. 2026 Oct 19;38(10):e70294. doi: 10.1111/jne.70294

Sustained induction of PMOS‐like phenotype in mice exposed to letrozole in drinking water

Kennedy A Stevenson ¹, Romy Kerbus ¹, Greg M Anderson ¹, Caroline Decourt ^1,^✉

PMCID: PMC13680047 PMID: 42896607

Abstract

Polyendocrine metabolic ovarian syndrome (PMOS; formerly polycystic ovary syndrome) is the most common endocrinopathy affecting women of reproductive age, characterized by ovarian dysfunction and hyperandrogenism, with metabolic disturbances including insulin resistance and obesity also present for 50% of PMOS patients. Effective preclinical models are crucial for evaluating therapeutic interventions. However, current mouse models of hyperandrogenemia induced by aromatase inhibition typically rely on subcutaneous letrozole implants, which require surgical procedures and struggle to sustain PMOS‐like phenotypes over an extended period necessary for thorough drug assessment. We hypothesized that an oral letrozole administration would provide extended maintenance of syndrome‐like features. Female mice received letrozole either via subcutaneous implant (~1.67 mg/kg/day for 70 days) or orally in drinking water (~1 mg/kg/day for up to 130 days). Estrous cyclicity was monitored by vaginal cytology. Hormonal, metabolic, and histological assessments were performed. While a subcutaneous letrozole implant only sustained hyperandrogenic, metabolic and reproductive phenotypes for ~50 days, oral letrozole administration sustained complete anovulation for at least 100 days, maintaining a persistent diestrus stage. The oral letrozole treatment produced comprehensive PMOS‐like features, including significantly elevated testosterone levels, increased luteinizing hormone concentration, pulse frequency and amplitude, enhanced hypothalamic androgen receptor expression, decreased hypothalamic progesterone receptor expression, cystic‐like ovarian morphology, increased body weight, impaired insulin sensitivity, and elevated adiposity during the entire time of the treatment period (18 weeks), compared to the placebo treatment. Uterine weight was not decreased, suggesting normal circulating estrogen concentrations. These findings demonstrate that oral letrozole delivery offers a more accessible method of administration, resulting in a sustained induction of PMOS‐like phenotypes. This approach effectively maintains both reproductive and metabolic features essential for comprehensive evaluation, thereby enhancing the model's utility for investigating therapeutic interventions in PMOS.

Keywords: drinking water, letrozole, mice, PCOS, PMOS

1. INTRODUCTION

Polyendocrine metabolic ovarian syndrome (PMOS), formely known as polycystic ovary syndrome (PCOS) is an intricate endocrinological disorder with complex interplays between reproductive, metabolic, and cosmetic manifestations. PMOS is the most common form of anovulatory infertility affecting women of reproductive age, with the global prevalence of 4%–20%. ¹ , ² According to the Rotterdam criteria, PMOS is diagnosed by the presence of two out of three key features ³ ; namely the presence of multiple cystic‐like ovarian morphology, irregular or absent menstrual cycles, and hyperandrogenism. Hyperandrogenism is a common hallmark of the condition, contributing to cosmetic manifestations of the syndrome such as hirsutism (excessive hair growth), acne, and alopecia. ⁴ Metabolic complications are also common in PMOS with approximately 50% of affected women exhibiting insulin resistance, ⁵ obesity, ⁶ and an elevated risk of cardiovascular diseases. ⁷ Current treatments are largely symptom‐based, combining oral contraceptives to regulate cycles, antihyperglycemic drugs to improve insulin sensitivity, and antiandrogens to reduce hyperandrogenism. ⁸ However, these therapies only target specific symptoms, require long‐term use, and do not address the underlying causes of the disorder, meaning there is currently no cure for PMOS.

Fertility is controlled by the hypothalamic–pituitary–gonadal (HPG) axis. When under normal physiological condition, the hypothalamus releases Gonadotropin‐releasing hormone (GnRH), influencing the anterior pituitary gland to then release luteinizing hormone (LH) and follicle‐stimulating hormone (FSH). High‐frequency GnRH pulses favor LH, while slower pulses favor FSH secretion. ⁹ The negative feedback loop between the ovaries and the hypothalamus and pituitary gland involves oestradiol and progesterone; however, in women with PMOS, there is an impaired sensitivity of the hypothalamus to negative feedback. ¹⁰ This results in the dysregulation of the HPG axis and leads to increased GnRH pulsatile secretion, triggering high‐frequency LH pulsatile release and increasing the ratio of LH:FSH. The peripheral effects of this include the increased production of ovarian androgen synthesis, resulting in a state of hyperandrogenism. ¹¹ , ¹² Our current understanding is that hyperandrogenism is not only a symptom of PMOS but also contributes to the onset of other PMOS manifestations. ¹³ , ¹⁴ Despite its high prevalence and multifaceted impact, the underlying causes of PMOS remain poorly understood. This lack of understanding underscores the need for robust and relevant animal models to unravel the mechanisms driving PMOS and to develop more effective therapeutic strategies.

Mouse models serve as a critical tool in investigating the effects of pharmacological interventions on the HPG axis, providing a controlled physiological environment that allows for translational insights into human reproductive and metabolic disorders. Among the various models used to study PMOS, the letrozole‐induced mouse model has emerged as a particularly valuable system due to its ability to recapitulate both the reproductive and metabolic phenotypes characteristic of the syndrome. ¹⁵ Unlike other PMOS models that predominantly focus on either reproductive or metabolic features, the letrozole model offers a more comprehensive representation of the syndrome, ¹⁵ , ¹⁶ making it a robust platform for preclinical drug evaluation. The current protocol uses subcutaneous implants containing letrozole, allowing a slow and continuous release over a 90‐day period. When implanted before puberty, mice develop all the neuroendocrine features of PMOS, such as hyperandrogenemia, cystic‐like ovarian morphology, decreased corpora lutea, increased body weight, increased adiposity, increased circulating levels of LH, ¹⁵ and increased LH pulsatility. ¹⁷ Data collection from this model is usually taken across the first 35 days (5 weeks) of treatment, and this model recapitulates the PMOS phenotype during that period in mice. However, the phenotype using 90‐day subcutaneous implants over extended treatment periods remains uncharacterized in mice, and our preliminary data suggest that the effect diminishes before the end of the 90‐day release window.

The limited duration of action of the subcutaneous letrozole implants, combined with the need for surgical implantation, restricts their suitability for long‐term studies. Therefore, we first assessed the PCOS‐like phenotype after 70 days of treatment and then hypothesized that an oral letrozole administration would provide a more sustained and reproducible PCOS‐like phenotype, offering a non‐invasive alternative better suited for evaluating the efficacy of therapeutic interventions over extended experimental periods.

2. MATERIALS AND METHODS

2.1. Mouse care

Wild‐type C57BL/6 female mice were used. All mice were bred and housed (2–4 per cage) at the University of Otago's Biomedical Research Facility. They had free access to food (standard rodent chow: Envigo Teklad rodent diet 2918), water, and standard enrichment materials (plastic tubes and wooden chew bars). The mice were subject to standard light/dark cycles (12 h on/off; lights on at 06:30 a.m.) and a temperature‐controlled environment (22 ± 1°C). Ethical approval for animal use and all experimental protocols was obtained from the University of Otago Animal Ethics Committee (AUP‐24‐09).

Experiment 1 tested the effects of letrozole delivery via subcutaneous implants. Twenty wild‐type C57BL/6 female mice aged between 25 and 26 days at the beginning of the experiment (Day 0) were randomly assigned to either a placebo group (n = 10) or a letrozole (LET‐implant) group (n = 10) for monitoring estrous cyclicity and testosterone levels. Each treatment group was then divided into two subgroups to assess either insulin resistance (insulin tolerance test [ITT]) or LH concentrations.

Experiments 2 and 3 tested the effects of letrozole delivery via drinking water. Experiment 2 assessed metabolic and reproductive phenotype over a 70‐day treatment period. Thirteen wild‐type C57BL/6 female mice aged between 25 and 26 days at the beginning of the experiment (Day 0) were randomly allocated to either a placebo group (n = 7) or letrozole (LET‐DW) group (n = 6). Experiment 3 assessed longer‐term metabolic and reproductive phenotype and LH pulsatility over a 130‐day treatment period. Ten wild‐type C57BL/6 female mice aged between 25 and 26 days at the beginning of the experiment (Day 0) were randomly allocated to either a placebo group (n = 5) or letrozole (LET‐DW) group (n = 5).

2.2. Subcutaneous implants of letrozole (Experiment 1)

Mice received a custom‐made 90‐day slow‐release subcutaneous implant (Innovative Research of America) containing letrozole (4.5 mg; AK Scientific Inc., #J10423; 99% purity) or placebo (matrix‐containing implant only) at 25–26 days of age. The letrozole implants provided a dose of approximately 0.05 mg/day, based on the 4.5 mg total letrozole content over the 90‐day release period. Briefly, mice were anaesthetized with 2% isoflurane, and a small incision was made in the medial dorsal region between the scapulae. A subcutaneous pocket was created caudal to the incision using blunt dissection, and the implant was inserted into the pocket; then the skin was closed using 1‐2 sutures and wound clips.

2.3. Oral administration of letrozole via drinking water (Experiments 2 and 3)

All mice received the treatment (0.03 mg/day) via their drinking water, assuming that they drink ~4 mL per day (from our previous observations using 0.4% Dimethyl sulfoxide (DMSO) in drinking water).

Letrozole (#J10423, 99% purity, AK Scientific Inc.) was used at a concentration of 7.5 μg/mL (0.4% DMSO). The treatments were prepared simultaneously and stored in aliquots of 375 μg letrozole in 200 μL of DMSO for letrozole groups, or just 200 μL of DMSO for the control group. Aliquots were stored at −20°C, and thawed immediately before use. Drinking water was changed two to three times per week, with a single 200 μL aliquot of letrozole or DMSO vehicle added to each 50 mL water bottle (1 per cage). To prevent light‐induced degradation, drinking bottles were wrapped in aluminum foil throughout the treatment period.

2.4. Experimental design

A timeline of the three experiments with the different parameters assessed is summarized in Figure 1.

Timeline of the three experiments, with the different parameters assessed. ITT, insulin tolerance test; LH, luteinizing hormone.

2.4.1. Plasma testosterone concentration

Blood was collected from the tail using heparin‐coated capillary tubes to prevent coagulation, and the plasma was harvested and frozen at −20°C until future testosterone level assessment using testosterone rat/mouse Enzyme‐linked immunosorbent assay (ELISA) kit (Labor Diagnostika Nord, catalog no. AR E‐8000, RRID:AB_2916330) following the manufacturer's instructions. The absorbance was measured at a wavelength of 450 nm using a SpectraMax® ABS Plus microplate reader.

The sensitivity of the assay was 0.024 ng/mL. The intra‐assay coefficient of variation was 3.6% for the low quality control sample (average 1.06 ng/mL), and 4.0% for the high quality control sample (average 3.30 ng/mL). The inter‐assay coefficient of variation was 3.3% and 4.0% for the low and high quality control samples, respectively.

2.4.2. Uterus weight as a marker of circulating estradiol levels

Estradiol levels are typically reported to remain within the normal range in women with PMOS. Because letrozole is an aromatase inhibitor that might lead to depletion of estradiol levels in blood, a biological index of estradiol levels was assessed using uterine weight, ¹⁸ as there is currently no reliable commercial assay that is sensitive enough to measure circulating estradiol levels in mice. At the end of the experiment, uteri were dissected, trimmed of surrounding fat, and weighed.

2.4.3. Brain tissue fixation and immunohistochemistry

Mice were deeply anaesthetized with pentobarbital injection (100 mg/kg) intraperitoneally, and perfused transcardially with 20 mL of 4% paraformaldehyde in 0.1 M phosphate buffer (PB). Once perfused, brains were dissected and stored in well plates containing 4% paraformaldehyde for 24 h at 4°C, then transferred into a 30% sucrose solution in 0.1 M PB at 4°C.

Coronal brain sections were collected using a microtome with a freezing stage, into three identical sets of 30 μm free‐floating sections through the arcuate nucleus.

For androgen receptor staining

Sections were washed three times for 10 min in Tris‐buffered saline (TBS), followed by incubation in TBS 0.3% Triton X‐100 (TX) for 10 min. The sections were then incubated in TBS 0.3% TX 1% Bovine serum albumin (BSA) with 10% Fetal bovine serum (FBS) for 60 min. After washing, endogenous peroxidases were deactivated using a solution of 1% hydrogen peroxide in TBS 0.3% TX with 40% methanol for 30 min. Following another wash, sections were incubated with an anti‐rabbit androgen receptor (AR) antibody (Abcam Cat# ab105225, RRID:AB_2782991), 1:1000 in TBS 0.3% TX 1% BSA and 2% normal goat serum for 48 h at 4°C on an orbital shaker. Sections were washed three times in TBS and incubated with biotinylated goat anti‐rabbit secondary antibody (Abcam Cat# ab6720, RRID:AB_954902) 1:200 dilution TBS 0.3% TX 1% BSA for 120 min. After washing, sections were incubated in A/B vectastain Elite solution (VECTASTAIN® Elite® ABC‐Horseradish peroxidase (HRP) Kit, Peroxidase (Standard) #PK‐6100) for 120 min, followed by a final wash. Finally, sections were incubated in DAB solution (SigmaFast™ 3,3'‐Diaminobenzidine (DAB) with metal enhancer, Sigma‐Aldrich, #D0426) until staining was judged to be adequate (approximately 40 min).

For progesterone receptor staining

Sections were washed three times for 10 min in TBS, followed by incubation in TBS 0.3% TX for 10 min. Endogenous peroxidases were deactivated using a solution of 0.9% hydrogen peroxide/40% methanol in TBS for 10 min. After washing, sections were incubated in TBS 0.3% TX 0.25% BSA and 2% normal goat serum for 60 min. Following another wash, sections were incubated with an anti‐rabbit progesterone receptor (PR) antibody (R.E. Campbell—University of Otago Cat# RC269, RRID:AB_2924988) 1:2500 dilution in TBS 0.3% TX 0.25% BSA and 2% normal goat serum for 72 h at 4°C on an orbital shaker. Sections were washed three times in TBS and incubated with biotinylated goat anti‐rabbit secondary antibody (Vector Laboratory, 3BA‐9200, RRID:AB_2313606) 1:250 dilution in TBS 0.3% TX 0.25% BSA for 60 min. After washing, sections were incubated in A/B vectastain Elite solution (VECTASTAIN® Elite® ABC‐HRP Kit, Peroxidase (Standard) #PK‐6100) 1:100 dilution for 60 min, followed by a final wash. Finally, sections were incubated in DAB solution (SigmaFast™ DAB with metal enhancer, Sigma‐Aldrich, #D0426) until staining was judged to be adequate (approximately 5 min).

2.4.4. Androgen receptor and progesterone receptor staining counting using QuPath

Image‐J (version 1.53t; https://imagej.net/) and QuPath (version 0.5.1; https://qupath.github.io/) software were used to quantify AR and PR staining within brain sections, focusing on the arcuate nucleus and adjacent areas. QuPath was employed for automated cell detection and the analysis of staining intensity across regions of interest. Image‐J was used for image processing and selecting and adjusting the region of interest. The settings were configured in QuPath to brightfield Nickel DAB with the correct pixel size based on the objective of the images taken (4× objective). The detection parameters were then optimized within QuPath to distinguish AR staining from background noise. The parameters used are shown in Supporting Information S1: Figure 1.

2.4.5. Body weight

Body weight was recorded six times throughout the duration of the experiment.

2.4.6. Fasted baseline blood glucose levels and insulin tolerance tests

Mice were fasted for 5 h (09:00 a.m.–2 p.m.) to assess the fasting level of glucose, and then insulin responsiveness was assessed by an ITT, which indicates the ability of insulin to decrease blood levels of glucose. Insulin was injected at 2 p.m. (0.25 U/kg, sc, Sigma‐Aldrich; #15523; insulin from porcine pancreas). Blood glucose samples were taken from the tail at −15 (baseline) and immediately prior to insulin injection, and then again at 15, 30, and 60 min following insulin injection, and analyzed using a glucometer (CareSens N, model GM505PAD).

2.4.7. Body composition assessment

Mice were briefly restrained (approximately 2 min) in a red cylindrical tube that limited voluntary movement. The restrained animals were then placed into the Bruker Minispec body composition analyzer to perform non‐invasive quantitative magnetic resonance (QMR) scanning. This method yields three primary output values: fat mass, lean mass, and free body fluid mass.

2.4.8. Vaginal cytology

The stage of the estrous cycle was determined by light microscopic analysis of the predominant cell type obtained from 5 μL physiological saline vaginal lavages. Mice were lavaged daily at ~09:00 a.m. during blocks of 10–20 days throughout the experiment. Proestrus was defined as the presence of predominantly nucleated and some cornified epithelial cells, estrus as mostly cornified cells, and diestrus/metestrus as some cornified epithelial cells and primarily leukocytes.

2.4.9. LH pulsatility

Blood samples were collected, in the diestrus stage, every 4 min over a 2‐h period (21 samples per mouse). A total of 4 μL of blood (per sample) was obtained from the tip of the tail and immediately transferred into pre‐prepared Eppendorf tubes containing 56 μL of Phosphate‐Buffered Saline with Tween 20 (PBS‐T) buffer and put in dry ice until the end of the collection time, and then transferred at −20°C until assay. To control for estrous cycle‐dependent fluctuations in LH secretion, vaginal cytology was performed prior to blood sampling to confirm that mice were in diestrus. Sampling occurred at approximately 1000–1200 h to minimize any circadian fluctuations.

2.4.10. LH Enzyme‐linked immunosorbent assay (ELISA)

For the LH ELISA, we followed the method described by Kreisman et al. ¹⁹ using a sensitive sandwich ELISA approach to assess whole blood LH concentrations. On Day 1, a 50 μL coating of bovine LH beta subunit (β) monoclonal antibody (#518B7 from Lillian E. Sibley, UC Davis, 1:1000 in 1× PBS) was added to each well and incubated overnight at 4°C. On Day 2, the plate was decanted and incubated with 200 μL of blocking buffer (5% skim milk powder in PBS‐T) per well, incubated at room temperature for 2 h (all incubations were within a humidified chamber on an orbital mixer set at 40–50 rpm), and then washed three times using PBS‐T. Standards and samples were loaded (50 μL/well) and incubated for 2 h at room temperature. After washing, 50 μL of biotinylated mouse monoclonal detection antibody (#5303 SPRN‐5, 1:1000 in blocking buffer with 0.2% BSA) was added, followed by a 90‐min incubation. A secondary poly‐HRP conjugated streptavidin antibody (50 μL, 1:8000 in PBS‐T) was added to each well and incubated for 90 min. After washing, 100 μL of o‐Phenylenediamine (OPD) substrate (one tablet of OPD in 6 mL of citrate buffer +6 μL of 30% H₂O₂) was added and incubated in the dark for 30 min, followed by stopping the reaction with 50 μL of 3 M HCl per well. Absorbance was read at 490 and 650 nm using the SpectraMax® ABS Plus microplate reader. A standard curve was generated using 4‐parameter logistic curve fitting. Output values were adjusted to account for the 15‐fold dilution. The sensitivity of the assay was 0.031 ng/mL, and the average intra‐assay coefficient of variation for the low (0.98 ng/mL) and medium (1.58 ng/mL) values were 9.38% and 2.49%, respectively. The inter‐assay coefficient of variation for the same values were 10.19% and 4.01%, respectively.

LH pulses were defined as an increase exceeding 30% above baseline and displaying an absolute increase from the preceding nadir greater than three times the intra‐assay coefficient of variation, corresponding to a minimum amplitude threshold of ~0.28 ng/mL at low concentrations. Additionally, all values considered for pulse detection were above the assay sensitivity (0.031 ng/mL).

For each identified pulse, amplitude was calculated as the difference between the peak LH concentration and the baseline value immediately preceding pulse onset. Mean LH levels were defined as the average of all measurements collected throughout the sampling period.

2.4.11. Ovarian histology

At the end of the experiment, ovaries were dissected and trimmed of surrounding fat and then stored within 70% ethanol for further histological analysis. Ovaries were then embedded within paraffin and sectioned at 5 μm intervals using a microtome, with every 10th section collected to capture the full extent of the ovary. Sections were stained with hematoxylin and eosin (H&E) using an Epredia Gemini automated slide stainer, followed by coverslipping with a mounting medium composed of dibutylphthalate, polystyrene, and xylene. Images of the sections were taken using an Olympus BX‐51 light microscope. Ovarian structures were classified across serial tissue sections into distinct morphometric categories based on defined cellular and structural criteria. To prevent double‐counting bias, each follicle was counted once, in the section where it was first identified. Preceding and following sections were then examined to confirm that the structure was a single continuous follicle rather than being recorded as separated structures across the series.

Primordial and primary follicles were excluded from analysis because they lack FSH receptors and their transition out of the resting pool is governed entirely by local intraovarian factors, independently of gonadotropins. ²⁰ Secondary follicles were identified by the presence of a central oocyte surrounded by two or more solid, concentric layers of cuboidal granulosa cells with no fluid space. Early antral follicles were defined by the initial appearance of small, isolated fluid‐filled pockets within the granulosa layers, which expanded into a single, continuous fluid cavity (antrum) in healthy antral follicles. Mature pre‐ovulatory follicles were characterized by an exceptionally large antrum, an eccentric oocyte anchored by an intact cumulus stalk. Cystic follicles were characterized as large, fluid‐filled follicles exhibiting a thickened theca cell layer but entirely lacking both a cumulus mass of granulosa cells and an oocyte. Finally, active corpora lutea were identified as large, solid clusters of luteal cells indicating recent ovulation, whereas regressing corpora albicans were classified as small, degenerated luteal scars from older reproductive cycles.

2.5. Data presentation and statistical analysis

All the graphs and statistical analysis were done using Prism (Graphpad Software, San Diego, CA, USA) data processing and management software. All data are presented as mean ± standard error of the mean (SEM). For all results, the significant threshold was placed at α = .05.

A Mann–Whitney test was used for single point measurements such as testosterone, uterine weight, glucose concentration area over the curve (AOC), LH concentrations, LH pulse frequency, LH pulse amplitude, body composition data, ovarian histology data, and fasting blood glucose. The Mann–Whitney test was chosen because the data distribution did not follow a Gaussian distribution and did not meet the assumptions of equal variance required for parametric tests.

A two‐way Analysis of variance (ANOVA) or mixed‐effect model with the Geisser–Greenhouse correction where appropriate was applied for data involving repeated measurements over time such as body weight, estrous cyclicity, testosterone, and ITT glucose concentrations, with time and letrozole treatment as factors. Šídák's multiple‐comparisons test was used for comparisons between treatment groups at each time point, with adjusted p‐values reported to control the family‐wise error rate.

3. RESULTS

3.1. Experiment 1: Limited effective duration from letrozole delivered via subcutaneous implants

3.1.1. Validation

There was no significant time × treatment interaction for plasma testosterone concentrations (F _1,18 = 1.189; p = .29). Post hoc comparisons showed that serum testosterone concentrations were significantly higher in the LET‐implant group compared to controls after 30 days of treatment (p < .05), but by 50 days of treatment this effect had disappeared (p = .53) and the levels were similarly low in both groups (Figure 2A).

Effect of subcutaneous implants of letrozole (LET‐implant; ~0.05 mg/day) or placebo in mice (Experiment 1). Testosterone blood levels after 30 and 50 days of treatment (A), uterus weight after 70 days of treatment (B), body weight during the experiment (C), blood levels of glucose after insulin injection performed after 70 days of treatment (D), representative example of an estrous cycle in each group during the 70 days treatment (E), quantification of the number of estrous cycles during 7–28, 28–49, and 49–70 days of treatment (F), individual profile of luteinizing hormone (LH) secretion in the blood, with each red dots representing a pulse of LH (G), LH blood concentration (H), and number of LH pulses in 120 min (I). Statistical comparisons were made using Mann–Whitney test for (A), (B), and (G), and using a two‐way ANOVA followed by a Sídák's multiple‐comparisons test, with time and treatment as factors, for (C), (D), and (F). *p < .05, **p < .01, ***p < .001. n = 10 per group for (A), (B), (C), and (E/F). The groups have been divided into two subgroups of five animals per group for (D) and (H) and (I). E, estrus; M/D, Met/diestrus; P, proestrus.

Uterus weight was used as a proxy for circulating estrogen concentrations. No significant difference in uterine weights was identified between LET‐implant treated‐mice and controls after 70 days of treatment (0.057 ± 0.006 g vs. 0.063 ± 0.004 g, respectively) (p = .10) (Figure 2B).

3.1.2. Metabolic phenotype

There was a significant effect of time (F _1.369,23.82 = 257.1; p < .001) and treatment (F _1,18 = 16.64; p < .001) on body weight, with mice receiving LET‐implant being heavier than control mice (F _1.369,23.82 = 7.180; p < .01) (Figure 2C). The mean body weight of LET‐implant mice was significantly heavier from Day 19 of treatment (p < .05) compared to control mice, but by Day 70, the effect of the letrozole implants was no longer evident (p = .32).

There was no statistically significant difference between LET‐implant and control groups in fasting blood glucose levels (time 0 values) on Day 70 of treatment (p > .99) (Figure 2D). An ITT was performed on Day 70 of the treatment. There was a main effect of time (F _2.378,19.02 = 19.72, p < .001) after insulin injection, but no main effect of the treatment (F _1,8 = 0.05, p = .82), and no main effect of time × treatment interaction (F _2.378,19.02 = 0.14, p = .89) (Figure 2D).

3.1.3. Reproductive phenotype

The number of estrous cycles observed during Days 7–28 and 28–49 of treatment differed significantly between the LET‐implant and control groups, with controls exhibiting more cycles than the letrozole‐treated mice (p < .001 for both intervals; Figure 2E,F). By Days 49–70, this difference was no longer statistically significant (p = .056; Figure 2F). Within the control group, the number of estrous cycles did not differ significantly across the three observation periods (7–28, 28–49 and 49–70 days; p = .776, p = .702 and p = .846, respectively). In contrast, the number of estrous cycles in the LET‐implant group increased significantly over time, with more cycles observed during Days 28–49 than Days 7–28 (p < .05) and during Days 49–70 than Days 28–49 (p < .01) (Figure 2E,F).

There was no significant difference in the mean LH concentrations between LET‐implant mice and control mice at diestrus stage after 65–70 days of treatment (0.82 ± 0.14 and 0.82 ± 0.12 ng/mL, respectively) (Figure 2G,H). Similarly, LH pulse frequency did not differ significantly between LET‐implant and control mice (2.2 ± 0.583 and 1.6 ± 0.245 pulses per 120 min, respectively) (Figure 2G,I).

3.2. Experiment 2: Efficacy of letrozole delivered in drinking water (LET‐DW)

3.2.1. Validation

There was no significant time × treatment interaction (F _1,11 = 3.476, p = .089) but there was a significant main effect of the LET‐DW treatment (F _1,11 = 170.3; p < .001). Serum testosterone concentrations were significantly higher in the LET‐DW group compared to controls at both Day 24 (p < .001) and Day 62 (p < .001) (Figure 3A).

Effect of letrozole (~0.03 mg/day during 60 days) and placebo treatment in drinking water in mice (Experiment 2). Testosterone after 24 and 62 days of treatment (A), and uterus weight (B). Representative example of androgen receptor (AR) immunostaining, top and middle panel, and of progesterone receptor (PR) staining, bottom panel, in the arcuate nucleus of mice treated for 70 days with placebo or letrozole (C), average androgen receptor positive (AR+) cells per section in each group (D), average progesterone receptor positive (PR+) cells per section in each group (E), body weight during the experiment (F), fat mass, free body fluid, and lean mass expressed in percentage of total body weight after 70 days of treatment (G), fasting blood levels of glucose after 62 days of treatment (H), blood levels of glucose after insulin injection performed after 62 days of treatment (I), analysis of insulin tolerance test with area over the curve (J), representative example of an estrous cycle in mice treated with placebo or letrozole in their drinking water during 70 days (K). Number of estrous cycles overserved during 60‐days period analysis (L). Time spent in each stage of the estrous cycle during the 60‐days period analysis (M). Number of cystic‐like follicles per ovary (N) and number of corpora lutea per ovary (O) in mice from each group, after 70 days of treatment. The insert shows high‐magnification image of the white dashed‐line square. Statistical comparisons were made using Mann–Whitney test for (B, D, E, G, H, J, L N, O), and using a two‐way ANOVA followed by a Šídák's multiple‐comparisons test, with time and treatment as factors, for (A, F, and I), or with stages of the estrous cycle and treatment as factors for (M). *p < .05, **p < .01, ***p < .01. n = 6–7 per group. Scale bar, 100 μm. E, estrous; M/D, metestrus/diestrus; P, proestrus.

Uterus weight was used as a proxy for circulating estrogen concentrations. No significant difference in uterine weights was identified between LET‐DW‐treated mice and controls after 70 days of treatment (0.056 ± 0.017 g vs. 0.077 ± 0.021 g, respectively) (p = .46) (Figure 3B).

3.2.2. Steroid receptors expression

Androgen receptor

Coronal brain sections demonstrated distinct AR immunoreactivity, especially in the rostral arcuate nucleus (Figure 3C, representative example for each group, with top and middle panel depicting the rostral and mid‐ARC regions, respectively). Quantitative analysis of AR positive cells revealed a significant increase in the LET‐DW groups compared to controls after 70 days of treatment (p < .001), with control animals showing an average of 224 ± 32.93 AR+ cells per section, and letrozole animals showing an average of 541 ± 58.60 AR+ cells per section (Figure 3D).

Progesteron receptor

PR expression in the arcuate nucleus was significantly reduced following chronic LET‐DW treatment. The number of PR‐positive cells was lower in LET‐DW‐treated mice compared with controls (279.8 ± 76.9 vs. 536.1 ± 135.7 PR + cells per section, respectively, p < .01) (Figure 3C, representative example for each group in the bottom panel, and Figure 3E).

3.2.3. Metabolic phenotype

There was a significant effect of time (F _1.643,18.73 = 357.6; p < .001), treatment (F _1,12 = 21.85; p < .001), and interaction of time × treatment (F _1.643,18.73 = 7.682; p < .01) on body weight, with mice receiving letrozole in drinking water being heavier than those in the control group (Figure 3F). The mean body weights of LET‐DW‐treated mice were significantly heavier compared to body weights of control mice, from 40 days of treatment (p < .05), until 70 days (end) of treatment (p < .01) (Figure 3F).

Analysis of body composition revealed significant alterations in body fat mass with LET‐DW‐treated mice exhibiting significantly higher fat mass percentage (14% of bodyweight) compared to the control group (10% of bodyweight) (p < .05) (Figure 3G, left panel). In contrast, free body fluid percentage (Figure 3G middle panel) and lean mass percentage (Figure 3G right panel) showed no differences between groups.

There was no statistically significant difference between each treatment group in fasting blood glucose levels after 62 days of treatment (p = .079) (Figure 3H). An ITT was performed on Day 62 and revealed significant main effects of time (F _2.310,25.41 = 24.31, p < .001), letrozole treatment (F _1,11 = 9.027, p < .05), and time × letrozole interaction (F _2.310,25.41 = 4.208, p < .05) (Figure 3I). The quantification of the glycemic responses to the insulin challenge, using the AOC, resulted in a significant difference between groups, with the LET‐DW group having a lower decrease of glucose levels compared to controls (p < .05) (Figure 3J).

3.2.4. Reproductive phenotype

All control animals (7/7) demonstrated a regular 4–5 day estrous cycle throughout the entire observation period of approximately 60 days (Days 10–70 of treatment). LET‐DW‐treated mice (6/6) showed persistent diestrus across the entire observation period (Figure 3K, representative example for each group) so that the control mice had significantly more estrous cycles compared to LET‐DW‐treated animals (p < .001) (Figure 3L).

The number of days spent in each stage of the estrus cycle was significantly different between control and LET‐DW groups (F _2,30 = 694.3, p < .001) (Figure 3M). Control animals spent significantly more time in proestrus (p < .001), more time in estrus (p < .001), and less time in diestrus (p < .001) compared to LET‐DW‐treated animals (Figure 3M).

Mice treated with LET‐DW had significantly more cystic follicles on their ovaries compared to control mice (p < .001) (Figure 3N), and significantly fewer corpora lutea (p < .01) compared to control mice (Figure 3O).

3.3. Experiement 3: Long‐term efficacy of letrozole delivered in drinking water

Next, we investigated whether the PMOS phenotype could be maintained over several months, whether letrozole treatment increased LH pulsatility, and how ovarian morphology was affected.

As was the case for mice treated with letrozole in the drinking water in Experiment 2, LET‐DW‐treated mice in Experiment 3 displayed an increased body weight that was still evident 130 days after letrozole treatment onset (p < .01; Figure 4A), an absence of cyclicity extending at least 80–95 days after treatment onset (p < .01; Figure 4B) and elevated testosterone measured at 127 days of treatment (p < .001; Figure 4C) compared to control mice.

Long‐term efficacy of letrozole (~0.03 mg/day during 130 days) delivered in drinking water (Experiment 3). Body weight during the experiment (A), number of estrous cycles observed during the 15 days period analysis, after 80 days of treatment (B), testosterone blood levels after 127 days of treatment in each group (C). Representative example of luteinizing hormone (LH) pulsatility in mice after 127 days of treatment with placebo or letrozole. Each red dots represents a pulse of LH (D). Average LH blood concentrations (E), total number of pulses during the 120 min sampling period (F), and average pulse amplitude (G) in mice from each group. Representative example of the ovarian morphology in mice treated with placebo or letrozole during 130 days, Scale bar, 200 μm. The insert shows high magnification image of the black dashed‐line square (H), length of the ovaries (I), total count of secondary, early antral, antral and pre‐ovulatory follicles, corpora lutea, corpora albicantia, atretic follicles, and hemorrhagic cysts per ovary (J) in mice from each group. Statistical comparisons were made using Mann–Whitney test for (B), (C), (E), (F), (G), (I), (J), and using a two‐way ANOVA followed by a Šídák's multiple‐comparisons test, with time and treatment as factors, for A. *p < .05, **p < .01, ***p < .001. n = 5 per group. AF, atretic follicle; CL, corpus luteum; HC, hemorrhagic cysts; SF, secondary follicle.

3.3.1. LH pulsatility

LH concentration profiles, after 127 days of treatment, demonstrated elevated baseline levels and increased pulse frequency in LET‐DW‐treated animals compared to controls (Figure 4D, representative example for each group). Quantitative assessment revealed significantly higher mean LH concentrations in LET‐DW‐treated mice (1.943 ± 0.035 ng/mL) compared to controls (0.372 ± 0.035 ng/mL, p < .01) (Figure 4E). Similarly, pulse frequency was significantly increased in the LET‐DW group (3.75 ± 0.32 pulses per 120 min period) compared to controls (0.80 ± 0.32 pulses per 120 min period, p < .01) (Figure 4F), accompanied by significantly greater pulse amplitude (0.737 ± 0.098 ng/mL vs. 0.194 ± 0.098 ng/mL, p < .01) (Figure 4G).

3.3.2. Ovarian morphology

Mice treated with LET‐DW had significantly enlarged ovaries compared to control mice (p < .05) (Figure 4H,I). Mice treated with LET‐DW also have significantly more atretic follicles on their ovaries (p < .001; Figure 4H,J), more hemorrhagic cysts (p < .001), significantly fewer corpora lutea (p < .05), and significantly fewer secondary follicles (p < .01) compared to control mice (Figure 4H,J).

4. DISCUSSION

The letrozole‐induced mouse model is a well‐established experimental paradigm that reliably produces reproductive and metabolic disturbances characteristic of PMOS. ¹⁵ , ¹⁶ However, the duration and consistency of the phenotype depend on the method of letrozole administration. In mice, letrozole is commonly administered by daily oral gavage ²¹ , ²² , ²³ or continuous subcutaneous slow‐release implants. Previous work in our laboratory has utilized custom‐made 4.5 mg, 90‐day continuous‐release letrozole implants in C57BL/6 mice, demonstrating initial efficacy. However, the present findings reveal important limitations of this approach: testosterone levels were no longer elevated beyond ~50 days of treatment, estrous cyclicity resumed around this time, and the metabolic phenotype (namely increased body weight and insulin resistance) was largely resolved by ~70 days. Although implants can induce a PMOS‐like phenotype, their short duration of action limits their use for long‐term studies. These findings therefore highlight the need for an administration strategy that can maintain letrozole efficacy over longer treatment periods without the additional procedural requirements associated with implantation.

In the present study, we addressed this limitation by delivering letrozole via the drinking water. The drinking water administration provides a practical advantage over existing approaches by avoiding either surgical implantation or stressful repeated oral gavage, ²⁴ , ²⁵ making prolonged letrozole treatment simpler and more accessible for laboratories without specialized surgical facilities by avoiding the material and labor costs associated with preparing and surgically implanting slow‐release devices, which may be particularly relevant for studies requiring larger cohorts or prolonged treatment periods. By eliminating these handling and surgical procedures, this method also minimizes physical and psychological stressors known to alter gonadotropin secretion and neuroendocrine outcomes. ²⁶ Although mice consume the majority of their daily water intake (~4 mL per mouse) during their active (dark) phase, testosterone remained elevated in samples collected during the light phase, supporting sustained pharmacological efficacy throughout the treatment period. This is notable given the relatively short half‐life of letrozole in mice compared with humans (~2.6 vs. ~42 h, reflecting faster hepatic and renal clearance), ²⁷ and indicates that repeated intake through drinking water was sufficient to maintain the hyperandrogenism state despite its rapid clearance in mice.

This approach maintained elevated testosterone concentrations throughout the full duration of treatment (70 and 127 days in Experiments 2 and 3, respectively), in contrast to the declining efficacy observed with implants. Importantly, estrous cyclicity remained disrupted throughout treatment, whereas cyclicity resumed after approximately 50 days with implant‐based delivery. This sustained disruption better reflects the chronic nature of PMOS, which manifests as a persistent endocrinopathy rather than a transient condition. ²⁸ Thus, letrozole administered in drinking water extended the effective duration of the phenotype beyond that achieved with the letrozole implants, providing an approach suitable for studies requiring prolonged treatment.

Our data provide some insight into the temporal development of the letrozole‐induced phenotype. Alterations in oestrous cyclicity were evident after approximately 10 days of treatment, whereas differences in body weight emerged only after approximately 40 days. This temporal pattern suggests that reproductive dysfunction develops earlier than the detectable metabolic phenotype in this model. However, these observations do not necessarily define the earliest onset of either phenotype, as changes may occur before the time points assessed. Moreover, because LH pulsatility, insulin sensitivity, and other metabolic and neuroendocrine parameters were not assessed longitudinally, the present study cannot determine when these components of the phenotype first emerge. Further time‐course studies will therefore be required to establish the sequence and timing of neuroendocrine, reproductive, and metabolic alterations following the onset of chronic hyperandrogenism.

Hyperandrogenism in this model was accompanied by profound reproductive dysfunction. Mice exhibited persistent diestrus with complete absence of proestrus and estrus, reflecting the oligo/anovulation characteristic of PMOS. ²⁹ Ovarian morphology was also altered, with increased cystic‐like ovarian morphology, absence of corpora lutea, reduction of secondary follicles and ovarian enlargement. ³⁰ The reduced number of secondary follicles may reflect disrupted follicular development and increased follicular arrest or atresia associated with chronic hyperandrogenism. Together these changes suggest altered follicular dynamics and impaired progression toward ovulation.

The extended phenotype recapitulated the neuroendocrine dysfunction and included metabolic abnormalities, hallmarks of PMOS. Letrozole‐treated mice displayed significantly elevated LH concentrations, along with increased LH pulse frequency and amplitude, consistent with dysregulated GnRH/LH secretion observed in women with PMOS and in other letrozole models. ¹⁵ , ¹⁷ Letrozole‐treated mice also exhibited significant weight gain and increased adiposity, without changes in lean mass or fluid content, indicating a selective fat‐centric metabolic alteration. Despite unchanged fasting glucose levels, insulin tolerance testing revealed marked insulin resistance in letrozole‐treated mice. In early or moderate insulin resistance, compensatory hyperinsulinemia can maintain normoglycemia, ³¹ providing a plausible explanation for the lack of treatment effect on fasting glucose levels. Unfortunately, insulin measurements were not conducted in our study to confirm this. This metabolic impact aligns with clinical observations that hyperandrogenism severity correlates with metabolic dysfunction in women with PMOS. ³² Consistent with clinical data, a substantial proportion of women with PMOS (estimated at 50%–80%) exhibit insulin resistance, with prevalence generally higher in those with obesity. ³³

The neuroendocrine findings were further supported by changes in AR expression in the arcuate nucleus. Elevated circulating testosterone was associated with increased AR positive cell density in the arcuate nucleus, which plays a central role in GnRH regulation. AR expression in this region is known to reflect circulating androgen levels, ¹⁶ , ³⁴ and may contribute to neuroanatomical remodeling that drives altered gonadotropin secretion. ³⁵ Similar central AR upregulation has been reported in other letrozole models and is thought to contribute to elevated GnRH/LH pulse frequency. ³⁶

Chronic letrozole treatment also significantly reduced PR expression in the arcuate nucleus. Progesterone signaling in this region contributes to negative feedback regulation of GnRH/LH pulsatility, with arcuate KNDy neurons playing a central role in the control of the GnRH pulse generator. Given that increased GnRH/LH pulsatility is a characteristic feature of PMOS, reduced PR expression may indicate altered progesterone responsiveness within the hypothalamic reproductive axis and provide a potential link between the letrozole‐induced androgenic phenotype and disrupted steroid feedback. However, reduced PR expression should not be interpreted as direct evidence of impaired progesterone feedback, as recent evidence indicates that PR deletion from most arcuate kisspeptin/KNDy neurons does not abolish progesterone‐mediated suppression of LH. ³⁷ This suggests that progesterone feedback may also involve other PR‐expressing neuronal populations or non‐genomic mechanisms.

Although estradiol concentrations were not directly measured, maintained uterine weights suggest that uterine estrogenic activity was not substantially reduced, as uterine weight is highly responsive to circulating estrogen levels. ¹⁸ However, uterine weight is not a specific measure of circulating estrogen levels, as other hormonal changes induced by letrozole may also contribute to maintaining uterine mass, including potential effects of AR signaling. Thus, the maintained uterine weight suggests that severe loss of uterine estrogenic stimulation is unlikely, but it cannot establish that circulating estrogen concentrations remained within the normal range. ³⁸ Direct measurement of circulating estrogen concentrations would be required to determine the extent to which estrogen levels were altered by chronic letrozole treatment.

Importantly, our findings provide evidence for an alteration in the hypothalamic component of the reproductive axis, but do not establish whether the increased LH pulsatility observed following chronic letrozole treatment results from enhanced sensitivity of the GnRH pulse generator, altered pituitary responsiveness, or both. Similarly, changes in circulating LH concentrations or LH pulse frequency cannot distinguish between altered hypothalamic GnRH drive and pituitary LH releasable pool. Future studies using exogenous kisspeptin and GnRH challenges could help distinguish altered hypothalamic drive from altered pituitary responsiveness.

Taken together, our findings demonstrate that delivering letrozole through drinking water provides a simple and sustained approach for generating a prolonged PMOS‐like phenotype in mice. This approach maintained reproductive, ovarian, neuroendocrine, and metabolic abnormalities for at least 127 days without surgical implantation or repeated gavage. Drinking‐water delivery therefore provides a practical and non‐invasive alternative for long‐term studies of chronic hyperandrogenism and PMOS.

AUTHOR CONTRIBUTIONS

KAS: investigation, formal analysis, visualization, writing—original draft. RK: investigation, formal analysis. GMA: funding acquisition, resources, supervision. CD: funding acquisition, project administration, supervision, conceptualization, methodology, investigation, formal analysis, visualization, writing—original draft—review and editing.

FUNDING INFORMATION

This study was supported by the Health Research Council of New Zealand (HRC) project grant (23/049).

CONFLICT OF INTEREST STATEMENT

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Supporting information

Figure S1. QuPath cell detection parameters used for androgen receptor and progesterone receptor staining analysis.

JNE-38-e70294-s002.pdf^{(60.7KB, pdf)}

Table S1. F statistics, degrees of freedom, and p values for all ANOVAs.

JNE-38-e70294-s001.pdf^{(22.4KB, pdf)}

ACKNOWLEDGMENTS

The authors thank Megan Inglis for technical assistance and the Biomedical Research Facility team for breeding and taking care of the mice. Open access publishing facilitated by University of Otago, as part of the Wiley ‐ University of Otago agreement via the Council of Australasian University Librarians

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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