining

H&E staining was performed in accordance with a standard protocol (23,24). Specifically, ovarian tissues were fixed in 4% paraformaldehyde (w/v in 0.1 M PBS, pH 7.4) at 4°C for a duration of 24 to 48 h, 5-µm-thick paraffin sections underwent three 5-min immersions in xylene. This was followed by a rehydration process through a graded series of ethanol concentrations: 100, 95, 80, and 70% (v/v), with each step lasting 2 min, ultimately concluding in distilled water. Next, the sections were stained with H&E (cat. no. L11021604l; Nanchang Yulu Experimental Equipment Co., Ltd.). Sections were stained with Mayer's hematoxylin at room temperature (23±2°C) for a duration of 3 min, and this was followed by staining with eosin Y alcoholic solution at the same temperature for 2 min. Ovarian follicles were quantified using systematic random sampling in every sixth serial section with a thickness of 5 µm. A manual counting method was employed, examining 30 non-overlapping fields per ovary at a magnification of 200× using a light microscope (Beijing Sunny Instruments Co., Ltd.). Follicles were identified across various stages, from primordial to antral, in accordance with the Amsterdam Consensus Criteria (25,26).

Estrous cycle staging

Estrous cycle staging was performed through daily vaginal smear cytology between 9:00-10:00 a.m. Vaginal lavage samples were obtained using 10 µl sterile PBS, placed onto poly-L-lysine coated slides, and fixed in 4% paraformaldehyde at 4°C for 15 min. Hematoxylin (3 min) and eosin (30 sec) staining at room temperature, adhering to standard protocols.

Statistical analysis

Data are presented as the mean ± SD based on a minimum of three independent experiments. Prior to analysis, normality was assessed using Shapiro-Wilk test (α=0.05) and the homogeneity of variance was confirmed via Brown-Forsythe test. Parametric tests were applied when assumptions were satisfied. Statistical analyses were carried out using GraphPad Prism 9.5 software (Dotmatics). For the comparison between two groups, a two-tailed unpaired Student's t-test was used, with Cohen's d effect sizes and 95% confidence intervals automatically calculated by the software. For the comparison of >2 groups, one-way ANOVA with Tukey's post hoc test (for all pairwise comparisons) or Dunnett's post hoc test (for comparisons vs. the control group) was applied, with partial η2 effect sizes and family-wise error rate control at α=0.05. P<0.05 was considered to indicate a statistically significant difference.

Results

Validation of PrPc knockdown and OE efficiency

To knock down PrPc in mGCs, the following lentivirus-based vectors were used: mPRNP sh-1, sh-2 and sh-3. Compared with those of mGCs infected with the NC vector (sh-NC), the mRNA expression levels of PRNP in the sh-1 group exhibited no significant change (P>0.05). However, infections with sh-2 and sh-3 resulted in a marked reduction in mRNA expression (sh-2:P<0.05; Sh-3:P<0.001), respectively, confirming successful knockdown of the PRNP gene (Fig. 1A). Supporting these findings, WB analysis revealed that the gray values of the PrPc protein bands in cells transfected with sh-2 and sh-3 were significantly reduced compared with those in the sh-NC group (P<0.05; Fig. 1B). Based on these results, sh-2 and sh-3 were selected as the most efficient shRNAs for subsequent experiments.

Figure 1. PrPc knockdown and OE efficiency in mGCs. (A) mRNA expression levels of PRNP and (B) protein expression levels of PrPc in mGCs transfected with different shRNAs. (C) mRNA expression levels of PRNP and (D) protein expression levels of PrPc in mGCs transfected with OE vectors. β-actin was used as an internal control. *P<0.05; **P<0.01; ***P<0.001. ns, non-significant; PrPc, cellular prion protein; PRNP, prion protein gene; mGCs, mouse ovarian granulosa cells; OE, overexpression; sh, short hairpin; NC, negative control.

In the PrPc OE group, the mRNA expression levels of PRNP were significantly increased compared with those in the OE-NC group (P<0.001; Fig. 1C), indicating that the OE vector effectively increased the transcription of the PRNP gene. Correspondingly, the expression levels of the PrPc protein in the OE group was significantly increased compared with that in the OE-NC group (P<0.01; Fig. 1D), aligning with the observed trend in mRNA expression levels.

PrPc does not alter mGC cycle progression or apoptosis

Flow cytometry analysis was carried out to assess cell cycle distribution in mGCs. The results revealed no significant differences in the proportions of cells in the G1, S and G2 phases among the sh-2, sh-3 and sh-NC groups (P>0.05; Fig. 2A). Likewise, no notable changes were observed in the distribution of cells across these phases in the OE group compared with OE-NC (P>0.05; Fig. 2B).

Figure 2. Cellular prion protein exhibits minimal influence on the cell cycle distribution and apoptosis of mouse granulosa cells. Flow cytometry was employed to analyze the cell cycle distribution, enabling quantification of cells in the G0/G1, S and G2/M phases after transfection with (A) different shRNAs or (B) OE vectors. Apoptotic and necrotic cells were assessed using Annexin V/PI staining after transfection with (C) different shRNAs or (D) OE vectors. ns, non-significant; sh, short hairpin; NC, negative control; OE, overexpression.

Analysis of apoptosis using the Annexin V/PI double staining method by flow cytometry revealed that, under normal culture conditions, the apoptosis rate of mGCs was consistent between the sh-NC and OE-NC groups. Moreover, sh-2, sh-3 and sh-NC did not exhibit any significant differences in the proportion of apoptotic cells (P<0.05; Fig. 2C). Similarly, the OE group demonstrated no notable variation in the proportion of apoptotic cells compared with OE-NC (P<0.05; Fig. 2D).

PrPc knockdown reduces AMH secretion while preserving P4 and E2 production

AMH levels averaged 56.10±3.73 ng/ml in the sh-NC group, while they significantly decreased to 42.27±7.51 and 42.7±6.85 ng/ml in the sh-2 and sh-3 groups respectively (P<0.05; Fig. 3A). However, PrPc expression was restored following knockdown in mGCs infected both with the PRNP knockdown and OE vectors (sh-2 + OE and sh-3 + OE; Fig. S1B), and AMH levels increased compared with those in the sh-2 and sh-3 groups (P<0.05; Fig. 3A). By contrast, the average AMH expression levels in the OE group was 46.96±4.22 ng/ml, exhibiting no significant difference compared with that in the OE-NC group (P>0.05; Fig. 3A).

Figure 3. Cellular prion protein knockdown specifically reduces AMH secretion in mGCs. (A) AMH, (B) P4 and (C) E2 levels in the culture medium of mGCs transfected with different shRNAs, OE vectors or their combination. *P<0.05. ns, non-significant; AMH, anti-Müllerian hormone; mGCs, mouse granulosa cells; P4, progesterone; E2, estradiol; sh, short hairpin; OE, overexpression; NC, negative control.

The average P4 level was 38.61±0.15 ng/ml in the sh-NC group, and 38.57±0.43 and 38.24±0.37 ng/ml in the sh-2 and sh-3 groups, respectively, with the latter showing no significant difference compared with the control group (P>0.05; Fig. 3B). Furthermore, compared with the sh-2 and sh-3 groups, the sh-2 + OE and sh-3 + OE groups did not exhibit significant changes in P4 levels (P>0.05; Fig. 3B). Similarly, the average P4 level in the OE group was 38.33±0.40 ng/ml, which was comparable with that in the OE-NC group (P>0.05; Fig. 3B).

The average E2 level in the sh-NC group was 67.33±10.54 pg/ml, while the sh-2 and sh-3 groups exhibited values of 68.11±10.07 and 68.74±5.81 pg/ml, respectively, both showing no significant differences compared with sh-NC (P>0.05; Fig. 3C). Similarly, compared with the sh-2 and sh-3 groups, no notable changes in E2 expression levels were observed in the sh-2 + OE and sh-3 + OE groups (P>0.05; Fig. 3C). Likewise, the average E2 level in the OE group was 65.67±11.67 pg/ml, which was not significantly different compared with that in the OE-NC group (P>0.05; Fig. 3C).

PrPc expression levels do not affect the classification count of ovaries

Ovaries from WT and KO mice were isolated and WB was carried out to assess PrPc protein expression levels. The results indicated that PrPc levels in the ovaries of KO mice were reduced compared with those in the WT group (Fig. 4A) confirming the effectiveness of the KO.

Figure 4. Prion protein gene KO mice exhibit normal ovarian morphology and follicular development. (A) PrPc protein expression in the ovaries of WT and KO mice. (B) Ratio of bilateral ovary weight to body weight showed no differences between groups. (C) H&E staining on mouse ovaries (scale bars, 200 µm for overviews and 100 µm for higher mag). The stages of follicular development are clearly indicated by color-coded arrows: Blue represents primordial follicles, which consist of a single layer of squamous granulosa cells; green denotes primary follicles, characterized by cuboidal granulosa cells; white signifies secondary follicles, identifiable by having ≥3 layers of granulosa cells and the early formation of an antrum; black marks antral follicles, distinguished by the presence of a fluid-filled cavity; and yellow indicates atretic follicles, which display pyknotic granulosa cell nuclei and fragmentation of the zona pellucida. (D) Follicle counts across developmental stages were comparable between WT and KO mice. ns, non-significant; WT, wild-type; KO, knockout; PrPc, cellular prion protein.

At 7 weeks of age, ovarian tissue was collected, and the wet weight index of ovaries was measured. The WT group had an ovarian weight of 0.390±0.08 mg/g, while the KO group exhibited a weight of 0.389±0.02 mg/g, with no statistically significant difference between the groups (P>0.05; Fig. 4B).

H&E staining revealed the pathological and morphological characteristics of the ovaries in each group (Fig. 4C). Although the KO group exhibited a slight reduction in the number of primordial and primary follicles, this difference was not statistically significant (P>0.05; Fig. 4D). Similarly, there were no significant differences in the numbers of secondary, antral or atretic follicles between the two groups (P>0.05; Fig. 4D).

PRNP deficiency elevates circulating FSH levels with unaltered AMH, LH and E2 levels

FSH levels in KO mice were significantly increased at 6.64±0.68 ng/ml compared with 4.52±1.18 ng/ml in the WT group (P<0.01; Fig. 5A). By contrast, E2 levels were not significantly different between the two groups (P>0.05; Fig. 5B). Similarly, AMH and LH levels were also not significantly different between WT and KO mice (P>0.05; Fig. 5C and D).

Figure 5. KO mice exhibit elevated FSH levels but normal E2, AMH and LH levels. Serum hormone levels of (A) FSH, (B) E2, (C) AMH and (D) LH in WT and KO mice. **P<0.01. ns, non-significant; FSH, follicle-stimulating hormone; LH, luteinizing hormone; E2, estradiol; AMH, anti-Müllerian hormone; WT, wild-type; KO, knockout.

PRNP KO mice exhibit normal reproductive performance

WT and KO mice exhibited a standard estrous cycle, typically spanning 4–5 days per cycle and encompassing four distinct phases: Proestrus, estrus, metestrus and diestrus (Fig. 6A). In the proestrus stage (P), the smears are primarily made up of nucleated epithelial cells, which are characterized by their polygonal shape, large spherical nuclei, and abundant cytoplasm. As the cycle progresses to estrus (E), there is a notable increase in keratinized epithelial cells, which display a distinct eosinophilic staining in their cytoplasm. This is followed by metestrus (M), where the number of keratinized cells begins to decline, and nucleated epithelial cells become more present, accompanied by a few clusters of leukocytes. Finally, in the diestrus stage, the cytological composition shifts to predominantly leukocytes, with few scattered nucleated epithelial cells present (27). Each stage in the WT mice was clearly defined, with smooth and consistent transitions between phases. Similarly, KO mice exhibited no notable alterations in cycle duration, with neither a marked prolongation nor a noticeable shortening of the cycle observed (Fig. 6B).

Figure 6. KO mice exhibit regular estrous cycle and fertility. (A) Vaginal secretion smears stained with H&E stain during estrous cycle phases: Proestrus, estrus, metestrus, diestrus (scale bar, 50 µm). (B) Regular estrous cycle patterns in mice spanning 9 consecutive days. (C) Interval from mating, following co-housing, to parturition. (D) Number of offspring at birth. ns, non-significant; WT, wild-type; KO, knockout; P, proestrus; E, estrus; M, metestrus; D, diestrus.

No statistically significant differences were observed in the litter intervals between the two groups (P>0.05; Fig. 6C). The average litter size remained consistent at 8–10 pups, and the mothers exhibited normal nurturing abilities. The offspring survival rate was high, with no notable differences detected between the groups (P>0.05; Fig. 6D).

Discussion

Previous studies have predominantly explored the role of PrPc within the nervous system (28–31). However, there is little research on the function of PrPc in non-neuronal tissues, especially in endocrine regulation. The present in vitro study revealed that PrPc can selectively affect the secretion of AMH, while no significant changes in AMH, P4, E2, folliculogenesis and the reproductive cyclicity in mice. These findings broaden the functional range of PrPc beyond its typical roles in neuroprotection and synaptic plasticity (32,33), indicating that PrPc may be a potential niche regulator of ovarian reserve biomarkers.

The present study revealed that alterations in the expression of PrPc in mGCs did not significantly influence the cell cycle or apoptosis, suggesting that PrPc may not regulate cellular homeostasis in mGCs. PrPc may not directly affect the signaling molecules involved in key cell cycle transitions, such as those between the G1/S and G2/M phases. However, its role varies significantly in other systems. In neural environments, PrPc acts as an accelerator of apoptosis during proteotoxic stress by impairing the ESCRT-0/AMPAR axis. In renal settings, it promotes regeneration through the activation of the PI3K/Akt-mTOR pathway. In cardiac contexts, PrPc is crucial for structural recovery, although it does not influence functional recovery. Additionally, in oncogenic environments, it serves as a key regulator of cell fate by modulating the dynamics between p53 and MDM2 during endoplasmic reticulum stress (34–36). Additionally, a previous study has highlighted the presence of compensatory mechanisms among members of the prion protein family, particularly PrPc and Shadoo, which aid in maintaining cell cycle progression and suppressing apoptosis (37).

The levels of AMH, which is considered a marker of the ovarian reserve function and is primarily secreted by GCs (15), were decreased when PrPc was knocked down. The effect of PrPc knockdown may be associated with disrupted TGF-β/bone morphogenetic protein (BMP)-SMAD superfamily signaling pathways (38). AMH transcription is strictly controlled by SMAD1/5/8 complexes, which are activated by BMP ligands (39). The present findings are consistent with mechanistic framework proposed by Puig et al (40), which suggests that PrPc plays a role in modulating the trafficking of FSHR and bone morphogenetic protein receptors (BMPR-II). Regarding the secretion of P4 and E2, no significant changes were observed after PrPc knockdown or overexpression, and the AMH-specific effect contrasts those of classical steroidogenic regulators, such as PPAR-γ agonists, miR-335-5p, retinoic acid, and artemisinins, which alter multiple hormonal axes (41–44). It may be hypothesized that PrPc stabilizes BMP receptor clusters on GCs, which activates SMAD phosphorylation and subsequent AMH synthesis. This hypothesis can be tested by mapping the interactions between PrPc and BMP receptor 2 and by quantifying the levels of phosphorylated-SMAD1/5 in PrPc knockdown models.

The results of the present study indicating that PrPc is dispensable for murine ovarian function contrasts with its reported roles in neuronal survival, underscoring profound tissue specificity (45). Although PrPc deletion in neurons induces apoptosis via Bcl-2 suppression, GCs may remain viable after PrPc depletion through compensatory mechanisms involving the Shadoo protein (46).

Despite minor changes, no statistically significant differences were observed in the ovary weight and the numbers of primordial and primary follicles in the PrPc KO mice compared with the WT group. Furthermore, there seemed to be no evident changes in the estrous cycle, litter interval or the number of pups per litter between the two groups. This suggests that PrPc may not be critical in maintaining the structure and reproductive activity of the ovaries. Vigorous compensatory mechanisms in the organism may have abolished the effect of PrPc deficiency on ovarian function.

Furthermore, the lack of PrPc does not seem to interfere with the estrous cycle or the reproductive functions, potentially because the hormonal regulatory pathways within the hypothalamic-pituitary-ovarian axis maintain normal reproductive rhythms and functions (47). Nonetheless, it cannot be ruled out that, under chemotherapy-induced ovarian damage or under hyperandrogenic conditions mimicking polycystic ovary syndrome (PCOS), PrPc may have a key impact on reproductive functions.

The mouse model used in the present study may not capture all the physiological and pathological intricacies of the human ovary. There are marked differences in the structural, functional and hormonal regulation, as well as the reproductive cycle between mouse and human ovaries. For example, folliculogenesis in humans is a process that spans 90 to 120 days, which is significantly longer than the 14 to 21-day cycle observed in mice. In humans, primordial follicles can remain dormant for several decades, a situation that can contribute to oxidative stress within the ovarian environment. Additionally, humans typically exhibit a single-wave, monovulatory pattern for the selection of the dominant follicle, in contrast to mice, which display multi-wave, polyovulatory cycles (48). In addition, there is an apparent limitation to the in vitro cell culture model used in the present study, since cultured cells may lose some of their physiological characteristics (49,50) and be affected by microenvironmental conditions (51,52), which may result in findings that do not represent the in vivo settings. Murine ovaries lack the protracted folliculogenesis and hormonal complexity of humans. Furthermore, environmental endocrine disruptors such as phthalates (53), which potently alter human AMH levels, were not examined in the present study.

PRNP KO mice seem to have phenotypic resilience, but this does not entirely invalidate their clinical relevance. Under stress, there can be a compensatory failure and PrPc may have a key effect on ovarian function. In mice, Shadoo-mediated compensatory mechanisms may take place (54), whilst in humans these mechanisms may not occur. When humans face prolonged oxidative stress, for example during chemotherapy or when there is age-related follicular depletion, Shadoo-mediated redundancy may not be sufficient. The observation of elevated (FSH) levels in PRNP knockout (KO) mice aligns with the clinical characteristic of increased early-follicular phase FSH levels seen in patients diagnosed with diminished ovarian reserve (DOR), as defined by established international diagnostic criteria (55). Additionally, PrPc expression in human follicular fluid may be associated with AMH levels or in vitro fertilization outcomes; therefore, cohort studies examining the association between PrPc concentrations in human follicular fluid and key reproductive parameters is warranted are warranted.

The anti-apoptotic role of PrPc in glioma cells via interaction with PRKC Apoptosis WT1 Regulator (PAWR) raises the possibility that PrPc may protect GCs from chemotherapy-specific observations in glioma models (56). This hypothesis aligns with clinical observations that the ovarian reserve declines post-chemotherapy (57–59), it was hypothesized that PrPc expression critically regulates chemotherapy-induced ovarian reserve depletion by modulating granulosa cell survival pathways. Inhibiting PrPc-PAWR interactions may sensitize ovarian cancer cells to chemotherapy while preserving normal ovarian tissue. Conversely, enhancing PrPc activity could protect ovarian follicles during oncotherapy.

In conclusion, the present study revealed that PrPc selectively disrupted AMH secretion in mGCs without affecting the levels of P4 and E2 or the reproductive cycle. Unlike its anti-apoptotic roles in neurons or glioma, the role of PrPc in AMH regulation was independent of cell survival. It was hypothesized that: Murine ovaries may exhibit compensatory mechanisms via Shadoo in the absence of PrPc, while human cells exposed to prolonged stress, such as chemotherapy, may rely more on PrPc for the regulation of AMH levels and cell survival. Clinically, PrPc could refine ovarian reserve diagnostics or inspire fertility preservation strategies. Furthermore, targeting the PrPc-BMP-SMAD interactions may also address AMH dysregulation in polycystic ovary syndrome or ovarian insufficiency.

Supplementary Material

Supporting Data

Acknowledgements

The authors would like to thank Professor Wen-Quan Zou (Jiangxi Academy of Clinical Medical Sciences of the First Affiliated Hospital of Nanchang University, Nanching, China), for technical discussion. The authors would also like to thank Professor Li Cui (Department of Neurology, First Hospital of Jilin University, Changchun. China) for their donation of FVB-PRNP tm1/ILAS congenic mice.

Funding

The present study was partially supported by the National Natural Science Foundation of China (grant no. 82260295).

Availability of data and materials

The data generated in the present study may be requested from the corresponding author.

Authors' contributions

CT interpreted data and revised the manuscript. QC and HW, who made contributions to the study's conceptualization and data validation. In vivo investigations, including surgical procedures and H&E staining, were carried out by QC, JH, and YW. QC, FL, TD and XY performed experiments. QY contributed to pathological assessments and the drafting of the manuscript. QC and HW confirm the authenticity of all the raw data. All authors participated in data analysis, manuscript review and they collectively assume accountability for the published work. All authors read and approved the final manuscript.

Ethics approval and consent to participate

The experimental procedures in the present study were approved by the Institutional Animal Care and Use Committee of The First Affiliated Hospital of Nanchang University (approval no. CDYFY-IACUC-202310QR030; Nanchang, China).

Patient consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

References