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Method Article

Production and Use of Customizable Agarose Molds for Scaffold-Free Mouse Ovarian Follicle Culture

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DOI:

10.3791/68871

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October 24th, 2025

 ,  ,  ,  ,  ,  ,  , 
1Department of Obstetrics and Gynecology, Feinberg School of Medicine, Northwestern University, 2Department of Biomedical Engineering, Northwestern University, 3Roy J. Carver Department of Biomedical Engineering, Pappajohn Biomedical Institute, University of Iowa

* These authors contributed equally

In This Article

Summary

We describe methods to culture ovarian follicles in a novel scaffold-free agarose mold that complement in vitro gametogenesis.

Abstract

The ovarian follicle is the functional unit of the ovary that produces hormones and gametes needed to sustain female reproductive function and health. The ability to recapitulate folliculogenesis, ovulation, and luteinization in vitro has broad basic, translational, and clinical utility. The most advanced in vitro follicle growth systems maintain the follicle's three-dimensional (3D) architecture, which is crucial for the development of meiotically competent metaphase II oocytes in humans. Recently, a scaffold-free method for in vitro follicle growth of mouse multilayer secondary follicles was developed and validated. For this, custom 3D printed molds were used to micropattern agarose with microwells that accommodate the volumetric expansion of follicles. Follicles grown in this scaffold-free environment showed comparable hormone production and viability relative to well-established alginate-based encapsulated in vitro follicle growth (eIVFG) systems. Importantly, agarose microwells are a scalable method, less technically demanding, and show improved follicle growth and ovulation rates relative to eIVFG. This methodology produces customizable molds that are biocompatible with the oocyte, a cell highly sensitive to material-specific leachates and other environmental contaminants. Further, follicles in this system are cultured in the same focal plane, enabling real-time timelapse imaging and analysis. To increase the accessibility of this new approach, this article details the methods needed to design and 3D-print master molds, create silicone molds for 24- or 96-well plates, and culture isolated multilayer secondary ovarian follicles in the agarose molds. This setup can also be integrated with a cost-effective time-lapse imaging system, enabling morphokinetic analysis. In addition, molds can be paraffin-embedded for downstream histological analyses. Overall, this user-friendly method is a versatile tool for follicle culture and can be customized further to promote the differentiation and maturation of germ cells within the context of the follicle to sustain complete in vitro gametogenesis.

Introduction

The oocyte, or female gamete, contributes the bulk of the cytoplasm and half of the genetic material to the embryo at fertilization, which is essential for the development of the next generation. Oogenesis starts during embryonic development when primordial germ cells enter meiosis and arrest in prophase I. These oocytes reside within primordial follicles and constitute the ovarian reserve, a finite and nonrenewable pool from which they are progressively recruited. Follicles develop in a process known as folliculogenesis, with the goal of ultimately producing a mature gamete capable of fertilization1,2,3,4,5. Follicle development begins with an early, gonadotropin-independent phase where paracrine signaling is essential for the survival and growth of the follicle. As follicles mature to the secondary stage, they become reliant on gonadotropins, marking a shift to endocrine signaling. A key characteristic of secondary follicles is an oocyte surrounded by a minimum of two complete layers of cuboidal granulosa cells, along with the newly developed theca layer. Theca cells form the follicle's outermost layer, providing androgens and structural support6,7. Finally, a fluid-filled antral cavity forms and ruptures in response to a luteinizing hormone (LH) surge during ovulation, releasing a fertilization-competent gamete arrested at Meiosis II2,8,9.

The oocyte achieves developmental competence for fertilization and the ability to support early preimplantation embryo development, referred to as nuclear and cytoplasmic maturation, during oogenesis and folliculogenesis10,11,12. Cytoplasmic maturation is apparent morphologically, as oocytes expand throughout folliculogenesis. As a result of somatic cell proliferation, including theca, granulosa, and cumulus cells, follicles themselves experience significant volumetric expansion. In mice, the diameter of the follicle increases fivefold from early secondary to pre-ovulatory follicles. Bi-directional communication between the oocyte and surrounding somatic cells is essential for oogenesis. In primary follicles, the zona pellucida forms from proteins secreted by the oocyte, and communication between these follicle compartments is partly mediated by transzonal projections, which provide physical contact between these two follicle compartments13,14. Hence, the follicular microenvironment is vital to correctly orchestrate the growth as well as nuclear and cytoplasmic maturation of the oocyte2,5. To better understand the discrete mechanisms that support these processes, there is a need for advanced and physiologically relevant in vitro culture techniques. The ability to culture follicles in vitro has broad applications, including assisted reproductive technologies and fertility preservation15,16,17,18, species conservation19,20, drug development21, and toxicology screening22,23.

In vitro follicle culture systems are also an essential intermediary step for the ability to generate gametes from pluripotent stem cells, also known as in vitro gametogenesis (IVG)24. Researchers have utilized embryonic stem cells, mesenchymal stem cells, and induced pluripotent stem cells to develop gametes in a variety of species, often relying on somatic cells to recreate the native follicle microenvironment25,26,27,28,29,30. While major proof-of-concept milestones have been achieved through these methods, there is still a challenge of low-quality oocytes, reflected in low embryo development and live offspring rates following in vitro fertilization in mice31,32. No live births have been achieved yet in other species, underscoring the need for continued refinement of protocols to fully support oogenesis and folliculogenesis.

Currently, several strategies are utilized for follicle culture. The technically least demanding and highest throughput approaches are scaffold-free methods, where follicles are placed in a plastic dish or membrane insert33,34. Here, granulosa cells adhere to the culture surface, losing contact with the oocyte33,34. The breakdown of the follicular architecture limits the usability and biological relevance of these techniques. However, scaffold-free culture in low attachment plates or inverted drops allows the follicle to maintain structural integrity in several mammalian species35,36,37. Unfortunately, with these techniques, it is difficult to control the interaction between follicles and the amount of mechanical support they receive. Hence, to better recapitulate in vivo physiology, culture methods have been developed where follicles are encapsulated in hydrogels with defined, tunable chemical compositions and/or mechanical properties33,34,38,39,40,41,42,43,44,45,46,47,48,49,50,51,52,53. For example, the alginate-encapsulated in vitro follicle growth system can recapitulate key transcriptomic signatures observed in vivo21.

Despite the success and value of encapsulated culture protocols, there are several disadvantages. They can be technically challenging, laborious, low-throughput, and are not compatible with automatic imaging methods. Further, encapsulated culture methods need multi-step approaches for larger mammalian species due to the higher volumetric expansion of the follicle37. Moreover, encapsulated techniques exert uniform support in all directions, and follicles need to be released from the hydrogel prior to ovulation. This does not occur in vivo, where the follicle interacts with the surrounding ovarian microenvironment, which provides dynamic biomechanical support54. For example, the stiff ovarian cortex maintains quiescence of primordial follicles, while large growing follicles are typically found in the softer medullary compartment55,56. Once a follicle with a competent oocyte has reached its terminal size, follicle wall and ovarian surface epithelium modifications allow for ovulation. Therefore, culture techniques where follicles are allowed to grow in one single direction with little resistance could better mimic in vivo physiology.

In a novel approach, a reproducible, scalable, technically less demanding, and cost-effective culture method for mouse multilayer secondary follicles was developed57. Here, ultra-low adhesion agarose microwells with precisely controllable geometry (shape, dimensions) and spatial arrangement (number, proximity) are manufactured using affordable methods. Follicles in the agarose molds experience mechanical support in all but one direction and are cultured in the same focal plane, enabling timelapse imaging for advanced morphokinetic analysis. Stereolithography (SLA) 3D printing was utilized as an approachable method to manufacture a master mold, followed by molding steps that create leachate-free silicone molds to cast the agarose. Improved follicle growth and ovulation outcomes were achieved using the new system compared to traditional eIVFG, without compromising overall follicle survival, maturation, or luteinization57. This protocol paper describes the steps necessary to customize and manufacture agarose molds, how to seed and culture multilayer secondary follicles, process, and finally perform timelapse analysis (Figure 1).

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Protocol

All animal procedures related to this protocol were approved by the Institutional Animal Care and Use Committee (IACUC) of Northwestern University and performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

1. Adapt master mold design

This section describes the first steps performed during the Design phase (Figure 1), where custom microwells are introduced in a computer-aided design (CAD) of a master mold for 24-well culture plates, called the 24-well master mold base design (Supplemental File 1). The dimensions of the microwells dictate the amount of support and maximal growth size of follicles, while the number of wells determines the maximal number of follicles per agarose mold. The adapted master mold design will then be used in the next section to create biocompatible silicone molds for agarose casting.

  1. Open the 24-well master mold base design .step file using CAD software (Figure 2A).
  2. Select the internal surface area of the object, then navigate to the Solid tab in the Design workspace, and select Create Sketch to insert the desired micromold design (Figure 2Bi).
  3. Introduce the desired xy dimensions and number of the microwells and select finish sketch on the toolbar (Figure 2Bii).
    NOTE: For multilayer secondary follicles, a 500 x 700 µm stadium geometric shape was used previously57. The printing resolution can vary depending on the resins and SLA printers used; allow for a minimal spacing of 400 µm between microwells.
  4. Create the 800 µm deep microwells by using the extrude function (click on Solid | Create | Extrude). Make sure to select the cut operation, creating the microwell cavity (Figure 2Biii).
  5. Select a 0.100 mm radius fillet (click on Design | Solid | Modify | Fillet) for the top of the microwells
    NOTE: To create a round-bottom microwell, use the fillet operation again for the bottom of the culture well and have the radius value equal to the radius of the microwell design. For example, when designing 500 µm x 700 µm wells, select a 0.250 mm radius value (Figure 2Biv).
  6. Export a copy of the new master mold design in .step format (click on File | export).

2. Creating silicone mold containers

NOTE: To prevent the leaching of cytotoxic compounds from the 3D printed parts, two biocompatible silicone molds will be made. This section describes the last steps in the design phase (Figure 1) and will result in the two containers used for silicone molding. Here, three different CAD files will be used, namely the new master mold design created from section 1, and 24-well silicone cast containers 1 and 2 (Figure 2Ci-ii and Supplemental File 2, Supplemental File 3, respectively).

  1. Open the 24-well silicone cast container 1.step file and insert the new master mold design as an external component (navigate to Solid | Insert | Insert Component).
    1. Right-click on the component and select Break Link to remove the linked reference to the new master mold design.
  2. Center the new master mold design with 24-well silicone cast container 1 surface, making sure it faces inwards.
    1. Right-click the new master mold design object in the browser and select the move/copy option.
    2. Select the Free Move operation in the Move Type section and rotate the object 180° on the x-axis if necessary.
    3. Select the Point to Point operation in the Move Type section with the middle of the new master mold design object as the origin point and the center of the 24-well silicone cast container 1 surface as the target point.
    4. Select OK to align the objects.
  3. Select the cut operation with the container as the target body and the master mold as the tool body design (Figure 2Ciii). This creates the container for the first step of silicone molding (Figure 2Civ).
    NOTE: The microwells that were designed in the first section are now micropillars that will be 3D printed as described in section 3. SLA printing is an additive manufacturing method; hence, printing micropillars improves print resolution compared to printing microwells.
  4. Save and export the resulting new 24-well silicone mold 1 as a .stl and .step file.

3. 3D printing of silicone containers

NOTE: This section describes the first steps performed during the Manufacture phase (Figure 1) where the new CAD designs with the desired microwell design features are 3D printed.

  1. Open the .stl files of new 24-well silicone mold 1 and 24-well silicone cast container 2 with 3D print preparation software.
  2. Orient the print with the micropillars facing upwards and use the drill hole function to create a 1 mm wide opening (Figure 2D). Make sure this channel is open to the side of the print.
    NOTE: The bore hole that avoids cupping effects during printing and can be filled after printing, or the excess silicone material can be removed after the first silicone molding.
  3. Print the two container designs at a 25 µm layer thickness (Figure 3A).
    NOTE: Print each model in triplicate to account for print imperfections or damage during processing.
  4. Remove the molds from the printing platform and wash in 95% isopropanol according to the manufacturer's instructions. Spray the micropillar section extensively with 95% isopropanol (Figure 3B). Remove any remaining ethanol with compressed air.
  5. Let the remaining ethanol evaporate for at least 30 min.
    NOTE: At this stage, the 1 mm-wide drill channel can be sealed. This can be done by applying a small drop of unpolymerized resin with a p200 pipette tip on the outside of the print. Capillary forces will fill the channel.
  6. Cure the 3D prints at 60 °C with ultraviolet light for 15 min.
  7. Inspect each print using a stereomicroscope and discard any 3D prints with imperfections (Figure 3C-F). Thoroughly inspect every print visually, ensuring that all micropillars are separated and uniform in size and appearance (Figure 3D). Discard defect master molds with merged micropillars (Figure 3E).
    NOTE: Common issues are damaged micropillars or bridges from remaining unwashed resin (Figure 3Fi,ii).
  8. Cover the outside of the print with parafilm if the drill channel is not filled and store until further use.

4. Silicone mold manufacturing

NOTE: This section describes the final steps of the manufacture phase (Figure 1), where two silicone molding steps will take place. This is necessary to prevent the release of toxic leachates by 3D printed parts.

  1. Prepare the silicone according to the manufacturer's instructions, in summary:
    1. Mix the contents of Part A and B containers thoroughly, place a beaker on a scale and pour into container at a 1A:1B ratio by weight. Make sure to have excess material.
    2. Mix thoroughly for 3 min, making sure to scrape the sides and bottom.
    3. Place the material in a vacuum desiccator and leave for 5 min to eliminate trapped air bubbles. Repeat degassing up to 2x if air bubbles remain.
  2. Pour the mixture with a uniform flow into silicone mold container 1. Ensure that the mixture level is at the same level as the 3D print or slightly below to ensure flatness of the silicone mold (Figure 4Ai,ii).
    1. Remove any trapped air using a p200 pipette tip and use vacuum degassing if necessary.
  3. Cure the silicone at room temperature for at least 5 h and preferably overnight before demolding.
  4. Remove the silicone from the 3D printed mold and visually confirm the integrity of the micromolds using a stereomicroscope (Figure 4Aiii,B). Remove excess material from the drill channel if necessary (Figure 4Aiv). Discard molds with bridging or compromised microwells (Figure 4C).
  5. Place the silicone mold in silicone mold container 2 for the second silicone molding step (Figure 4Av). Silicone container 2 has a 1 mm opening in the side wall that will be covered once the silicone mold is placed.
  6. Lightly spray the silicone mold with embryo-safe mineral oil and remove any excess oil from the microwells. This will act as a release agent to separate both silicone parts after curing.
  7. Repeat the mixing steps described in step 4.1 and pour the mixture into silicone mold container 2. Ensure that the mixture level is below or at the same height as the 3D print to ensure flatness of the silicone mold (Figure 4Avi).
  8. Allow the silicone to cure at room temperature for at least 5 h, preferably overnight, before demolding.
  9. Remove the silicone from the 3D printed mold and separate silicone molds 1 and 2 from each other (Figure 4Avii). Visually confirm the integrity of the micropillars using a stereomicroscope (Figure 4D). Discard the mold if bubbles or excess oil were present in the microwells (Figure 4E).
  10. Wash the silicone mold with 70% ethanol and let air dry for 30 min in a laminar flow hood.
  11. Place in a sterilization pouch and autoclave using a dry cycle to sterilize (Jacket pressure: 20 psi, Chamber temperature: 250 °F, Sterilizing time: 15 min).

5. Preparation of culture media

NOTE: Digestion of ovarian tissue for isolation and culture of follicles, in vitro maturation of oocytes, and luteinization requires six types of media: Dissection Media (DM), Enzymatic Media (EM), Maintenance Media (MM), Growth Media (GM), Maturation Media (IVM), and Luteinization Media (LM). Preparation of these media can be found as previously described by Converse et al.38 with an explanation of the importance of different media components analyzed by Simon et al.33, and their purpose is briefly mentioned in Table 1.

  1. Prepare the media up to 1 week in advance and store at 4 °C.

6. Casting and storage of custom agarose mold and follicle culture

NOTE: This section describes how to cast custom agarose molds for follicle culture from the silicon molds generated in section 4. Agarose molds can be made and stored at 4 °C up to 2 weeks before follicle culture.

  1. Place molds into a sterilization pouch and autoclave using a dry cycle as described in section 4.11. Allow the molds to cool down before use.
  2. Make agarose solution by dissolving sterile 1.5% agarose (w/v) solution in Phosphate Buffered Saline (PBS) in a microwave or hot plate. Utilize a sterile conical tube that is placed in a beaker with water to allow for more even heating of the agarose and to slow down solidification of liquid agarose solution.
  3. Pipette the sterile agarose solution into silicon casts in a laminar flow hood. Pipette slowly and smoothly to minimize bubbles from forming and ensure that the top of the mold is flat and not convex or concave. The top when casting will be the bottom of the mold when culturing and should lie flat. Use approximately 600 µL of agarose for a 24-well plate micromold and 85 µL for a micromold that fits in a 96-well plate.
  4. Let micromolds cool for ~3 min until solidified. Invert silicon cast to expel micromolds onto a 100 mm Petri dish.
  5. Assess the micromolds under a microscope for any deformities, including cracks, merging of microwells, or incomplete borders. Place the micromolds in sterile PBS + 1% Penicillin-Streptomycin and store at 4 °C for up to 2 weeks prior to use.
    NOTE: Make more molds than necessary to account for degradation over time or damage to micromolds when transferring between plates.
  6. Equilibrate micromolds in 750 µL of Maintenance Media 2x for at least 1 h (total of two incubations) on the day of follicle isolation.
  7. Remove all maintenance media, add 750 µL of Growth Media and place in incubator for 1 h.
  8. Replace with fresh and equilibrated Growth Media prior to seeding.

7. Isolation and culture of late secondary follicles

NOTE: This section explains how to isolate and seed follicles in the agarose micromolds for follicle culture. Multilayer secondaries are cultured for 8 days, must have over 80% viability, and produce mature oocytes. Isolation and culture of secondary follicles have been described in significant detail elsewhere38. Isolation and selection of follicles for established alginate encapsulation and for seeding in agarose molds are consistent.

  1. Dissect ovaries from mice and remove from periovarian adipose tissue and bursa. Place ovaries in 2.5 mL of Enzymatic Media and incubate in an orbital shaker at 37 °C for 20 min as previously described57.
  2. Pipette 2 mL of Maintenance Media in an IVF dish and place in an incubator (37 °C, 5% CO2) prior to follicle isolation.
  3. Quench reaction with 250 µL of FBS. Select for late secondary follicles after enzymatic isolation and transfer them to an IVF dish with DM.
  4. Transfer follicles to an IVF dish with pre-equilibrated maintenance media and place in an incubator (37 °C, 5% CO2) for 1 h.
    NOTE: The starting size of late secondary follicles is 150-180 µm with more than two layers of granulosa cells present.
  5. Add 750 µL of fresh GM to agarose micromolds and store in the incubator to prepare for seeding. Add sterile PBS to adjacent wells surrounding agarose micromolds to maintain humidity and minimize evaporation of media throughout culture period.
  6. Seed follicles into agarose molds that have been pre-equilibrated and are submerged in GM. Transfer high-quality multilayer secondary follicles to agarose micromolds under a microscope utilizing a 200 µm stripper tip. Ensure that 10 follicles are transferred to each micromold in a 24-well plate and that each follicle is in a separate but adjacent microwell.
    1. Bend the stripper tip to allow for more precise manipulation and transfer of the follicles into the microwells.
    2. Transfer follicles quickly to minimize temperature and pH changes as MM and GM are CO2 buffered.
    3. Ensure that follicles within a micromold are of a similar starting size once all 10 follicles are transferred.
  7. Place the seeded micromolds back into the incubator for at least 1 h to allow follicles to recover prior to imaging.
  8. Image follicles at 4x and 10x magnification every other day to quantify follicle survival and growth outcomes.
    NOTE: Timelapse imaging can also be set up to perform morphokinetic measurements and are described in more detail in section 9.
  9. Change 50% of culture media every other day.
  10. Snap-freeze spent media with dry ice and then store at -80 °C. Use this conditioned media to assess hormone production through estradiol ELISAs.
    NOTE: If GM is made on the day of seeding (beginning of the culture period), the media can be stored at 4 °C and utilized for the entire culture period.
  11. Remove and snap-freeze all growth media (750 µL) on day 8 of culture.
  12. Replace growth media with IVM media to allow for in vitro ovulation.
    NOTE: Ovulation can also occur under timelapse imaging to assess timing and rupture features.
  13. Remove cumulus-oocyte complexes (COCs) 14-16 h after IVM media has been added and score maturation status.
    1. Define the maturation status is as follows: identify MII by extrusion of the first polar body (expected outcome for healthy oocytes), GVBD/MI by the lack of a polar body or germinal vesicle (delayed maturation), GV by the spherical nucleus visible in the center of the oocyte (lack of meiotic progression) or degenerate-flattened and/or darkened oocyte.
  14. Optional: As spindle parameters of MII oocytes, including chromosomal alignment, shape, and volume, are tightly correlated with oocyte quality and embryo outcomes58,59, use them to assess the success of in vitro oogenesis. To assess spindle morphology, fix and stain MII eggs through whole-mount immunocytochemistry.
    1. Fixation: Fix the eggs in 3.8% PFA with 0.1% Triton X-100 at 37 °C for 20 min. Wash the eggs 3x in blocking buffer (PBS with calcium and magnesium (PBS +/+) with 10% Tween-20 and 0.3% BSA).
    2. Spindle staining:
      1. Incubate the eggs in permeabilization buffer (PBS +/+ with 0.1% Triton X-100 and 0.3% BSA) for 15 min at room temperature
      2. Rinse 2x in blocking buffer.
      3. Incubate the eggs in Alexa Fluor 488-conjugated anti-alpha-tubulin at a concentration of 1 µg/mL on a rocker wrapped in foil for 2 h at room temperature.
      4. Wash the eggs for 3 x 20 min in blocking buffer prior to mounting on microscope slides with DAPI-containing mounting medium.
      5. To observe the oocyte cytoskeleton, include 1:50 rhodamine phalloidin during staining.
    3. Image samples on a confocal microscope using a 63x objective with a z-stack at a width of 1.0 µm throughout the entire spindle.

8. Downstream applications of custom agarose molds

NOTE: Agarose micromolds can be utilized for histological analysis, allowing quantification and localization of different genes or proteins of interest. The micromolds create a follicle microarray that allows for multiple follicles to be stained in parallel. Optical coherence tomography (OCT) can also be performed without having to transfer follicles (Figure 1, Application). Optical Coherence Tomography is an imaging technique that allows for 3D reconstruction of samples and has higher penetration depth and resolution compared to confocal microscopy, allowing for improved imaging of 3D samples such as follicles cultured in vitro60.

  1. For histology applications, make an agarose solution (1.5% w/v) as described in step 6.2.
    1. Remove all media surrounding agarose micromolds in the well and carefully remove media in the micromold cavity utilizing a p200 pipette at the corners of the cavity.
    2. Carefully pipette 100 µL of liquid agarose solution to fill and seal the microwells and cavity without agitating the follicles; ensure that the top is flat. Remove any bubbles with a sterile P10 pipette tip.
    3. Let the agarose solidify at room temperature for 2 min. Ensure under a microscope that the micromold is fully sealed.
    4. Fix the micromolds overnight at 4 °C with gentle rocking in 750 µL of fixative (Modified Davidson's Solution or 3.8% paraformaldehyde).
    5. Aspirate the fixative and wash the sealed micromolds in ascending concentrations of ethanol, starting with 50% and finally 70% EtOH.
    6. Process using an automated tissue processer utilizing standard processing protocols (Table 2) and embed in paraffin wax. When embedding, make note of the orientation of the micromolds. Ensure that there is wax before and after the sample during the embedding process to minimize tissue loss during sectioning.
    7. Serially section embedded samples at 5 µm thickness and place on charged microscope slides.
    8. Perform H&E staining by an automatic stainer and seal with mounting medium prior to imaging.
  2. When performing OCT, place the micromold with follicles in a 50 mm dish containing IVM media.
    1. To follow the procedure used to generate the representative OCT images, image using a custom-built OCT system operating within the visible-light spectral range with an axial resolution of 1.3 µm and a lateral resolution of 9.4 µm61,62.
    2. Scan follicles in a 1.58 x 1,58 mm2 field of view, consisting of 512 A-lines and 512 B-scans, repeated 2x per B-scan and 3x per volume.
    3. Process OCT images using ImageJ to visualize the 3D structure of follicles and quantify volumes.
      1. Upload images into ImageJ as a Z-stack by dragging files into open application.
      2. Visualize the z-stack as a 3D image by going to Plugins|3D|Volume Viewer. Rotate the 3D image to view different angles at different time points.
      3. Navigate to File| Save As |Tiff to save the image to the desired location.

9. Timelapse analysis of follicle culture

NOTE: Follicles grown in the scaffold-free culture system reside in the same focal plane and are compatible with timelapse imaging in the incubator. Timelapse imaging can be started after seeding follicles in step 7.7 (Figure 1, Application).

Make sure that all changes are accurate throughout the timelapse. Hence, pass through the entire stack after every step to ensure there are no discrepancies in the measurements.

  1. Timelapse setup:
    1. Place a Light focuser/concentrator cap on the handheld microscope.
    2. Put the handheld microscope with mount holder in the incubator and connect to a laptop with software installed.
    3. Place and align the culture well under the microscope and adjust the height and focus of the microscope so all follicles are in focus. Using Auto White Balance (AWB), choose optimal lighting conditions using LED Control, turn Auto Exposure (AE) off, and select the optimal exposure time.
      NOTE: Make sure that the alignment of the mold is the same before and after each media change; otherwise, analysis will have to be done separately for every 2 days of culture.
    4. Start timelapse imaging, selecting a duration of 8 days and interval of 30 min. Select photo and Turn off LED when not taking pictures.
  2. Specify which measurements will be recorded in the set measurements dialog box (click Analyze | set measurements) before ImageJ analysis.
    NOTE: For more information on possible measurements incorporated into ImageJ software, see the user guide subsection 30.763.
  3. Place all images in a separate analysis folder to start analysis.
  4. Drag and drop the analysis folder into ImageJ software to open the timelapse as a stack. Ensure that sort names numerically is selected and if image sequences do not fit in the RAM of the computer, select virtual stack.
  5. Use the rectangular selection tool to select an area where all follicles throughout the timelapse are present. For this, use the last image first and scroll through the stack to make sure none of the follicles will be cut off. Then, crop the selection (navigate to Image | Crop or Ctrl+Shift+X, Figure 5Ai,ii).
  6. Set the scale bar for the timelapse stack (Analyze | Set Scale). Utilize the day 0 image made with the imager and measure three identifiable features such as follicles or microwells. Use the same day 0 image from the timelapse stack and define the scale for one of the measurements; use the other two measurements to confirm the accurate scale is set.
  7. Align all images using the SIFT plugin (navigate to Plugins | Registration | Linear Stack Alignment with SIFT, (Figure 5Bi,ii). Use the following standard settings, which can be adjusted at a per case basis: Linear Stack Alignment with SIFT, initial_gaussian_blur = 1.60 steps_per_scale_octave = 3 minimum_image_size = 64 maximum_image_size = 1024 feature_descriptor_size = 4 feature_descriptor_orientation_bins = 8 closest/next_closest_ratio = 0.92 maximal_alignment_error = 25 inlier_ratio = 0.05 expected_transformation=Rigid interpolate.
  8. Convert the stack to 8-bit grayscale (Image | Type | 8 Bit).
  9. Remove the background utilizing the Rolling Ball Background Subtraction (click Process | Subtract Background, Figure 5Ci,ii) using the following settings, which can be adjusted at a per case basis: Rolling ball radius=70.0 pixels; select disable smoothing.
  10. Threshold the image (navigate to Image | Adjust | Threshold) to divide the timelapse images into two classes of pixels, using the following settings: setThreshold(35, 255, "raw"); run("Apply LUT", "stack"); setOption("BlackBackground", true); run("Convert to Mask", "method=Intermodes background=Dark calculate black").
  11. Measure each follicle individually by using the rectangle selection, then select the particle analysis function (click Analyze | Analyze Particles). Use the following settings: Analyze Particles...", "size= 8000-Infinity display exclude include summarize overlay add composite stack (Figure 5Di,ii).
    1. Before saving the results, confirm that the ROI for each image is accurate (Figure 5Ei,ii). Save Summary and Results in the .CSV format.
    2. Repeat this for all other follicles.

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Results

To determine whether follicle survival and growth were supported using this novel technology as described in protocol section 7, follicles from the same mice were isolated and cultured in parallel in agarose micromolds or encapsulated in 0.5% alginate. Representative images of follicles from every other day of culture demonstrated normal development, growth, and formation of an antral cavity throughout the culture period (Figure 6A). Previously published results demonstrated that follicle su...

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Discussion

In vitro gametogenesis and maturation are key technologies for fundamental understanding of oocyte biology, with potential applications in assisted reproductive technology, fertility preservation, and species conservation. The methods to support IVG often rely on co-culture with somatic cells to create reconstructed ovaries30. The importance of these support cells is well-known in vivo, where the follicular somatic microenvironment is critical to healthy oocyte growth and develop...

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Disclosures

The authors have no conflicts of interest to disclose.

Acknowledgements

This work was funded by the National Institutes of Child Health and Human Development (R01HD105752 to F.E.D and T32HD094699 to E.J.Z) and the NIH Common Fund's SenNet program (U54AG075932 and UH3CA268105 to F.E.D.) and the Bill & Melinda Gates Foundation Grant (INV-003385 to F.E.D.). Under the grant conditions of the Bill & Melinda Gates Foundation, a Creative Commons Attribution 4.0 Generic License has already been assigned to the Author Accepted Manuscript version that might arise from this submission.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Materials and Chemicals
 50 mL Polystyrene Centrifuge TubesGlobe Scientific941-11017-CS
1.5 mL Microcentrifuge TubesFisher ScientificMCT-150To snap freeze and store media
100 mm Dish, Non-TreatedFisher Scientific08-757-100D To make and sstore agarose micromolds
100% EthanolMercedes Scientific1200Tissue Processing
1000 µL  Pipette Tips, Low RetentionMIDSCIPR-1000BKCulture tips 
200 µL  Pipette Tips, Low RetentionMIDSCIPR-200BKCulture tips 
24-well plateCorning353047Culture
35 mm dishesFalcon351008To isolate ovaries and incubate in enyzmatic media
95% EthanolMercedes Scientific1210Tissue Processing
AgaroseHoeferGR140Micromolds
BioXtra Mineral Oil, Light Oil, Suitable for mouse embryo cell cultureSigma-AldrichM5310-500MLUsed as a biocompatible demolding agent
Bovine Serum Albumin MP Biomedicals103700Growth Media
Cleaning Dusters, 10 Oz., Pack Of 6Office Depot 110284Compressed air
Clear V4 Resin 1 LFormlabs, IncRS-F2-GPCL-043D printing resin
Corning Reusable Plastic Low Form 100 mL Beaker, Polypropylene, GraduatedCorning1000P-100Beaker for mixing silicone components 
DNAse IQiagen79254Enzymatic Media
Ecoflex 00-45 Near ClearSmooth-On, IncB09M8Y9PTVSilicone material
F-12 + GlutaMAXGibco31765-035Growth Media
Fetal Bovine FetuinSigmaF-3385Growth Media
Fetal Bovine Serum (FBS)Peak SerumPS-FB2Dissection Media, Enzymatic Media,  Maintenance Media & Maturation Media
Fisherbrand Sterilization PouchesFisher Scientific01-812-50 Sterilization Pouches
Follicle stimulating hormone (FSH)EMD Serono, Inc.Gonal-F Rx only Growth Media & Maturation Media
Gibco DPBS, calcium, magnesiumFisher Scientific14-040-182 Dulbecco's phosphate-buffered saline
Human chorionic gonadotropin (hCG)Sigma AldrichC1063-1VLMaturation Media
Insulin-transferrin-selenium (ITS)ThermoFisher41400045Growth Media
Invitrogen Molecular Probes Rhodamine PhalloidinFisher ScientificR415Egg Cytoskeleton Stain
IVF DishesThermo Scientific150260To isolate, select for, and pre-equilibrate follicles
Leibovit'z 15 (L15)Gibco11415-064Dissection Media & Enzymatic Media
LiberaseSigma Aldrich5401119001Enzymatic Media
MEM? + GlutaMAXGibco32561-037Maintenance Media,  Growth Media & Maturation Media
Mouse epidermal growth factor (EGF)BD Biosciences354010Maturation Media
Nalgene Transparent Polycarbonate Classic Design DesiccatorThermo Scientific5311-0250Vacuum Desiccator 
ParaffinFisher Scientific83-30Tissue Processing & Embedding
Penicillin-streptomycinGibco15140-122Dissection Media, Enzymatic Media & Maintenance Media
Stripper tipsOrigioMXL3-200Culture
VECTASHIELD PLUS Antifade Mounting Medium with DAPIVector LaboratoriesH-2000Egg nucleus stain
XyleneMercedes Scientific9840Tissue Processing
α-Tubulin (11H10) Rabbit mAb (Alexa Fluor 488 Conjugate)Cell Signaling Technology5063SEgg Tubulin stain
Equipment
Build Platform 2Formlabs, IncBP-F3-02-01 3D printer component
Dino-Lite Edge AF4915ZTL, USB 2.0Dino-LiteAF4915ZTLImager for timelapse setup
Form 3 Resin Tank V2.1 Formlabs, IncRT-F3-02-013D printer component
Form 3B+ 3D PrinterFormlabs, IncF3B-P-PRINTERFormlabs' stereolithography (SLA) 3D printer
Form cure Formlabs, IncFH-CU-01Curing platform 
Form washFormlabs, IncFH-WA-01Washing platform
Spray bottle w/ pump vaporizerVWR0309-3005To apply demolding agent
Light focuser/concentrator cap for Edge series (excluding T5, T8 models)Dino-LiteN3C-RImager component for timelapse setup
Mount Holder bundle combining the RK-10 rack and RK-10-EX Arm ExtensionDino-LiteRK-10AImager component for timelapse setup
Tissue EmbedderLeicaHistoCore ArcadiaTo embed micromolds
Tissue ProcessorLeicaTP1020To process micromolds
211DS Shaking Incubator (Orbital Shaker)Labnet InternationalI-5211-DSTo incubate ovaries in enzymatic media for release of follicles
Software 
Autodesk FusionAutodesk, IncComputer-aided design (CAD) software. Autodesk offers free educational access through the Autodesk Education program.
DinoCapture 3.0 version 1.1.1.3 Dino-Lite
ImageJNational Institutes of HealthTo analyze OCT images and measure follicles
PreFormFormlabs, IncFree print preparation software designed to work with Formlabs' stereolithography (SLA) printers. 

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Agarose MicromoldsScaffold Free CultureOvarian Follicles3D Printed MoldsTime Lapse ImagingMultilayer Secondary FolliclesSilicone Mold FabricationFolliculogenesis In VitroOptical Coherence Tomography
Production and Use of Customizable Agarose Molds for Scaffold-Free Mouse Ovarian Follicle Culture
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