Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

All mice used were of the genetic strain C57BL/6J (see the Table of Materials). This strain has been fully sequenced and is standard for many studies on ovarian structure and function. Mice were housed according to NIH guidelines, and procedures performed were approved by the Institutional Animal Care and Use Committee of The Jackson Laboratory. Reagents and compositions used in this protocol are listed in Table of Materials and Table 1, respectively.

1. Preparation of reagents

1. Fixative: Prepare 4% paraformaldehyde (PFA) in 1x PBS. For example, for 24 samples (0.5 mL/sample = 12 mL total volume), add 3 mL of 16% PFA electron microscopy-grade to 9 mL of 1x PBS.
2. Permeabilization buffer
   1. Measure polyvinyl alcohol (PVA) into a 250 mL plastic beaker containing 1x PBS the day before the permeabilization buffer is needed. Stir the solution overnight to allow PVA to dissolve completely.
   2. Add sodium borohydride to the dissolved PVA and allow the mixture to homogenize for approximately 3-5 min. Expect the solution to foam.
   3. Add Triton X-100 to the solution and use the buffer once the Triton X-100 is dissolved.
      NOTE: PVA takes a long time to dissolve (at least 3 h), and the penetration of the antibody depends on how well it dissolves. For larger ovaries, it is crucial to dissolve PVA completely.
3. Blocking buffer
   1. Measure bovine serum albumin (BSA) into a beaker with 1x PBS. Stir the solution until the BSA is dissolved.
   2. Add glycine, Triton X-100, Penicillin-streptomycin, and sodium azide. Stir until the solution homogenizes. Transfer the buffer into a bottle and store at 4 °C. Add normal goat serum before use.
4. Washing buffer: Measure PVA into a beaker containing 1x PBS. Stir the solution overnight to allow the PVA to dissolve. Add Triton X-100 and sodium azide. Once the buffer is mixed, store it at 4 °C.
5. ScaleS4(0) solution (pH 8.1): Dissolve D-sorbitol in PBS in a beaker with stirring at ≤100 °C. Add urea to the solution, and once the urea dissolves, turn off the heat and allow the solution to stir until completely dissolved. Add glycerol and dimethyl sulfoxide, and stir until the solution homogenizes. Store the solution at room temperature in the dark.
   NOTE: ScaleS4(0) solution must be made fresh on the first day of clearing ovaries.
6. ScaleCUBIC-1 solution
   1. Dissolve urea in a beaker containing PBS with stirring at temperature ≤100 °C. Measure N,N,N′,N′-tetrakis(2-hydroxypropyl)ethylenediamine into a separate beaker, then add the dissolved urea, and stir at a temperature of ≤100 °C until the reagents completely dissolve. Once all the reagents are in solution, turn off the heat, and cool at room temperature. Store in the dark.
   2. Degas the solution by transferring it into a filtering flask. Attach the flask to a vacuum with tubing, cover the flask, and turn on the vacuum until bubbles in the solution disappear. Use the solution after this degassing.
      ​NOTE: The solution can be stored in the dark to use for up to a month.
7. Sucrose solution: Dissolve sucrose in 1x PBS; then degas the solution before use.
   NOTE: Prepare the solution freshly before use.
8. ScaleCUBIC-2 solution
   1. Dissolve sucrose in a beaker containing PBS with stirring at a temperature ≤100 °C. Add urea and stir at a temperature of ≤100 °C until the urea dissolves. Turn off the heat.
   2. Add triethanolamine and Triton X-100. Stir until the solution homogenizes. Allow the solution to cool to room temperature and store in the dark. Degas the solution before using.
      NOTE: The solution can be stored in the dark to use for up to a month.

2. Dissection and fixation of prenatal ovaries (Figure 1A)

1. Mate adult females (≥8 weeks) and males according to the approved institutional protocol. Check for vaginal plugs every morning.
   NOTE: The morning of a vaginal plug is designated as 0.5 days post coitum (d.p.c) or E0.5.
2. On the day of dissection, prepare 4% paraformaldehyde (PFA) and aliquot ~0.5 mL into microcentrifuge tubes, placed on the bench side (~4 per pregnant female).
   NOTE: PFA is a hazardous substance and must be handled under a chemical hood.
3. Prepare three 100 mm plates containing ~10 mL of 1x phosphate-buffered saline (PBS) for each pregnant female (for harvesting uterus, for dissecting mesonephros-ovary complex, and isolating ovaries).
4. Euthanize pregnant females that carry embryos of the preferred developmental stage and proceed to dissection. Euthanize no more than 1-2 females at a time. Use an approved protocol for the euthanasia of pregnant mice (e.g., CO₂ exposure followed by cervical dislocation).
5. Once a female is euthanized, spray the abdomen with 70% ethanol. Using forceps, lift the abdominal skin and use scissors to make a V-shape incision through the abdomen and body wall. Cut along the two sides of the incision and peel the skin back to expose the internal organs and fetuses in the uterus.
6. With forceps and scissors, cut and separate the uterus with the fetuses from the abdominal fat and ovarian arteries and veins. Remove the uterus and place it into a 100 mm plate containing 1x PBS. Release the fetuses into PBS by cutting the uterine wall and puncturing the yolk sac. Cut the umbilical cord.
   NOTE: For fetuses younger than 15 days in gestation, euthanasia of the mother and removal from the uterus ensures rapid death of the fetus. Fetuses older than 15 days in gestation to birth must be euthanized by decapitation.
7. Transfer a fetus into a new plate with fresh 1x PBS. To decapitate an E15.5 fetus, delicately pin the two hindlimbs apart on the plate with forceps to immobilize the fetus. Using a second pair of forceps held in the dominant hand, hold the neck between the forceps prongs and squeeze to cut off the head of the fetus, or use scissors to cut the neck.
8. With the dominant hand forceps, cut below the forelimbs and gently pull the upper body away. Next, insert one endpoint of the dominant hand forceps into the peritoneal cavity vertically into the opening of the abdominal region while the other endpoint of the forceps remains outside. Close the forceps, pinch the body wall to cut it, and expose the internal organs.
9. With the same forceps, gently remove the organs, including the intestines and liver, until the kidney and the reproductive organs become visible. Determine the sex of the fetus: at E15.5, ovaries have an elongated sausage-like shape while the testes are oval.
   NOTE: At earlier stages, genotyping may be needed to determine the sex.
10. Using one pair of forceps, hold the female fetus down and use the other forceps to grasp each mesonephros-ovary complex and gently separate it from the kidneys and the Müllerian ducts. Place the ovaries in a new plate with 1x PBS, leave the mesonephros attached, but remove the surrounding tissues to ensure that the ovaries are exposed. Place both ovaries into ~0.5 mL of 4% PFA in microcentrifuge tubes and fix at 4 °C overnight. The next day, replace PFA with 70% ethanol.
11. For the dissection and fixation of E18.5 ovaries, separate the fetuses from the uterus as described above and then follow the protocol for prepubertal pups described in section 3 (Figure 1B).

3. Dissection and fixation of prepubertal ovaries (Figure 1B)

1. Prepare 4% PFA and aliquot ~0.5 mL into 1.5 mL tubes and place them on the bench side (~one tube per pup). Prepare 35 mm plates with 1x PBS (~one per pup). Euthanize the pups using the approved institutional protocol.
   NOTE: Decapitation is the preferred method of euthanasia for mice 8 days of age or younger. Neonatal mice can be decapitated using either 3-4" dissecting scissors or 5-7" Mayo scissors.
2. Determine the sex of pups by measuring the anogenital distance, which is greater in males than in females. Use sharp decapitation scissors to decapitate female pups. Place the body, open-cut side down, on a paper towel to drain the blood (~5-10 s).
3. Secure the pup on its back by placing a pin in the paw pad of each foot. Spray the abdomen with 70% ethanol. Use forceps to hold the pup's skin away from the body and use scissors to make a small V-shaped incision in the skin of the lower abdomen.
4. Insert the scissors into the incision under the skin and cut along both sides of the abdomen. Use the forceps to gently peel back the skin and reveal the lower abdomen. Gently move the intestine up and locate the uterine horns next to the bladder. Follow each uterine horn towards the kidney and locate the ovary with surrounding adipose tissue below each kidney.
5. To dissect the ovaries, hold the uterine horn below the ovary and oviduct with one pair of forceps (forceps A, Figure 1B) and gently lift it away from the body. Using another pair of forceps (forceps B), gently grab underneath the first pair of forceps so that the ovary and oviduct in the adipose pad are visible above the second pair of forceps. Then, cut below the second pair of forceps with scissors or carefully pull to detach the ovaries and attached tissues.
6. Place the isolated organs in a plate with 1x PBS. Trim away the extra adipose tissue, oviduct, and uterine horn to clean and isolate the ovary. Delicately open the bursa surrounding the ovary and expose the whole ovary.
7. Trim the bursa membrane without damaging the ovary and leave tissue around the hilum (ligament-like structure between the ovary and the reproductive tract) to handle the ovary, as shown in Figure 1B, P5. Once the ovaries are trimmed, place them in 4% PFA and fix the ovaries at 4 °C overnight. The next day, replace 4% PFA with 70% ethanol and store the ovaries at 4 °C until the immunostaining protocol step.

4. Perfusion, dissection, and fixation of pubertal ovaries (Figure 1C)

1. Perfuse mice according to institutionally approved protocols with the mouse under deep anesthesia in a chemical fume hood. Prepare the perfusion equipment and reagents. See the detailed protocol at ⁴².
   NOTE: Perfusion is a terminal euthanasia procedure that replaces the blood throughout the vascular system with a tissue fixative.
2. Weigh a pubertal female mouse (P28) and record the mouse's weight (typically between 13 g to 15 g). Calculate the amount of the approved anesthesia agent to be used.
   NOTE: Here, 0.35 mL of tribromoethanol per 10 g of body weight was used in this protocol.
3. Use a 10 mL disposable syringe with a 20 G needle to fill the syringe with the approved anesthetic agent. Replace the 20 G needle with a 26 G x 3/8 needle.
4. Carefully restrain the mouse by scruffing the dorsal neck skin with one hand. Turn the mouse over to expose the abdomen. Inject the previously calculated amount of anesthetic into the peritoneal cavity. Place the mouse in a container with a cover until the anesthesia takes effect (i.e., the mouse no longer moves).
5. Once the mouse stops moving, gently pinch its feet to check that there is no response and ensure that the mouse is fully anesthetized before initiating the perfusion. Place and secure the mouse on its back by gently pinning all four legs through the paw pads onto a board. Ensure that the legs are pinned in a relaxed position, not overstretched.
6. Spray the mouse with 70% ethanol from the abdomen to the chest region. Use forceps to gently pinch and lift the abdominal skin and then use scissors to make a small V-shaped cut through the skin and body wall.
7. Lift away the skin flap from the body cavity and insert the scissors below the skin and body wall. Cut the skin and body wall straight up toward the head to the edge of the thoracic cavity at the sternum.
8. Open the thoracic cavity by carefully cutting the ribs on both sides of the sternum to just below the scapula. Take care not to puncture the lungs below the rib cage. Grab the skin/rib flaps with forceps and pin them to expose the internal organs.
9. Gently trim any tissue, such as the thymus, that may obstruct clear access to the heart. Expose all organs, including the gastrointestinal tract, to allow visual confirmation that the whole body is perfused, including the lower abdomen.
10. Use dissection scissors to snip the right atrium of the heart and release blood from the system. Gently hold the heart with forceps and insert a perfusion needle (connected to a peristaltic pump controller) into the left ventricle.
11. Pump ~10 mL of PBS, with gentle but constant pressure, until tissues such as the liver, kidneys, and reproductive tract (Figure 1E,F) turn from pink to white. Next, replace the PBS with freshly made 1% PFA and continue perfusion with approximately 10 mL of PFA or until twitching occurs in the lower body of the mouse (e.g., tail).
    NOTE: These fixation tremors indicate successful perfusion.
12. Once perfusion is completed, locate the fat pad with the ovaries below the kidney and gently dissect the entire reproductive tract, including the fat pad, ovaries, and uterine horns, and place the tract in a vial with 4% PFA. Fix the ovaries at room temperature overnight (~16-24 h). Then, replace the 4% PFA with 70% ethanol for at least one day and trim the ovaries (Figure 1E,F), as described above, before starting the immunostaining protocol.

5. Whole-mount ovary immunostaining (Figure 2A)

NOTE: Practice sterile techniques during the immunostaining protocol, especially when changing buffers, to prevent contamination during extended incubation periods.

1. Perform all immunostaining steps in a 24-well plate, as shown in Figure 2A. Adjust reagent volumes for other formats. Change buffers carefully to avoid losing small prenatal and prepubertal ovaries.
   NOTE: Other formats, such as 48- and 96-well plates, can be used, but the deeper wells and narrower openings make it difficult to aspirate buffers without losing or damaging ovaries.
2. Day 1
   1. Pipette 1 mL of PBS/well into the number of wells needed in a clean 24-well plate. Cut off the tip of a 200 µL pipette tip to make a wide opening and use it to gently transfer prepubertal ovaries from 1.5 mL tubes with 70% ethanol into the wells. For pubertal ovaries, use a 1000 µL pipette tip with the tip cut off for the transfer.
      NOTE: The wider opening of the tip prevents tissue damage during transfer.
   2. Once all the samples are transferred into the wells, use a new tip to aspirate the PBS and replace it with fresh 0.8 mL of PBS/well. Incubate the samples at room temperature on a shaker at ~100-150 rpm overnight. Start preparation of the permeabilization buffer (see Table 1 and section 1).
      ​NOTE: The new tip prevents ovaries from sticking to the tip, and the PBS incubation step allows the ovaries to equilibrate in PBS and rehydrate before staining.
3. Day 2
   1. Finish preparation of the permeabilization buffer. Aspirate PBS from the wells and replace it with 0.8 mL of the permeabilization buffer per well. Place the plates back on the shaker and incubate the samples at room temperature for 4 h.
   2. After 4 h of incubation, completely aspirate the permeabilization buffer and replace it with 0.5 mL of blocking buffer (see Table 1 and section 1). Seal the plate with parafilm to prevent evaporation, place in a securely covered container on a shaker, and incubate the samples with gentle agitation in blocking buffer overnight (16-24 h) at room temperature.
4. Day 3
   1. Prepare the primary antibodies by dilution in blocking buffer. For visualizing oocytes, use oocyte markers, e.g., rat anti-germ cell nuclear acidic peptidase (GCNA)/TRA98 for both prenatal and prepubertal ovaries and rabbit anti-DEAD-box helicase 4 (DDX4)/mouse vasa homologue (MVH) for both prepubertal and pubertal ovaries. For quantifying LINE-1 ORF1p expression in prenatal ovaries, use rabbit anti-LINE-1 ORF1p or another available antibody.
      NOTE: GCNA/TRA98 signal is not detectable in the pubertal ovaries.
   2. Aspirate the blocking buffer from the samples and replace it with 0.5 mL of diluted primary antibodies. Seal the plate with parafilm to prevent evaporation. Place the plates in a container with a cover to prevent the drying up of samples during incubation. Place the container on the shaker and incubate the samples for 3-4 days at room temperature.
5. Day 7
   1. Gently remove the parafilm from the plates and aspirate the primary antibody mixture from the samples. Store the primary antibody mixtures at 4 °C (for up to a month or three uses) and reuse them later by supplementing with fresh antibodies (e.g., 2 µL of fresh antibody aliquot per 5 mL of the previously used mixture).
   2. Add 1 mL of washing buffer (see Table 1 and section 1) per well to the samples and gently rock the plates for a few seconds to rinse away the residual antibodies. Carefully aspirate and replace the washing buffer with 0.8 mL of fresh washing buffer and shake the plates at room temperature overnight.
6. Day 8
   1. Aspirate the overnight washing buffer, replace it with 0.8 mL of fresh washing buffer, and shake at room temperature for 2 h. Repeat the washing step with fresh washing buffer for another 2 h.
   2. After the last wash, replace the washing buffer with 0.5 mL of secondary antibody mixes diluted in a blocking buffer, e.g., goat anti-rat Alexa Fluor 555 and goat anti-rabbit Alexa Fluor 647 at 1:1,000 dilution. Prepare the secondary antibody mixture fresh for each immunostaining experiment.
   3. Seal the plate with parafilm to prevent evaporation during incubation. Incubate the samples at room temperature in the dark or in plates covered with aluminum foil with gentle agitation for 3 days.
      NOTE: This step and all subsequent steps take place in the dark or with aluminum foil wrap.
7. Day 11
   1. Aspirate the secondary antibody mixture and perform an initial wash with 1 mL of washing buffer. Gently rock the plates for a few seconds to rinse away the residual secondary antibodies.
   2. Aspirate the washing buffer, replace it with 0.8 mL of fresh washing buffer, and incubate the samples at room temperature for 2 h with shaking, protected from light. Repeat the washing step 2 times for a total of 3 washes. After the third wash, proceed to the clearing step (section 6).

6. Clearing of immunostained whole-mount ovaries (Figure 2A).

NOTE: Perform all clearing steps in the dark by wrapping the plates in aluminum foil or placement in opaque containers. The clearing steps differ for prepubertal and pubertal ovaries.

1. Day 11 [Prenatal and prepubertal ovaries are cleared in a one-step protocol]
   1. After the last wash, aspirate the washing buffer and replace it with 0.8 mL of freshly made ScaleS4(0) solution (see Table 1 and section 1). Seal the plate with parafilm and incubate the samples in the dark with shaking at room temperature for three to four days before imaging.
   2. Replace the ScaleS4(0) solution with a fresh solution daily if needed; Take care while replacing the solutions, as cleared ovaries become transparent and are difficult to see. After clearing, proceed to sample set up and imaging (section 7).
      NOTE: Changing solutions did not significantly improve image quality. Daily changes increase the likelihood of losing small and mostly transparent samples.
2. Days 11-15 [Pubertal ovaries are cleared in a two-step protocol]
   1. After the last immunostaining wash, aspirate the washing buffer and replace it with 0.8 mL of ScaleCUBIC-1 (see Table 1 and section 1). Seal the plate with parafilm and gently shake samples at 37 °C for 3 days in the dark. Replace the ScaleCUBIC-1 solution with the fresh solution daily.
   2. On day 15, wash the samples 3 times for 10 min each time in 0.8 mL of PBS at room temperature with gentle shaking. Replace the PBS with 0.8 mL of degassed 20% sucrose prepared freshly with PBS. Shake gently at room temperature for 1 h.
   3. Replace the 20% sucrose solution with 0.8 mL of ScaleCUBIC-2 Solution (see Table 1 and section 1), seal the plate with parafilm, and shake gently at room temperature with daily changes of the ScaleCUBIC-2 Solution. Clear for 4-5 days and proceed to image. Take care when exchanging solutions to avoid sample loss as cleared ovaries are transparent and difficult to see. Proceed to sample set up and imaging (section 7).

7. Sample setup and imaging with a multiphoton microscope

NOTE: All steps described below were performed with a Leica DIVE/4TUNE/FALCON with two tunable mode-locked Ti:Sapphire multiphoton lasers with a pulse duration of 120 fs with a multi-immersion 16x/NA0.6 objective (immersion liquid = glycerol) with a maximal working distance of 2.2 mm. See the Table of Materials for details about the image acquisition software. Supplemental Table S1 and Supplemental Figure S1 show the settings used for this protocol. For other imaging platforms, consult with the microscopy core or follow the manufacturers' specifications/recommendations.

1. Sample setup
   1. Prepare the setup for mounting the samples. For example, use a sticky and flexible silicone gasket to create an adhesive well. Place the adhesive well on one 25 mm x 25mm No. 1.5 micro coverslip to create a well in which the ovaries will be placed (Figure 2B).
      NOTE: A gasket of 1 mm thickness accommodates all sizes of ovaries. This setup is recommended as it eliminates ovary movement during imaging, which can occur when ovaries are placed in mountant in a glass-bottom dish.
   2. Using a pipette tip with the tip cut off (20 µL for prenatal and prepubertal), transfer the ovaries with ~5-10 µL of ScaleS4(0) solution into the middle of the adhesive well as shown in Figure 2B. For pubertal ovaries, use a 200 µL pipette tip with the tip cut off far enough to transfer the ovaries. Add ~15-20 µL of the ScaleCUBIC-2 Solution into the middle of the adhesive well.
      NOTE: Too much solution may cause leaking onto the microscope stage, and too little solution will result in poor image quality. Use a dissection microscope to transfer samples as cleared ovaries become transparent and are difficult to see.
   3. Once the ovaries are transferred into individual wells with a drop of clearing solution, gently place another cover glass on the adhesive well, press to create a seal, and keep the samples in place in a small pool of clearing solution sandwiched between the two coverslips (Figure 2B). For imaging, place the coverslip "sandwich" in a 3D printed microscope adapter slide (e.g.,⁴³ ).
      NOTE: Adhesive wells are reusable after deconstructing the "sandwich" and washing with water.
2. Imaging (Supplemental Table S1 and Supplemental Figure S1)
   1. Turn on the microscope and software according to the guidelines of the microscopy core or the manufacturer. Place the mounted samples onto the microscope stage and look through the eyepiece to locate the sample lit with a low-power LED fluorescence lamp.
   2. Once the samples are located, turn on the laser(s) and assign each detector to a specific Alexa Fluor dye/marker. For multiple lasers, allow for sequential image acquisition. Acquire the whole stack prior to switching lasers.
      NOTE: For this protocol, one laser was used for the image acquisition of both 555 nm and 647 nm fluorophores.
   3. Set up the parameters for image acquisition according to the microscopy core's or the manufacturer's specifications. Use bidirectional image acquisition, if available, to reduce the scanning time and improve efficiency. Adjust other settings, including the line average (1), frame average (1), and frame accumulation (1).
      ​NOTE: Bidirectional image acquisition allows lasers to scan in both directions as they move on the X-axis.
   4. Set up the Z-Stack by identifying the beginning (bottom of the sample/bottom glass cover) and the end position (top of the sample/top glass cover). Select a Z-step size of 2 µm for prepubertal ovaries and 5 µm for pubertal ovaries.
   5. Activate the Z-compensation feature, and within this feature, activate the excitation gain. Set the Z-compensation for the bottom and top of the sample by selecting a laser intensity for both the bottom (low intensity) and top stack (high intensity). See Supplemental Figure S1, Linear Z-compensation, where excitation gain box is selected, and laser intensity for the bottom (0) and top (347.75) is set as 10 and 12, respectively.
   6. Use the tiling mode for larger samples that cannot be captured in a single field of view. Activate the image navigator and indicate the number of tiles needed to capture the entire sample using the rectangular marking tool.
   7. Start the image acquisition. For images with multiple tiles, start image acquisition with the navigator to capture all tiles. Once the image acquisition is completed, first save all images, and if multiple tiles were acquired, then run the mosaic-merge tool to merge the tiles into a single image.
   8. Save the merged files into a separate project folder to make it easier to work with the Gigabyte-sized images. Transfer the files for image processing with 3D image visualization and analysis software. Figure 3 shows representative images taken at different stages with two markers (GCNA and DDX4).

8. Image processing

NOTE: All steps described below were developed and performed using IMARIS 3D image visualization and analysis software.

1. Import the image(s) into the 3D image visualization and analysis software and, if needed, convert the files from the original format (e.g., .lif, .czi, .lsm, .lei, .oib, .oif, .nd2) to the .ims format.
2. If needed, process the images with the gaussian filter to reduce pixelation (Figure 4A). Using the 3D view feature, position the image and deselect the frame option (to remove the frame).
   NOTE: There are other optional filters, that can also be used to decrease pixelation and reduce background intensity.
3. Enlarge the image to a specific scale and use the Snapshot feature. Set the preferences as follows: select 300 DPI, save as TIFF Image, and Transparent Background. Take a snapshot of the image and save it. Figure 3 shows complete assembled images.

9. Oocyte Quantification

NOTE: Whole ovary immunofluorescence and 3D image visualization and analysis can be used for the estimation of oocyte numbers in whole ovaries (Figure 3 and Figure 4) using the Spot feature. The GCNA signal can be used to quantify oocytes in prenatal and prepubertal ovaries, as shown in Figure 4 (P5). In pubertal ovaries, use the DDX4 signal to quantify two oocyte populations in non-growing follicles ("ring-like" structure, closed arrow) and growing follicles (large structures, open arrow, Figure 4, P28).

1. Determine the size of oocytes that will be used as a parameter to quantify oocytes with the Spots feature in step 9.2.
   1. Open the image and select the Slice option to open the Z-stack image.
   2. Select the Line option in the Measure panel and measure the distance by drawing a line from one edge of an oocyte/marker to the other edge at the widest point to determine the XY diameter of the oocyte type/size to be counted. Move through the stack and obtain the range of diameters for multiple oocytes of the same type. Use the shortest length as the size selection criterion for oocyte counts.
2. Spots feature: Select the 3D View option and activate the Add new Spots function in the Scene panel to open the Algorithm panel. Deselect all the algorithm settings as the size selection filter with the oocyte size obtained in step 9.1 will be used.
   1. Click on the single forward arrow to move to the Source Channel. Select the Channel with the preferred marker and type in the Estimated XY Diameter obtained in step 9.1. For example, use an estimated XY diameter of 7 µm for prenatal oocytes and 11 µm for P5 oocytes (GCNA = green signal; Figure 3 and Figure 4A).
   2. Use 30 µm for the DDX4 signal to estimate the number of oocytes in growing follicles in P5 ovaries (DDX4 = magenta signal; Figure 3 and Figure 4A). For pubertal ovaries, use the DDX4 signal for oocyte quantifications. For example, use XY diameters of 15 µm and 80 µm to identify oocytes within non-growing follicles and growing follicles, respectively, as shown in Figure 4A.
      NOTE: The size selected may vary due to image acquisition criteria and the type of microscope used. The selection may also vary among genetic strains as ovaries with more oocytes will require smaller size selections for better automated oocyte detection.
   3. After size selection, activate the Model PSF-elongation and Background subtraction, which are automatically determined. Select the single forward arrow to move to the Filter Spots panel. Add the Quality filter and determine a threshold. To ensure the accurate estimation of oocyte numbers, enlarge the image as the threshold is adjusted to choose a threshold value that selects most of the oocytes. Click on the double arrow to finish automated counts.
   4. Manually check the selected oocytes using the Edit tab to select missed oocytes or deselect non-oocyte particles selected by the automatic threshold. Record the data in the Statistics tab under the Overall tab for further analysis.
      NOTE: The parameters used for counting the spots can be saved and used for another sample set but it is recommended to do a manual check before running batch analysis.
   5. To store the parameters, click on the Creation tab and click on Store Parameters for Batch.
      ​NOTE: Nuclear markers (GCNA+ oocytes, Figure 4A) are easily identified by the spot feature and may not require much manual adjustment. However, the cytoplasmic marker (DDX4+ oocyte, Figure 3 (P28, closed arrow) and Figure 4A) will require more manual adjustment to ensure the identification of the correct size/type of oocytes.

10. Quantifying protein expression in ovaries

NOTE: There are several ways to quantify oocyte expression of specific markers using both the Spots feature (section 9 and step 10.1) and Surfaces feature (step 10.2). The Spots feature can be used for proteins with distinct localization patterns such as nuclear markers (GCNA), and the Surfaces feature can be used for proteins with non-uniform localization patterns as shown in Figure 5A where LINE-1 ORF1p intensities in E15.5 and E18.5 ovaries were measured. To calculate and compare the intensity of the protein of interest between two samples (e.g., timepoints, treatments, or genotypes), collect images with the same properties. Use samples with a more intense signal to determine the parameters that can be stored and used for the other samples.

1. To obtain the protein expression with the Spots feature, first identify oocytes as highlighted (section 9).
   1. Then under the Spots feature, select the Statistics tab, followed by the Detailed tab. Click on the down arrow and select Specific Values, which will open another tab beneath it containing different measurements. Select the Intensity Mean of the channel used for the surface generation (or select other intensity measurements such as the Intensity Median).
   2. To obtain the combined/average statistical information, select the Average Values criterion. Once done, export and save data for further analysis.
2. To obtain the protein expression with the Surfaces feature, open the image, select the 3D View option and activate the Add new Surfaces function in the Scene panel to open the Algorithm panel. Deselect all the algorithm settings if not needed and click on the single forward arrow to move to the Source Channel.
   1. Select the Channel with the antibody signal to measure. Other properties to select are user-specific options. Here, the LINE-1 signal was used as a marker. The Surfaces Detail and Background Subtraction (Local Contrast) values were kept at default values of 1.42 and 5.33, respectively.
   2. Click on the forward arrow to move to the Threshold panel. Choose a threshold value that is comparable in both samples (e.g., here, LINE-1 expression in E18.5 was used to choose a threshold). Select Enable with a default Seed Points Diameter of 7.10 (this value was found optimal for the images but may depend on the expected pixel-size of features).
   3. Click on the forward arrow to move to the Filter Surfaces panel. Use a Quality filter was used, and base the value, as with the Threshold value, on the expression of the proteins in both ovary samples. Once the filter property is selected, click on the double forward arrow to complete the generation of the surface.
   4. To obtain the intensity values, select the Statistics tab, followed by the Detailed tab. Click on the down arrow and select Specific Values, which will open another tab beneath it containing different measurements.
   5. Select the Intensity Mean of the channel used for the surface generation (or select other intensity measurements such as the Intensity median).
   6. To obtain the combined/average statistical information, select the Average Values criterion. Once done, export and save the data for further analysis (Figure 5C).

11. Estimation of total oocyte numbers in damaged ovary with computational correction

NOTE: If a minor ovary damage occurs during dissection, it may be possible to computationally estimate total oocyte count. It is recommended to use intact ovaries from the same strain and developmental stage for oocyte number estimation as shown in Figure 6. Simulations performed with ovaries at E15.5 indicate that correcting for a ≥30% loss results in a significant deviation from actual numbers (Figure 6C).

1. Open images of intact ovaries with IMARIS and select the Frame feature. Under the Frame Settings, select the Box, Grid, Tickmarks, and Axis labels. Under the Position X/Y/Z tab, select 200 µm to generate tickmarks with 200 µm spaces in all X/Y/Z positions.
   NOTE: The tickmarks create a grid/ruler in X, Y and Z planes to identify oocytes within a specific region.
2. Activate the Spots feature to identify the oocytes as highlighted in section 9 (Figure 4). Create a 3D model using the oocytes identified in all intact ovaries being used for the simulation by selecting Sphere under the Points Style/Quality tab in the Spots feature.
   NOTE: The Pixel Width can also be changed under the Points Style/Quality tab.
3. With the box generated in step 11.1, align the 3D models of the intact ovaries in the same orientation as shown in Figure 6A.
4. Using the 200 µm tickmarks (step 11.1), select oocytes that fall within 50% of the volume of each ovary. Click on the Spots feature and select Edit Labels to classify the oocytes that fall within the 50% region by color, as shown in Figure 6A.
   NOTE: Once oocytes are classified in a region, the number of oocytes that fall within each class will be provided.
5. Zoom into the 50% region and change the tickmarks from 200 µm to 50 µm to make a smaller ruler for oocyte identification. Maintain the zoom % in all images. With the 50 µm tickmarks as a guide, divide the 50% region into five equal parts.
   NOTE: Oocytes within each part will represent 10% of the volume as highlighted in Figure 6A, where an intact ovary shows oocytes in the top half of the ovary classified by five different colors with each color representing oocytes within a 10% volumetric region.
6. Record the oocyte numbers in each 10% region as shown in Figure 6A,B. Calculate the average number of oocytes for 10, 20, 30, 40, and 50% increments.
7. Use the highlighted in steps 11.1 to 11.6 to obtain oocyte numbers in the damaged partial ovary. Position the damaged ovary in the same orientation as the intact ovaries and estimate missing volume/percentage as described above.
8. To estimate the total oocyte number in the whole ovary, add the average number of oocytes calculated for a similar volume to the number obtained from the partial ovary (Figure 6B).