Neonatal Mouse Ovary Culture and Whole-mount Follicle Quantification: An Efficient In Vitro Approach for Studying Primordial Follicle Regulation

Summary

This study presents protocols for culturing postnatal mouse ovaries on membrane inserts and for whole-mount follicle quantification in an entire ovary. Results demonstrated that technical handling during ovary culture significantly affects primordial follicle counts, highlighting the importance of maintaining consistency to minimize non-biological variation.

Abstract

In female mammals, primordial follicles form during fetal ovarian development and serve as the sole source for sustaining adult ovarian function. Mechanisms underlying how primordial follicles assemble, maintain dormancy, activate for follicular development, and undergo cell death are important for understanding ovarian physiology and pathological conditions. This study presents a protocol for culturing postnatal mouse ovaries on membrane inserts, an approach that enables the culture and pharmaceutical treatment of intact ovaries for up to 10 days, depending on the developmental stage of the ovary. Changes in the culture conditions can be achieved by transferring inserts containing cultured ovaries between wells on a plate, thereby avoiding physical interference with the tissues during culture. P5 ovaries were isolated and cultured on a 12 mm insert in a 24-well plate as an example. Each ovary was separated within a droplet of DMEM/F12 medium supplemented with 10% FBS, 3 mg/mL BSA, 10 mIU/mL FSH, and Antibiotic-Antimycotic, and gently stabilized on the membrane insert. The medium was changed every two days, and the culture was maintained for a total of five days. Following the culture, ovaries were fixed in 4% paraformaldehyde for 2 h and processed for whole-mount antibody staining of the oocyte marker DDX4. Follicles were staged and quantified based on oocyte size and the nuclear morphology of somatic follicle cells. The results showed that the number of primordial follicles in each ovary was significantly affected by the proper placement of tissues on the membrane insert. In addition, differences in the number of ovaries on each insert may introduce non-biological variations and should be avoided.

Introduction

Oogenesis is a highly regulated and complex process that results in the production of mature oocytes. In mammals, oogenesis initiates in fetal ovaries, where primordial germ cells (PGCs) differentiate into primary oocytes. Each primary oocyte is enclosed by a layer of squamous pregranulosa cells, forming a primordial follicle. Most of these primordial follicles become dormant after formation, serving as the ovarian reserve^1,2,3. In the adult ovary, periodic activation of primordial follicles for follicular development, known as folliculogenesis, is essential for sustaining the production of mature oocytes and ovarian steroid hormones ^4,5. In the absence of activation signals, human primordial follicles can remain quiescent for up to 50 years. The prolonged dormancy may contribute to reduced oocyte quality, as well as an increased rate of primordial follicle loss in aging ovaries⁶. The mechanisms underlying how a specific pool of follicles becomes activated while others stay quiescent and undergo periodic cell death remain open questions.

The process of oogenesis is highly conserved in mammals, making mice an ideal model for studying mammalian oogenesis⁷. In mice, primordial follicle formation is complete by postnatal day 4 (P4). A small proportion of primordial follicles undergo follicular development immediately after their formation. During follicle development, the oocyte increases in size due to enhanced organogenesis, and mRNA and protein synthesis⁴. Squamous granulosa cells transition to cuboidal granulosa cells and become proliferative⁸. These developing follicles grow to the ovulatory stage around day 21, which is when female mice reach puberty. This phase of follicular development in postnatal ovaries is commonly referred to as 'first-wave folliculogenesis.' Since first-wave folliculogenesis closely mirrors the process of follicle development, postnatal mouse ovaries provide an ideal model for studying primordial follicle regulation and ovarian folliculogenesis⁹.

The classification of ovarian follicles is primarily based on the nuclear morphology of follicle somatic cells and the size of the oocyte. In mouse ovaries, the dormant primordial follicle is identified by a single layer of squamous pregranulosa cells that encase a primary oocyte, which has a diameter of approximately 20 µm. Developing follicles can be categorized into primary, secondary, tertiary, and antral follicles. The primary follicle contains an oocyte that is typically over 25 µm in diameter and is surrounded by a single layer of cuboidal granulosa cells (Supplementary Figure 1). The secondary follicle features two layers of granulosa cells, while the tertiary follicle comprises three or more layers of granulosa cells. The antral follicle is larger than the tertiary follicle, contains multiple layers of granulosa cells, and has a fluid-filled cavity known as the antrum. As follicles develop, the oocyte increases in size, reaching approximately 80 µm by the time of ovulation. Additionally, the developing follicles are encircled by several layers of thin theca cells, which are located outside the granulosa cells^10,11,12.

The relatively small size of postnatal mouse ovaries makes whole ovary culture a practical approach for research. This method effectively enables the study of ovarian and follicle development within an intact ovary, as it preserves the physical and physiological microenvironments while minimizing interference from surrounding tissues¹². This approach allows for experiments that would be challenging to conduct in in vivo models. Examples include live imaging of ovarian development, time-controlled multi-drug treatments, and the analysis of ovarian secretory activity through protein profiling of the culture media^13,14. Furthermore, this approach can be applied to investigate the effect of secretory factors without direct cell-cell interaction through co-culturing experiments. In these experiments, different types of tissues can be placed on separate membrane inserts within a shared medium¹⁵. Additionally, using conditional gene-knockout mouse ovaries in vitro could help elucidate key mechanisms involved in primordial follicle activation and growth.

In this article, we introduce methods for culturing postnatal mouse ovaries using membrane inserts and for whole-mount follicle quantification in an entire ovary (Figure 1). Techniques outlined here include 1) dissection of neonatal mouse ovaries, 2) culture of neonatal mouse ovaries, 3) media change during culture, 4) tissue fixation and whole-mount antibody staining, and 5) tissue imaging and follicle quantification. Additionally, this study employed doxorubicin to experimentally induce primordial follicle loss, providing a model to investigate the mechanisms underlying ovarian reserve depletion. The results showed that the number of ovaries per insert and their positioning within the insert during culture influenced primordial follicle counts. This highlights the importance of maintaining a consistent culture setup to prevent non-biological variations in the experimental outcomes.

Protocol

CD-1 male and female mice were housed together at a 1:1 ratio as breeding pairs. The date of birth of new pups was designated as postnatal day 0. All animal experiments were approved by the Institutional Animal Care & Use Committee (IACUC) at the University of Missouri (protocol number: 36647).

1. Reagents and Culture Media

1. All reagents used in this study are listed in the Table of Materials. Prepare all materials and media before ovary culture.
2. Culture media: Make ~5 mL of DMEM/F-12 media supplemented with 3 mg/mL bovine serum albumin (BSA), 10% fetal bovine serum (FBS), and 10 mIU/mL follicular-stimulating hormone (FSH).
   NOTE: The media should be freshly prepared by adding the BSA (30 mg/mL), FBS, and FSH stocks (10 IU/mL) to DMEM/F-12 media. Equilibrate the media in a petri dish for about 30 min in a 5% CO₂ incubator at 37 °C. Store the stocks at -20 °C and thaw them immediately before use. Aliquot stocks in proper volumes to avoid repeated freezing and thawing of the stocks.

2. Ovary culture

1. Euthanize P5 female mice by decapitation. Remove the ovaries and the attached bursa tissue using surgical scissors and forceps. Place the ovaries in a 60 mm petri dish with pre-chilled Dulbecco's phosphate-buffered saline (DPBS) containing Antibiotic-Antimycotic (Figure 2A,B).
2. Using a pair of insulin syringes (1 mL), dissect and remove the bursa tissue from the ovary under a stereoscope (Figure 2C,D). Transfer the ovaries using a 1 mL pipette to a 30 mm petri dish containing 3 mL of culture media and place the dish in a CO₂ incubator.
   NOTE: Avoid poking or squeezing the ovaries. Make clean and uniform incisions through the ovarian bursa using the insulin syringes to release the ovary. Prefill the pipette tip with approximately 50 µL of media, place the pipette tip next to the ovary, and aspirate it into the tip for transfer.
3. Under the laminar flow hood, prepare the plate by placing 400 µL of media and an insert in each well (Figure 2E,E'). The media is outside the insert, and the membrane of the insert turns opaque once it is soaked by the media.
4. For the culture with 24 h Dox treatment, prepare media containing Dox at 0.1 mg/mL concentration freshly by diluting Dox stock (10 mg/mL) with the media.
   NOTE: Media containing Dox should be disposed of properly in accordance with the guidelines of the institutional committees for biosafety and environmental health and safety.
5. Under a stereoscope, transfer up to three ovaries to an insert using a 1 mL pipette (Figure 2F,G). Minimize the amount of media transferred into the insert and immediately remove any excess media after transfer. Once the ovaries are gently placed in the insert, use a 200 µL pipette to transfer approximately 200 µL of media from the well into the insert, ensure the membrane is completely covered by the media. Transfer the media back to the well and center the ovaries on the membrane. Fill the pipette tip with culture media to prevent the ovary from being sucked into the tip when centering it. Make sure to leave adequate space between the ovaries and that they are stabilized on the membrane insert (Figure 2H), rather than remaining suspended in the medium (Figure 2I). Take care not to poke the ovaries and the membrane.
6. In a 5-day culture, change media every two days by removing 200 µL of media (half of the total amount) from the well and adding 200 µL of fresh culture media back to the well (Figure 2J).
7. For the culture with 24 h drug treatment, transfer the insert with ovaries to a new well with 400 µL of fresh culture media after the initial 24 h, using the following steps:
   1. Remove the insert containing the ovaries from the well using a pair of forceps.
   2. Rinse the bottom of the insert briefly by submerging it in a 60 mm dish with fresh culture media.
   3. Gently tap the bottom of the insert on an empty 60 mm dish to remove excess media on the insert. Place the insert into the well with fresh culture media.
      NOTE: Remove excess media from the insert to ensure the ovaries are stabilized on the membrane immediately after changing media or transferring the insert.

3. Fixation and whole-mount immunostaining

1. To fix the cultured ovaries, remove the insert from the plate using a pair of forceps and place it into a 60 mm petri dish containing 5 mL of DPBS. Transfer approximately 100 µL of DPBS next to the ovaries and resuspend them using a 1 mL pipette.
2. Using a 1 mL pipette, transfer the ovaries from the insert into a 1.5 mL Eppendorf tube. Remove DPBS from the tube and add 500 µL of 4% paraformaldehyde. Fix the ovaries at 4°C for 2 h.
   NOTE: The end of the pipette tip may need to be trimmed wider to prevent damaging the tissues if the ovaries are too large to pass through it.
3. Wash the fixed ovaries in 1 mL of PBST₂ (PBS, 0.1%Tween [v/v], 0.5% Triton [v/v]) at least three times, 1 h each on a rotating shaker. It is recommended to leave the tissues in PBST₂ overnight (12-16 h) at 4 °C to enhance antibody penetration.
   NOTE: This can be a stopping point; the tissues may be stored in PBST₂ up to three days before primary antibody incubation.
4. Incubate the ovaries with anti-dead-box helicase 4 (DDX4) in 100 µL of antibody dilution buffer (1% BSA [w/v], 10% Donkey serum [v/v], 0.1 M glycine, 0.1% Tween, and 0.5% Triton in PBS) at a dilution of 1/400 (2.3 µg/mL) at 4 °C for 12-16 h.
5. On the next day, remove the primary antibody and add 1 mL of PBST₂ to the tube. Wash the tissues at least three times, 1 h each on a rotating shaker.
   NOTE: This can be a stopping point; the tissues can remain in PBST₂ up to three days after primary antibody incubation.
6. Dilute the secondary antibody donkey anti-rabbit AF488 at 1/500 dilution (1.5 µg/mL) in PBST₂. Incubate the tissue with at least 100 µL of the diluted secondary antibody at 4 °C for 12-16 h.
7. On the next day, wash the tissues extensively in PBST₂ at least three times, 1 h each on a rotating shaker_. It is recommended to wash the tissues for up to 8 h to reduce the background. Make sure to cover the tubes with foil to protect them from light from this step forward.
8. Remove PBST₂ and add 100 µL of DAPI at a dilution of 1/1000 (1 µg/mL) in PBS to the tube. Incubate the tissues in DAPI for 30 min at room temperature.
9. Wash the ovaries in PBST₂ once for 30 min and mount them on a microscopic slide with an imaging spacer. Transfer the ovaries to the center of the spacer using a 1 mL pipette and remove excess PBST₂. Add about 5 µL of the antifade mounting medium and place a microscope slide on the top of the spacer. Seal the edges of the spacer with clear nail polish for long-term storage.
   NOTE: Before imaging the slides, ensure nail polish is completely dry and that there is no mounting medium on the outside of the cover slip. These reagents may damage the objectives of the microscope. Additionally, avoid using excess mounting media, as the ovaries need to remain stationary during imaging.
10. Image the ovaries with a confocal microscope at 10x magnification with a Z-step size of 10 µm. Adjust settings and use the Z-compensation (excitation gain) to image the entire ovary.This can be done using the following steps:
    1. Adjust the excitation power and detector gain by monitoring signal saturation using the histogram, as it may vary between tissues. Begin with low laser power and gradually increase as needed.
    2. For imaging deeper regions, enable Z-compensation for the gain detector, as signal intensity may decrease with depth. Use a frame averaging of at least 10 and enable bidirectional X-scanning at a speed of 600 Hz.
       NOTE: With these settings, imaging should not exceed 10 min per ovary (~200 µm). Whole-mount immunofluorescent staining of ovaries with anti-DDX4 is typically robust and rarely photobleaches.

4. Follicle quantification

1. To quantify follicles in each ovary, open the images using ImageJ software. Upon opening, ensure that the color mode is selected as colorized and that split channels is checked in the box under split into separate windows. Select the image of interest, then use the image/transform/rotate tool to place the ovary symmetrically in the image.
   NOTE: The images acquired by confocal can be viewed and analyzed in 3D using Imaris software (Figure 3A,B) or ImageJ software¹⁶.
2. Use the analyze/tools/grid tool to divide the ovary into grids (Figure 3C,C').
3. Use the plugins/analyze/cell counter tool to mark all the grids covering the ovary (Figure 3D,D', grids marked by 4). Of these grids, mark every other grid in each row for follicle quantification (Figure 3E,E', grids marked by 8). Grids on the edge of the ovary that have fewer than 10 oocytes are not included for follicle quantification.
   NOTE: Initialize the cell counter and choose the counter types that you consider easy to read.
4. Oocytes can be identified by the presence of a DDX4-positive ring-shaped structure surrounding a nucleus that is positive for DAPI. Primordial follicles have oocytes at a size of approximately 20 µm in diameter surrounded by squamous somatic cells. Developing follicles have oocytes that are greater than 25 µm in diameter and surrounded by one or more layers of cuboidal granulosa cells (Supplementary Figure 1).
5. Count the primordial follicles in each marked grid (as seen in Figure 3E,E' with counter Type 8). Go over the follicles by using optical sections/stacks in the grid, and mark them using the cell counter tool, respectively. Ensure not to double-count primordial follicles on two consecutive optical sections/stacks. Count developing follicles in all the grids marked with counter Type 4.
   NOTE: Use the zoom option to help visualize and count the oocytes through the stacks.
6. Calculate the number of primordial follicles in the entire ovary (nf) by multiplying the average number of primordial follicles per grid (nfgrid) by the number of grids encompassing the ovary (ngrids). nf = nf grid × ngrids.
   NOTE: Statistical analyses were performed, and graphs were generated using Prism 10.2.0 software (GraphPad Software, version 10, Inc., CA). Differences among groups were evaluated using one-way analysis of variance (ANOVA) followed by Tukey's multiple comparison test. Statistical significance was considered at p < 0.05. Data is presented as the mean ± standard error (SE).

Results

In this article, we demonstrate the methods of neonatal ovary culture, whole-mount antibody staining, confocal imaging, and follicle quantification to assess whether the number and position of the ovaries in the insert during culture affect the experimental outcome, specifically, follicle counts in the ovary. We cultured ovaries with the following setups: one, two, or three ovaries per insert for stabilized culture, and three ovaries per insert for floating culture. The ovaries were cultured under two conditions: a contr...

Discussion

In this study, we demonstrated methods for neonatal mouse ovary culture and for whole-mount ovarian follicle quantification. The culture approach can sustain primordial and early developing follicles in vitro. The whole-mount follicle quantification method requires minimal tissue processing, which saves time and preserves tissue integrity, ensuring accurate follicle quantification.

The number of primordial follicles observed in the control group suggests that, while there is a slight reduction...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
24-well cell culture plates	Fisher scientific	09-761-146
Alexa Fluor 488 Donkey Anti-Rabbit	Jackson ImmunoResearch Laboratories	711-546-152
Anti-Anti (100X)	Gibco	15240-062
Bovine Serum Albumin	Sigma-Aldrich	A1470-25G
CD-1 IGS Mouse	Charles River Laboratories	022	Mice
Cell Culture Inserts	Millipore	PICM01250
Comfort Point Insulin Syringe With Needle 1cc 28G x 1/2	Exel	26027
DAPI	Thermo Scientific	62248
DDX4 Rabbit mAb	abcam	ab13840
DMEM/F12 (1:1) (1X)	Gibco	11320-033
Doxorubicine hydrochloride	Sigma-Aldrich	PHR1789
DPBS (1X)	Gibco	14190-144
Fetal Bovine Serum	Sigma-Aldrich	F0926-500ML
Fiji-ImageJ	NIH, USA (https://imagej.net/ij/)	ImageJ 1.52p
Fisherbrand Nutating Mixers - Variable Speed	Fisher scientific	88-861-043	Shaker
Follicle Stimulating Hormone	Sigma-Aldrich	F4021-10UG
Forma Steri-Cycle i160	Thermo Scientific™	i160	CO2 Incubator
Glycine	Fisher scientific	BP381-500
Imaris	Oxford Instruments (https://imaris.oxinst.com/)	Imaris 10.2
Leica TCS SP8	Leica Microsystems	SN 8100001409	Confocal
Normal Donkey Serum	Jackson ImmunoResearch Laboratories	017-000-121
Olympus SZ61	Olympus Microscopes	SZ61	Stereoscope
Paraformaldehyde	Fisher scientific	15714
SecureSeal imaging spacer	Grace Bio-Labs	654002
Triton X-100	Sigma-Aldrich	X100-500ML
TWEEN 20	Sigma-Aldrich	P1379-500ML
VECTASHIELD Antifade Mounting Medium	Vector Laboratories	H-1000