Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

Cornell University's Institutional Animal Care and Use Committee (IACUC) has approved all the methods described here, under protocol 2004-0038 to JCS.

1. Preparation of Instruments and Culture Media

1. Wipe the working area clean with 10% bleach and allow the bleach to remain on the work area surface for at least 5 min. After 5 min, remove the excess bleach with clean paper towels and 70% ethanol (EtOH). Clean the dissecting microscope thoroughly with 70% EtOH.
   NOTE: It is important that the work area be at least semi-sterile to avoid contamination of organ cultures.
2. Wrap clean forceps and scissors with a paper towel dampened with 70% EtOH until the time of use.
   NOTE: The ideal dissection tools may vary according to the age/size of the ovary. Best results are obtained using one sharp micro dissecting scissor, two fine tip forceps (either number 5 or 55) and one micro dissecting iris scissor.
3. In a conical tube, make 50 mL of ovary culture media and warm in a 37 °C water bath.
   1. To make ovary culture medium⁴, supplement 1x Minimal Essential Media (MEM) with 10% Fetal Bovine Serum (FBS), 25 mM (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 100 units/mL penicillin, 100 µg/mL streptomycin, 0.25 µg/mL Amphotericin B (e.g., 45 mL of MEM, 5 mL of heat inactivated FBS, 1.25 mL of HEPES, and 0.5 mL of 100x stock of penicillin, streptomycin, and Amphotericin B).
4. Prepare plates and inserts prior to the ovary culture.
   1. In a tissue culture hood, add 1 mL of ovary culture medium to a 35 mm tissue culture plate.
   2. Pre-soak insert membrane with the culture medium. Add sufficient media, usually about 0.55 mL per well, to the 24-well carrier tissue culture plate, and place a cell culture insert in the well. Make sure the insert's membrane touches the media.
   3. Prepare the number of 35 mm plates and wells with cell culture inserts according to the expected number of ovaries in the experiment. Place the 35 mm plates containing 1 mL of culture media and the 24-well carrier plate containing culture media and inserts in a 37 °C, 5% CO₂, and atmospheric O₂ incubator.
5. Arrange a hot plate next to the dissection scope and set the temperature to 37 °C. Keep a 37 °C, 5% CO₂ and the atmospheric O₂ incubator is close to the dissection room.

2. Ovary Dissection and Organ Culture

NOTE: Ovaries can be dissected at room temperature as long as the exposure to the non-ideal temperature is minimal.

1. Euthanize the female mice at post-natal day 5, by decapitation, one at a time.
   NOTE: Euthanasia must be conducted according to IACUC guidelines present at the facility where the research is being performed.
2. Spray the animal with 70% EtOH. Place the animal on the top of the dry paper towel under the dissection microscope. Place a 35 mm tissue culture dish containing ovary culture media close to the dissection microscope.
3. With forceps or a micro dissection iris scissor, make an incision through the skin of the animal and into the abdominal cavity, using caution not to cut into the intestines. Push the intestines out of the abdominal cavity and locate the ovaries. Extract the ovaries and place them into the 35-mm tissue culture dish containing culture media.
4. Once the ovaries have been dissected from the animal, increase the magnification of the dissection microscope in order to better visualize ovaries and any attached tissue. With clean fine tip forceps, remove all non-ovarian tissue, such as bursa and fatty tissue. Be careful to keep the ovaries intact when removing the attached non-ovarian tissue.
5. Once the ovaries are isolated, place the pair into the pre-soaked cell culture inserts of the carrier 24-well plate (refer to steps 1.4.2 and 1.4.3) and adjust the volume of the medium to ensure that it is sufficient to keep the organ moist (without completely submerging it).
   Caution: Be careful not to poke a hole in the membrane of the cell culture insert when transferring the ovaries.
6. Place explanted organs in a 37 °C, 5% CO₂, and atmospheric O₂ incubator.
7. Repeat steps 2.1 – 2.6 until the ovaries of each animal are dissected and placed on the cell culture insert of the 24-well plate containing the ovary culture medium.
8. Change the ovary culture medium every 2 days, using warmed ovary media aliquots from a 37 °C water bath and proceed to Part 3 of the protocol at the desired experimental time point.

3. Tissue Fixation

1. In a 15 mL conical tube, prepare 4% paraformaldehyde (PFA) in 1x phosphate-buffered saline (PBS) (e.g., add 2 mL of 16% PFA electron microscopy grade to 6 mL of 1x PBS).
   Caution: PFA is a hazardous substance and should be handled under a chemical hood.
2. Fix the cultured ovaries, without removing the tissue from the culture insert, by covering the tissue with the freshly prepared 4% PFA solution. Seal the edges of the plate with plastic adhesive (to avoid evaporation) and place the samples at 4 °C overnight.
   NOTE: In order to facilitate handling and tissue integrity, avoid displacing ovaries from the cell culture insert throughout the entire procedure. Cultured ovaries attached to the surface of the insert are advantageous for handling without damaging or losing the organ.
3. Rinse the tissue 3x with 70% EtOH and either proceed to step 4.1 or store it in scintillation vials filled with 70% EtOH at 4 °C until further processing.
   NOTE: If using tissue endogenously expressing fluorescent proteins, storing the samples with 70% EtOH may greatly reduce the signal. Alternatively, tissue can be rinsed and stored in the PBS with 0.2% sodium azide (NaN₃). NaN₃ is, however, highly toxic and poses a serious inhalation hazard. Make the stock solution in the fume hood and handle wearing appropriate personal protective equipment such as a laboratory coat and nitrile gloves.

4. Whole Mount Immunofluorescence

1. Place the fixed tissue in 1x PBS at RT for a minimum of 4 h prior to the permeabilization step 4.3.
2. Prepare permeabilization solution and blocking solution.
   1. Prepare permeabilization solution as described. To 1x PBS, add 0.2% polyvinyl alcohol (PVA), 0.1% sodium borohydride (NaBH₄) solution (from 12 wt.% in 14 M sodium hydroxide (NaOH) solution), and 1.5% polyethylene glycol tert-octylphenyl ether (non-ionic surfactant detergent). In a 50 mL conical tube, add 5 mL of 10x PBS, 0.1 g of PVA, 50 µL of NaBH₄, and 750 µL of non-ionic surfactant detergent, then add ultrapure water up to 50 mL and agitate well at room temperature until the mixture is entirely in solution.
   2. Prepare blocking solution as described. To 1x PBS, add 0.1% non-ionic surfactant detergent, 0.15% of 2.5M glycine pH 7.4, 10% normal goat serum, 3% bovine serum albumin (BSA), 0.2% NaN₃, 100 units/mL penicillin, 100 µg/mL streptomycin, and 0.25 µg/mL Amphotericin B. Prepare a stock solution of 2.5 M glycine by dissolving 93.8 g of glycine in ultrapure water to a 500 mL final volume (pH 7.4) and a stock solution of 10% NaN₃ by dissolving 5 g in 50 mL of ultrapure water; then, in a 50 mL conical tube prepare the blocking solution by adding 5 mL of 10x PBS, 50 µL of non-ionic surfactant detergent, 750 µL of glycine stock solution, 5 mL of goat serum, 1.5 g of BSA, 1 mL of NaN₃ stock solution, and 0.5 mL of 100x stock of penicillin, streptomycin, and Amphotericin B . Add ultrapure water up to a final volume of 50 mL and syringe filter the final solution.
3. Remove and discard the 1x PBS from the vial, and then add enough permeabilization solution to completely submerge the ovaries. Place the sample in permeabilization solution on an orbital shaker (e.g., nutator) at room temperature for 4 h.
   NOTE: Incubation time may vary according to the organ thickness.
4. Replace the permeabilization solution with enough blocking solution to completely submerge the ovaries. Seal the vial with the plastic adhesive and leave the tissue incubating for a minimum of 12 h at room temperature on an orbital shaker.
5. Prepare primary antibody dilution in blocking solution as per the manufacturer's datasheet.
   NOTE: Note that not all antibodies are compatible with the clearing agent. The average volume needed per sample is approximately 750 µL.
   1. For oocyte quantification, use TAp63 (nuclear oocyte marker) and MVH (cytoplasmic germ cell marker).
      NOTE: The antibodies used in the immunofluorescence images are mouse anti-p63 and rabbit anti-MVH). The primary antibody solution can be reused 2x or more if stored at 4 °C between staining protocols and properly handled to avoid bacterial growth.
6. Place the insert containing the tissue in a 24-well plate and add about 750 µL of primary antibody solution. Ensure that ovaries are completely submerged. Seal the plate with plastic adhesive to minimize evaporation and leave the tissue incubating for 4 days at room temperature on an orbital shaker.
7. Prepare washing solution.
   1. Prepare washing solution as described. Make 1x PBS, add 0.2% PVA and 0.15% non-ionic surfactant detergent. In a 50 mL conical tube, add 5 mL of 10x PBS, 0.1 g of PVA, 75 µL of non-ionic surfactant detergent and ultrapure water up to a final volume of 50 mL.
8. Remove primary antibody dilution and add a generous amount of washing solution. Always make sure to submerge the ovaries. Allow the ovaries to soak in the solution overnight at room temperature on the orbital shaker.
9. Replace the washing solution and incubate on the orbital shaker for 2 h or more.
10. Repeat the step 4.9 one additional time.
11. Prepare the secondary antibody dilution in blocking solution.
    NOTE: The secondary antibodies used are anti-mouse 488 fluorescent dye raised in goat and anti-rabbit 594 fluorescent dye raised in goat. See step 4.5. for the average volume needed per sample.
12. Remove washing solution from the ovaries and add secondary antibody solution. Protect the samples from light (wrap plate with aluminum foil) and seal the plate with plastic adhesive to minimize evaporation. Incubate samples on an orbital shaker at room temperature for 2 days.
    NOTE: Continue protecting the samples from light using aluminum foil during all subsequent incubation steps.
13. Remove secondary antibody solution from the samples and add washing solution. Incubate on an orbital shaker for 8 h.
    NOTE: If 4,6-Diamidino-2-phenylindole (DAPI) staining is desired, add 50 ng/mL of DAPI to the first wash step (~8 h incubation).
14. Replace the washing solution for an overnight wash.
15. While preparing the clearing solution (see section 5), replace the overnight washing solution with fresh solution and keep the samples on the orbital shaker at room temperature.

5. Ovary Clearing and Imaging

1. Prepare ScaleS(0) clearing solution.
   1. Prepare ScaleS(0) clearing solution¹⁰ by adding reagents in the given proportion: 40% D-(-) sorbitol (w/v), 10% glycerol, 4.3 M urea, and 20% dimethyl sulfoxide (DMSO), pH 8.1 (e.g. in a 50 mL conical tube, add 2.5 mL of glycerol, 10 g of sorbitol, 5 mL of DMSO, and 12 mL of 9 M urea for a total volume of 25 mL. Mix solution by inversion at 50 °C for 30 min and degas prior to use.)
2. Remove the washing solution and submerge the samples in clearing solution. For better results maintain samples on the orbital shaker.
3. Replace the clearing solution 2x daily until the tissue becomes transparent (approximately 2 days).
4. Once the tissue is cleared, prepare the sample for imaging by carefully removing the insert membrane containing the cultured ovaries with a fine tip scalpel and placing the sample on a glass slide.
5. Proceed to imaging the samples on a microscope capable of optical sectioning.

6. Oocyte Quantification

NOTE: There are many different computer programs that are able to quantify cells. Described below is a protocol for oocyte quantification using the non-commercial image processing package, FIJI-ImageJ¹⁴.

1. Using a flattened maximum intensity projection of the Z-stack image series for the specific cellular markers of interest (without the DAPI channel), convert the image into a black and white 8-bit image under Type in the Image drop-down menu.
2. Under the Image drop-down menu, highlight the Adjust tab and then select Threshold.
3. Manually adjust the threshold to a level that best identifies each individual oocyte. Once the level is determined, click Apply to generate a black and white image. Once completed, quantify the sample by delineating using the Oval or Freehand icon.
   NOTE: FIJI-ImageJ has different options to fine tune the image that will be used for the particle quantification. For instance, in Process | Binary tab, there are different options that can be used to improve the detection of what will be counted. In Figure 4, the options used to better individualize follicles was Erode, followed by Fill holes, and lastly Watershed.
4. Under the Analyze drop-down menu, select Analyze particles. Set the parameters according to the desired resolution.