Isolation of Small Preantral Follicles from the Bovine Ovary Using a Combination of Fragmentation, Homogenization, and Serial Filtration

Summary

Advancing the study of preantral folliculogenesis requires efficient methods of follicle isolation from single ovaries. Presented here is a streamlined, mechanical protocol for follicle isolation from bovine ovaries using a tissue chopper and homogenizer. This method allows collection of a large number of viable preantral follicles from a single ovary.

Abstract

Understanding the full process of mammalian folliculogenesis is crucial for improving assisted reproductive technologies in livestock, humans, and endangered species. Research has been mostly limited to antral and large preantral follicles due to difficulty in the isolation of smaller preantral follicles, especially in large mammals such as bovine species. This work presents an efficient approach to retrieve large numbers of small preantral follicles from a single bovine ovary. The cortex of individual bovine ovaries was sliced into 500 µm cubes using a tissue chopper and homogenized for 6 min at 9,000-11,000 rpm using a 10 mm probe. Large debris was separated from the homogenate using a cheese cloth, followed by serial filtration through 300 µm and 40 µm cell strainers. The contents retained in the 40 µm strainer were rinsed into a search dish, where follicles were identified and collected into a drop of medium. The viability of the collected follicles was tested via trypan blue staining. This method enables the isolation of a large number of viable small preantral follicles from a single bovine ovary in approximately 90 min. Importantly, this method is entirely mechanical and avoids the use of enzymes to dissociate the tissue, which may damage the follicles. The follicles obtained using this protocol can be used for downstream applications such as isolation of RNA for RT-qPCR, immunolocalization of specific proteins, and in vitro culture.

Introduction

Ovarian follicles are the functional units of the ovary, responsible for production of the gamete (oocyte) as well as hormones critical for reproductive function and overall health. Primordial follicles form in the ovary during fetal development or in the neonatal period depending on the species¹, and they constitute a female's ovarian reserve. Follicular growth begins with the activation of primordial follicles that leave the resting pool and enter the growing phase. Preantral folliculogenesis, encompassing all follicle stages before antrum development, is a highly dynamic process that requires synchronous morphological and metabolic changes in the oocyte and the surrounding granulosa cells, driven by tight communication between these two cell types^2,3. Preantral follicles constitute the majority of the follicular units found in the ovary at any given time⁴. Development through the preantral stages of folliculogenesis is estimated to be several weeks longer than antral development^5,6, and this time is necessary for the oocyte and somatic cells to acquire sufficient maturity to enter the final stage of development (i.e., the antral stage), and prepare for ovulation, fertilization, and embryonic development^7,8,9.

Much of the current knowledge about ovarian preantral folliculogenesis comes from mouse models^10,11,12,13, due in part to the ease in recovering a large number of these follicles from a smaller and less fibrous ovary. Although reports of isolation of large numbers of preantral follicles from bovine ovaries date back approximately 30 years¹⁴, a more complete understanding about the processes regulating the development of these early-stage follicles has remained unrealized, largely due to the lack of optimized, efficient, and repeatable methods to retrieve sufficient numbers of viable preantral follicles, particularly at early stages of development. With the increasing interest in preserving the ovarian reserve for future use in assisted reproduction in humans, cows become an attractive model due to their more similar ovarian structure¹⁵. However, the bovine ovary is markedly richer in collagen compared to the mouse ovary¹⁶, making mechanical isolation using methods described for the mouse very inefficient. Efforts to expand fertility preservation techniques include complete in vitro growth of preantral follicles to the antral stage, followed by in vitro maturation (IVM) of the enclosed oocytes, in vitro fertilization (IVF), and embryo production and transfer¹⁷. Thus far, this entire process has only been achieved in mice¹⁸. In cattle, the progress toward follicle growth in vitro is limited to a few reports with variable follicle stages at the start of culture, as well as variable length of culture between protocols^17,19.

The methods described in the literature for the harvest of preantral follicles from the bovine ovary have mostly used mechanical and enzymatic techniques, either isolated or in combination^2,14,17,20. The first report of a protocol for bovine preantral follicle isolation used a tissue homogenizer and serial filtration to process whole ovaries²⁰. This study was followed by reports combining mechanical and enzymatic procedures that utilized collagenase¹⁴. A recurrent theme when utilizing collagenase to digest the ovarian tissue is the potential risk for damage of the follicular basement membrane, which may compromise follicle viability^14,21,22,23. Therefore, different combinations of mechanical methods have been employed, such as the use of a tissue chopper and repeated pipetting or a tissue chopper combined with homogenization^20,24,25,26. Another mechanical technique that has been described utilizes needles to dissect preantral follicles directly from the ovarian tissue, which is especially useful for isolating larger (>200 µm) secondary follicles. However, this process is time-consuming, inefficient for isolating smaller preantral follicles, and is skillset-dependent when attempted in bovine ovaries^19,27,28.

Taking advantage of the different techniques described in the literature, this protocol aimed to optimize the isolation of preantral follicles from single bovine ovaries in a simple, consistent, and efficient manner that avoids incubation in enzymatic solutions. Improving the methods to isolate preantral follicles will provide an opportunity to enhance the understanding of this stage of folliculogenesis and enable the development of effective culture systems to develop preantral follicles to the antral stage. The detailed procedures described herein for the isolation of preantral follicles from a large mammal such as the bovine species will be vital for researchers aiming to study early folliculogenesis in a non-murine species that is translatable to humans.

Protocol

Bovine (Bos taurus) ovaries were sourced from a local abattoir and transported to the laboratory within 6 h of collection. Due to the large number of animals processed in the facility, the age, breed, and stage of the estrus cycle of the animals are unknown. Because no live animals were used in these experiments, an approved animal care and use protocol was not required.

1. Preparation of equipment and reagents

1. Cover a 2 ft wide section of a lab bench with bench paper.
2. Obtain a scalpel handle, sterile scalpel blade, hemostat, pair of dissection forceps, 20 mL luer-lock syringe, 18 G needle, two 200 mL beakers, 500 mL Erlenmeyer flask, 104 mm diameter plastic funnel, plastic cutting board, one 22 cm² layer of cheesecloth (the cheesecloth can be sterilized by autoclaving prior to use) per ovary being processed, a 300 µm cell strainer, and a 40 µm cell strainer (see Table of Materials).
3. Transfer all equipment onto the bench paper.
4. Use the hemostat to slot the scalpel blade onto the scalpel handle. Align the angled base of the blade to the angled indicator on the handle, then slide the blade into the groove of the handle.
5. Place the funnel into the Erlenmeyer flask and cover the funnel opening with the cheesecloth.
6. Place one 50 mL conical tube per ovary to be processed in a water or bead bath set to 38.5 °C.
7. Place one 100 mm x 15 mm square Petri dish per ovary being processed on a slide warmer set to 38.5 °C.
8. Add 10 mL of Penicillin-Streptomycin (PenStrep; 10,000 U/mL penicillin and 10,000 µg/mL streptomycin) to 1 L of 1x phosphate buffered saline (PBS). Warm the PBS + PenStrep in a water or bead bath set to 38.5 °C at least 2 h prior to ovary processing.
   ​NOTE: PBS + PenStrep solution is imperative for washing ovaries when isolated follicles will be cultured, and it is still recommended for any downstream experiments to mitigate microbial contamination.
9. For collecting processed ovary filtrate, use Follicle Wash Medium (FWM) consisting of TCM199 with Hank's Salts (see Table of Materials) containing 3 mg/mL bovine serum albumin (BSA), 25 mM HEPES buffer, 100 UI penicillin/100 µg/mL streptomycin, 1 mM sodium pyruvate (NaPyr), and 100 nM non-essential amino acids (NEAA).
   1. Transfer sterile TCM199, a 250 mL bottle, and a 100 mL graduated cylinder to a biosafety cabinet (BSC). Transfer 194 mL of TCM199 to the bottle.
   2. Remove the beaker of TCM199 from the BSC and bring to a stir plate. Add 600 mg of BSA, 1.19 g of HEPES buffer, and an autoclaved stir bar to the bottle and stir until dissolved.
   3. Once the BSA and HEPES buffer have completely dissolved, add 1 N sodium hydroxide (NaOH) to the medium until it reaches a pH of 7.6-7.8, as measured by a pH meter.
   4. Wipe the bottle of medium, a vacuum filtration device, four 50 mL conical tubes, and the bottles of PenStrep, NaPyr, and NEAA with 70% ethanol before transferring to the BSC.
   5. Add 2 mL each of PenStrep (10,000 U/mL penicillin and 10,000 µg/mL streptomycin), 100 mM NaPyr, and 100x NEAA to the bottle of TCM199 + 3 mg/mL BSA + 25 mM HEPES. Sterile-filter the final medium and aliquot into the 50 mL conical tubes. Store the medium at 4 °C for up to 2 weeks.
   6. Warm one 50 mL conical tube of medium per two ovaries in a bead bath set to 38.5 °C at least 1 h prior to ovary processing.

2. Tissue chopper setup

1. Ensure the tissue chopper (see Table of Materials) is plugged in and turned on.
2. Set the slice thickness to 500 µm, the blade force control knob to 20°, and the speed control knob to 90° according to the manufacturer's specifications.
3. Insert a 60 mm plastic Petri dish into the plate holder and insert the plate holder onto its stage.
4. Raise the chopping arm as high as it will go by turning the manual operating knob clockwise.
5. Using forceps, place a double-edged razor blade (see Table of Materials) onto the screw inserted into the chopping arm. Place the blade clasp over the blade and secure with the washer and nut. Leave the nut a quarter turn loose.
6. Rotate the manual operating knob until the chopping arm snaps the blade flat onto the Petri dish. Tighten the nut the rest of the way with the nutdriver.
7. Raise the chopping arm as high as it will go using the manual operating knob. Move the table release knob all the way to the left until it notches into place.

3. Ovary preparation

1. Transfer ovaries in the laboratory to warm (38.5 °C) sterile PBS + PenStrep.
   NOTE: It is recommended to process the ovaries for follicle isolation as soon as it is feasible after removal from the animal. In this protocol, ovaries were processed within 6 h of harvesting. Ovaries were transported from the abattoir to the laboratory in thermoses containing sterile 0.9% saline solution at approximately 38.5 °C.
2. If possible, select small (≤ 4 cm x 3 cm x 3 cm) ovaries containing small (3-5 mm) antral follicles, no large (≥8 mm) antral follicles, and no prominent corpus luteum (Figure 1). These criteria are recommended to ensure that a minimal amount of non-follicle debris, such as stromal cells and extracellular matrix, is included in the resulting square dish containing isolated follicles.
   NOTE: Antral follicles can be identified as spherical, fluid-filled vesicular structures on the surface of the ovary. Corpora lutea can be identified as red, orange, or yellow stiff structures protruding from the surface of the ovary.
3. Use scissors to remove any excess connective tissue and fat from the ovaries.
4. Wash the ovaries for 30 s in 70% ethanol in a beaker.
5. Wash the ovaries 3x for 2 min each in beakers of warm (38.5 °C) PBS + PenStrep, using fresh PBS + PenStrep for each wash.
6. Keep the ovaries in warm (38.5 °C) PBS + PenStrep until ready for processing.
   NOTE: Distance between the laboratory and ovary source can be variable. Therefore, it is important to complete the protocol in a timely manner to ensure maintenance of follicular viability.

Figure 1: Anatomy of the bovine ovary. The bovine ovary consists of two main regions enclosed in an epithelial layer. The cortex, comprised of the tissue to the left of the dashed line, contains ovarian follicles from the primordial stage to the antral stage. Preantral follicles are too small to see with the naked eye; antral follicles are marked with asterisks. The medulla, comprised of the tissue to the right of the dashed line, contains blood vessels, lymphatic vessels, and nerves. Please click here to view a larger version of this figure.

4. Chop procedure

NOTE: Only process one ovary at a time. Process ovaries quickly to avoid temperature decreases, which may affect follicle viability.

1. Transfer one ovary to the cutting board on the bench paper (Figure 2A) and prepare the tissue chopper (Figure 2B).
2. Using forceps and a scalpel, cut the ovary in half and remove the medulla from each half, leaving only the cortex at a thickness of approximately 1 mm as shown in Figure 2C.
   1. Cut the ovary in half longitudinally from one ligament attachment site to the opposite attachment site.
   2. Keep one half of the ovary on the cutting board to be processed and place the other half of the ovary back into warm (38.5 °C) PBS + PenStrep.
   3. With the exposed medulla facing upward, slice along the curvature of the ovary approximately 2 mm away from the surface of the ovary without cutting through the cortex.
   4. Use the slice along the curvature of the ovary as a guide to deepen the slice, still following the curvature of the ovary to separate the cortex from the medulla.
   5. Dissect and discard any corpora lutea from the ovary by cutting along the border of the corpus luteum.
   6. Flip the ovary half over so that the epithelium faces upward and use the scalpel to finish cutting the medulla away from the cortex. Trim away any remaining white connective tissue around the edge of the ovary piece that was connected to the ligaments.
   7. Once the majority of the medulla is removed, use the scalpel to cut the cortex to approximately 1 mm thickness. Manipulate the scalpel with small, back-and-forth motions to shave away the remainder of the medulla.
      NOTE: The medulla is the inner portion of the ovary containing large blood vessels. The cortex is the outer portion of the ovary, lying directly underneath the outermost surface epithelium. The cortex is approximately 1 mm thick in the bovine ovary, and thus cutting the ovary to a thickness of 1 mm will remove the medulla.
3. Cut the cortex into pieces no larger than 2.5 cm x 2.5 cm. Keep the cortex pieces in warm (38.5 °C) PBS + PenStrep until ready to chop.
4. Fill a beaker with at least 50 mL of warm (38.5 °C) PBS + PenStrep and obtain a plastic transfer pipette.
5. Transfer a single piece of cortex to the Petri dish on the tissue chopper and wet the tissue with three or four drops of warm (38.5 °C) PBS + PenStrep.
6. Hold the piece of tissue steady with a pair of forceps and press the reset button once to start the tissue chopper. Stabilize the Petri dish with one hand while continuing to stabilize the tissue with the forceps. Move the forceps left along the tissue as needed to avoid the blade hitting the forceps. The resulting strips will be approximately 500 µm in length.
7. Once the entire piece of cortex has been cut into strips, use the blade holder knob to lift the blade off the Petri dish and the forceps to remove any tissue from the blade.
8. Rotate the plate holder 90°.
9. Press the reset button once. Stabilize the Petri dish with one hand while using the forceps to push the tissue strips into the path of the blade.
10. Pass the blade entirely through the tissue strips. Use the blade holder knob to lift the blade off the Petri dish and the forceps to remove any tissue from the blade.
11. Use the transfer pipette and warm (38.5 °C) PBS + PenStrep to wash the chopped tissue (final size of tissue: 500 µm x 500 µm x 1 mm cubes) into a pre-warmed (38.5 °C) 50 mL conical tube. Return the conical tube to the water or bead bath to keep the chopped tissue warm (38.5 °C).
12. Use the nutdriver to remove the nut from the chopping arm, and remove the washer and blade clasp. Using forceps, remove the blade from the chopping arm, flip it over so that the unused edge is facing the Petri dish, and place it back onto the chopping arm. Replace the blade clasp, washer, and nut, and reset the table release knob as described in steps 2.5-2.7.
13. Repeat steps 4.5-4.12 for all remaining pieces of cortex from the ovary, replacing blades with new ones after each cutting edge has been used.
14. Dispose of all used blades in a hard-walled, plastic sharps container.

5. Homogenization procedure

1. Ensure the homogenizer unit (see Table of Materials) is plugged in and the speed is set to the second bar (9,000-11,000 rpm). Insert the 10 mm generator probe into the unit according to the manufacturer's specifications.
2. Set a timer for 1 min and insert the probe into the 50 mL conical tube containing the chopped cortex tissue from one ovary (step 4.11) and enough PBS + PenStrep to fill the tube to the 25 mL line. The depth to which the probe is inserted must be 1/3 of the liquid's height measured from the bottom of the chamber. Position the probe slightly off center to minimize vortexing.
3. Start the timer and turn on the homogenizer. Ensure the bottom of the probe does not touch the tube and hold the tube still while the homogenizer is turned on.
4. After 1 min of homogenization, remove the probe from the tube. Using forceps, remove any connective tissue clogging the venting holes and the space between the rotor knife and rotor tube. If any pieces of cortex are stuck in the probe, remove them with forceps and place them back into the tube.
5. Repeat steps 5.2-5.4 an additional 5x for a total of 6 min of homogenization.
6. Place the tube with homogenized tissue into the water or bead bath to keep the tissue warm (38.5 °C). After processing the last ovary, immediately disassemble, clean, and dry the generator probe according to the manufacturer's specifications.

6. Filtration procedure

1. Pour the dispersed tissue into the cheesecloth-covered funnel inserted into the Erlenmeyer flask. Rinse the contents of the tube into the funnel using warm (38.5 °C) PBS + PenStrep until no tissue fragments remain in the tube.
2. Force the tissue fragments to pass through the holes of the cloth by twisting the cheesecloth around the tissue fragments and squeezing until all excess fluid and tissue are removed from the cheesecloth.
3. Reopen the cheesecloth over the funnel, rinse the cheesecloth with PBS + PenStrep using a transfer pipette, and again squeeze any residual tissue fragments through the cloth.
4. Use a hemostat to hold the 300 µm cell strainer over a 200 mL beaker. Pour the filtrate in the Erlenmeyer flask through the cell strainer. Rinse the contents of the flask into the cell strainer using warm (38.5 °C) PBS + PenStrep until no tissue fragments remain.
   1. If the cell strainer becomes clogged with tissue, gently tap the cell strainer against the beaker to ensure all liquid has filtered into the beaker, and then turn the cell strainer upside down and tap out the large tissue debris onto the bench paper. Return the cell strainer over the beaker and continue pouring the filtrate through it. Repeat as necessary until all filtrate from the Erlenmeyer flask has been filtered through.
5. Use a hemostat to hold the 40 µm cell strainer over a second 200 mL beaker. Pour the filtrate in the first 200 mL beaker through the cell strainer. Rinse the contents of the beaker into the cell strainer using warm (38.5 °C) PBS + PenStrep until no tissue fragments remain. Do not discard the contents of the 40 µm cell strainer.
6. Fit the 18 G needle to the 20 mL syringe. Fill the syringe with FWM. Turn the 40 µm cell strainer upside down over a square Petri dish and use the syringe to wash out the contents of the cell strainer into the dish. Refill the syringe and rinse the cell strainer as needed until no tissue fragments remain.
   NOTE: Typically, 25 mL of FWM is sufficient to fully rinse out the contents of the 40 µm cell strainer.

Figure 2: Workspace setup for ovary processing and protocol workflow. (A) Bench setup for cutting ovaries prior to chopping and for filtering the ovary homogenate. (B) Tissue chopper and homogenizer set up, with Styrofoam support to reduce vibrations of the homogenizer stage. (C) Schematic illustrating the workflow for the processing of one whole ovary. Ovaries are trimmed of excess connective tissue and then cut in half, and the medulla is removed until a ~1 mm thick slice of cortex remains. The cortex is cut into 2.5 cm x 2.5 cm pieces and chopped in a tissue chopper set to a cut interval of 500 µm. The pieces are then homogenized, and the homogenate is filtered through cheesecloth followed by filtration through 300 µm and 40 µm cell strainers. The contents of the 40 µm cell strainer are rinsed into a square Petri dish, which is searched for follicles using a stereomicroscope. Created with BioRender.com. Please click here to view a larger version of this figure.

7. Searching and collecting follicles

1. Transfer the square Petri dish (step 6.6) to a stereoscope with a warmed stage set to 38.5 °C. The stereoscope magnification should be set between 1.25x and 3.2x depending on the searcher's preference.
2. Pipette 10 µL drops of FWM into a 60 mm Petri dish and cover the drops with mineral oil to prevent drying out. Place the Petri dish with media drops onto a warming plate set to 38.5 °C.
   NOTE: A 4-well plate can be used to collect follicles. Add 500 µL of wash media to one or two wells. Place on the warming plate set to 38.5 °C.
3. Obtain a micropipette plunger and tip.
   NOTE: A glass 1-5 µL micropipette (see Table of Materials) is recommended because the follicles are less likely to adhere to the glass pipette and be lost when transferred between solutions. It is also a small enough instrument to allow easier and more precise micromanipulations of the follicles.
4. Identify follicles from the square Petri dish and transfer to media (FWM) drops using the micropipette. Many follicles are likely to be embedded in tissue debris and may be retrievable using one of two methods as described below.
   NOTE: Follicles are oblong, rather than perfect spheres, and typically have an oocyte, presenting as a solid white circle in darker contrasts, toward the center of the follicle (Figure 3A-C). Take care to avoid confusing follicles with denuded oocytes. Oocytes tend to be perfect spheres and are surrounded by a thick, clear membrane (the zona pellucida). An inverted microscope with a magnification of 10x (or greater) can be used for closer examination of follicles (Figure 3D).
   1. Carefully separate the follicles from debris using the tip of the micropipette or fine (27 G) needles.
   2. Alternatively, use a glass Pasteur pipette with a rubber bulb to take up and squirt out the debris in the dish multiple times to dislodge follicles from the debris.
5. Work quickly, taking no longer than 30 min, to search the Petri dish to help preserve the follicle viability.
6. Place a maximum of only five follicles per 10 µL drop, as a higher density can increase the likelihood of follicles adhering together.

8. Trypan blue exclusion viability test

NOTE: Use the lid of a Petri dish or a 4-well plate for all the following steps, as the follicles stick less to the plastic of the lid than they do to the plastic of the actual dish.

1. Prepare PBS + 0.2% polyvinylpyrrolidone (PVP) by dissolving 100 mg PVP in 50 mL of PBS.
   NOTE: PVP is used here to reduce the likelihood of follicles attaching to the dish.
2. Use the micropipette to transfer all follicles (average of 40) from the media drops into a 50 µL drop of PBS + 0.2% PVP.
3. Wash the follicles 2x by transferring them sequentially to fresh 50 µL drops of PBS + 0.2% PVP.
4. Transfer the follicles to a 285 µL drop of PBS + 0.2% PVP.
5. Add 15 µL of trypan blue to the 285 µL drop of PBS + 0.2% PVP (final concentration of 0.05% trypan blue) and carefully mix the drop using a 200 µL pipette tip set to 100 µL.
   NOTE: If using the 4-well plate for the trypan viability test, add 475 µL of PBS + 0.2% PVP and 25 µL of trypan blue to one well.
6. Incubate the follicles for 1 min in the trypan blue drop, and then transfer the follicles to a 50 µL drop (or 500 µL well) of PBS + 0.2% PVP.
7. Wash the follicles 3x according to step 8.3 with fresh 50 µL drops (or 500 µL per well) of PBS + 0.2% PVP.
8. Discard any follicles that still appear blue after three washes in PBS + 0.2% PVP, as these are non-viable. Any follicles that do not retain blue coloration after three washes are viable and may be used for immunofluorescence, culture, or other procedures (Figure 3E). Snap-freeze the follicles in liquid nitrogen and store at -80 °C until further use if required.
9. Perform RT-qPCR analysis and immunofluorescence staining of the follicles as described in steps 9 and 10.

Figure 3: Isolated follicles and trypan blue exclusion test. (A-C) Isolated follicles were imaged through a stereomicroscope at several magnifications. (A) Isolated follicles among debris within the initial search dish. Individual follicles are circled in red. Scale bar = 2,000 µm. (B) Isolated follicles and debris within a droplet of follicle wash medium covered with mineral oil. Scale bar = 1,000 µm. (C) Isolated follicles without debris at a higher magnification. Scale bar = 1,000 µm. (D) Isolated follicles imaged using an inverted brightfield microscope. Scale bar = 100 µm. (E) Representative images of viable (unstained) and non-viable (blue stain) follicles imaged using an inverted brightfield microscope and a 20x objective. Scale bar = 100 µm. Please click here to view a larger version of this figure.

9. RT-qPCR analysis

1. Isolate RNA from viable follicles (from step 8.8) using an RNA isolation reagent (see Table of Materials). Purify the RNA and treat with DNase using a commercially available cleanup kit (see Table of Materials) according to the manufacturer's instructions.
2. Elute the RNA with 14 µL of RNase-free water and quantify using a spectrophotometer. The RNA can be stored at -80 °C until cDNA synthesis.
3. Perform cDNA synthesis from equal amounts of RNA extracted from primary and early secondary follicles, using a commercially available cDNA synthesis kit (see Table of Materials) according to the manufacturer's instructions. Incubate the reaction mixture for 5 min at 25 °C followed by 60 min at 42 °C, then terminate the reaction by heating at 70 °C for 5 min.
4. Perform RT-qPCR with the synthesized cDNA (5 ng per reaction) and primers (Table 1) using a commercially available reaction mix (see Table of Materials). Use thermal cycling conditions: 30 s at 95 °C for polymerase activation, followed by 40 cycles of amplification, where each cycle included 15 s at 95 °C for denaturation and 30 s at 60 °C for annealing/extension. Analyze the RT-qPCR by quantifying cycle threshold (Ct) values and/or view PCR products using agarose gel electrophoresis.
   NOTE: Transcript expression of the granulosa cell marker FSHR and germ cell marker DAZL were evaluated in this study. Reference genes were H2A and ACTB.
5. Run a melt curve analysis by increasing the temperature from 65 °C in 0.5 °C increments every 5 s until it reaches 95 °C.

10. Immunofluorescence analysis

1. Fix viable follicles (from step 8.8) for 15 min in a 100 µL drop of 4% (v/v) paraformaldehyde (PFA) at room temperature (RT), followed by washing 3x in 100 µL drops of PBS + 0.1% BSA + 0.1% Tween 20.
2. Block the follicles for 1 h at RT in a blocking buffer consisting of 1x PBS + 5% (v/v) normal donkey serum (NDS). After blocking, incubate the follicles overnight at 4 °C in a 100 µL drop of 4 µg/mL rabbit anti-human CX37 antibody or 4 µg/mL rabbit isotype IgG (negative control) diluted in blocking buffer.
3. Wash the follicles 3x in 100 µL drops of PBS + 0.1% BSA + 0.1% Tween 20, and then incubate them for 1 h at RT in the dark in a 100 µL drop of 2 µg/mL donkey-anti-rabbit AlexaFluor 488 secondary antibody diluted in blocking buffer.
4. Incubate the follicles for 5 min at RT in the dark in a 100 µL drop of 1 µg/mL Hoechst 33342 diluted in blocking buffer to label DNA.
5. Transfer the follicles to a 5 µL drop of mounting media (see Table of Materials) on a glass microscope slide and cover with a cover slip. Leave the slides to cure at RT overnight, followed by sealing with nail polish. Store them at 4 °C until imaging.
6. Image all slides within 48 h of cover slipping. Perform imaging using an inverted epifluorescence microscope (see Table of Materials) under DAPI (excitation 380 nm and emission 450 nm) and FITC (excitation 470 nm and emission 525 nm) filters.
7. Fix the exposure time for both channels. Adjust the FITC (CX37) exposure time based on the rabbit isotype negative control. Use a 20x objective and the DAPI channel set to 50 ms exposure time to identify rabbit isotype-labeled follicles.
8. Image these follicles under the FITC channel, and decrease the exposure time until all background green signal is abolished. Note this exposure time.
9. Image all CX37 antibody-labeled follicles using the exposure time set for the isotype FITC channel and 50 ms exposure time for the DAPI channel.
10. Process the signal intensity, as measured by mean gray area after thresholding, using a computer image processing program²⁹ (see Table of Materials).
    1. Adjust the tiff file of the DAPI image for each follicle such that the entire follicle is outlined. Use the Analyze Particles function of the program to select the entire follicle as a region of interest (ROI).
    2. Open the tiff file of the FITC image for the corresponding follicle and overlay the ROI generated from the DAPI image on top of the FITC image. Use the Measure function of the program to quantify the mean gray area of the FITC image, which represents signal intensity.

Results

Overview and critical steps
Using this protocol, small bovine preantral follicles can be reliably isolated from single ovaries in experimentally relevant numbers. From a total of 30 replicates, an average of 41 follicles were obtained per replicate, with a range of 11 to 135 follicles (Figure 4A). In 14 replicates, the follicles were characterized for stage of development as previously described²⁶ by measuring the follicle diameter using a 1
...

Discussion

The present protocol details a reproducible method to retrieve early stage preantral follicles, specifically at primary and early secondary stages, from the bovine ovary. This protocol builds on previous reports^{20,25,30,34,35,36} and provides optimizations that result in the isolation of a meaningful number of follicles from a...

Materials

Name	Company	Catalog Number	Comments
5-3/4" Soda Lime Disposable Glass Pasteur Pipette	Duran Wheaton Kimble	63A54	Pasteur pipette that can be used to dislodge follicles from debris while searching within the petri dish
16% Paraformaldehyde	Electron Microscopy Sciences	15710	Diluted to 4%; fixation of follicles for immunostaining
20 mL Luer-lock Syringe	Fisher Scientific	Z116882-100EA	Syringe used with the 18 G needle to dislodge follicles from the 40 μm cell strainer
#21 Sterile Scalpel Blade	Fisher Scientific	50-365-023	Used to cut the ovaries and remove the medula
40 μm Cell Strainer	Fisher Scientific	22-363-547	Used to filter the filtrate from the 300 μm cell strainer
104 mm Plastic Funnel	Fisher Scientific	10-348C	Size can vary, but ensure the cheese cloth is cut appropriately and that the ovarian homogenate will not spill over
300 μm Cell Strainer	pluriSelect	43-50300-03	Used to filter the filtrate from the cheese cloth
500 mL Erlenmeyer Flask	Fisher Scientific	FB500500	Funnel and flask used to catch filtrate from the cheese cloth
Air-Tite Sterile Needles 18 G	Thermo Fisher Scientific	14-817-151	18 G offers enough pressure to dislodge follicles from the 40 μm cell strainer
Air-Tite Sterile Needles 27 G 13 mm	Fisher Scientific	14-817-171	Needles that can be used to manipulate any debris in which follicles are stuck
BD Hoechst 33342 Solution	Fisher Scientific	BDB561908	Fluorescent DNA stain
Bovine Serum Albumin (BSA)	Sigma-Aldrich	A7030-100G	Component of follicle wash media
Cheese Cloth	Electron Microscopy Sciences	71748-00	First filtering step of the ovarian homogenate meant to remove large tissue debris
Classic Double Edge Safety Razor Blades	Wilkinson Sword	N/A	Razor blades that fit the best in the McIlwain Tissue Chopper and do not dull quickly
Donkey-Anti-Rabbit Secondary Antibody, Alexa Fluor 488	Fisher Scientific	A-21206	Secondary antibody for immunostaining
Eisco Latex Pipette Bulbs	Fisher Scientific	S29388	Rubber bulb to use with Pasteur pipettes
HEPES Buffer	Sigma-Aldrich	H3375	Component of follicle wash media
Homogenizer	VWR	10032-336	Homogenize the ovarian tissue to release follicles
ImageJ/Fiji	NIH	v2.3.1	Software used for analysis of fluorescence-immunolocalization
McIlwain Tissue Chopper	Ted Pella	10184	Used to cut ovarian tissue small enough for homogenization
Microscope - Stereoscope	Olympus	SZX2-ILLT	Dissection microscope used for searching and harvesting follicles from the filtrate
Microscope - Inverted	Nikon	Diaphot 300	Inverted microscope used for high magnification brightfield visualization of isolated follicles
Microscope - Inverted	ECHO	Revolve R4	Inverted microscope used for high magnification brightfield and epifluorescence visualization of isolated follicles
Mineral Oil	Sigma-Aldrich	M8410-1L	Oil to cover the drops of follicle wash medium to prevent evaporation during searching
Non-essential Amino Acids (NEAA)	Gibco	11140-050	Component of follicle wash medium
Normal Donkey Serum	Jackson ImmunoResearch	017-000-001	Reagent for immunostaining blocking buffer
Nunc 4-well Dishes for IVF	Thermo Fisher Scientific	144444	4-well dishes for follicle isolation and washing
Penicillin-Streptomycin Solution 100x	Gibco	15-140-122	Component of follicle wash medium
Petri Dish 60 mm OD x 13.7 mm	Ted Pella	10184-04	Petri dish that fits the best in the McIlwain Tissue Chopper
Phosphate Buffered Saline (PBS)	Fisher Scientific	BP665-1	Washing buffer for ovaries and follicles
Plastic Cutting Board	Fisher Scientific	09-002-24A	Cutting board of sufficient size to safely cut ovaries
Polyvinylpyrrolidone (PVP)	Fisher Scientific	BP431-100	Addition of PVP (0.1% w/v) to PBS prevents follicles from sticking to the plate or each other
ProLong Gold Antifade Mountant	Thermo Fisher Scientific	P36930	Mounting medium for fluorescently labeled cells or tissue
Qiagen RNeasy Micro Kit	Qiagen	74004	RNA column clean-up kit
R	The R Foundation	v4.1.2	Statistical analysis software
Rabbit-Anti-Human Cx37/GJA4 Polyclonal Antibody	Abcam	ab181701	Cx37 primary antibody for immunostaining
RevertAid RT Reverse Transcription Kit	Thermo Fisher Scientific	K1691	cDNA synthesis kit
Rstudio	RStudio, PBC	v2021.09.2	Statistical analysis software
Sodium Hydroxide Solution (1N/Certified)	Fisher Scientific	SS266-1	Used to increase media pH to 7.6-7.8
Sodium Pyruvate (NaPyr)	Gibco	11360-070	Component of follicle wash medium
Square Petri Dish 100 mm x 15 mm	Thermo Fisher Scientific	60872-310	Gridded petri dishes allow for more efficient identification of follicles
SsoAdvanced Universal SYBR Green Supermix	BioRad	1725271	Mastermix for PCR reaction
Steritop Threaded Bottle Top Filter	Sigma-Aldrich	S2GPT02RE	Used to sterilize follicle wash medium
SYBR-safe DNA gel stain	Thermo Fisher Scientific	S33102	Staining to visual PCR products on agarose gel
TCM199 with Hank’s Salts	Gibco	12-350-039	Component of follicle wash medium
Triton X-100	Fisher Scientific	BP151-100	Detergent for immunostaining permeabilization buffer
Trizol reagent	Thermo Fisher Scientific	15596026	RNA isolation reagent
Trypan Blue Solution, 0.4%	Gibco	15-250-061	Used for testing viability of isolated follicles
Tween 20			Detergent for immunostaining wash buffer
Warmer Plate Universal	WTA	20931	Warm plate to keep follicles at 38.5 °C while searching under the microscope
Wiretrol II Calibrated Micropipets	Drummond	50002-005	Glass micropipettes to manipulate follicles