Chromatin Spread Preparations for the Analysis of Mouse Oocyte Progression from Prophase to Metaphase II

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of Johns Hopkins University. Experiments were performed on wild-type C57BL/6J mice.

1. Harvesting Fetal or Neonatal Ovaries and Preparation of Prophase Chromatin Spreads

1. To extract embryos at 14.5–19.5 days post-coitum (dpc) sacrifice the pregnant females via cervical dislocation or CO₂ asphyxiation according to IACUC guidelines.
   NOTE: For postnatal day 1–5 ovaries skip to step 1.3. Figure 1 summarizes the meiotic prophase stages enriched at different embryonic and post-natal ages.
2. Open the abdominopelvic cavity using sterile scissors, making a V-shaped opening. Dissect out the maternal uterine horn, separate the embryos from the placenta, and transfer embryos into 35 mm petri dishes containing 3 mL of pre-warmed 1x phosphate buffer saline (PBS) at 37 °C.
   NOTE: A fliptop incubator set at 37 °C can be used to maintain temperature. In addition, a temperature regulated glass stage can also be used to maintain temperature during ovary manipulation.
3. With 3.5 inch surgical scissors, sacrifice embryos or pups via decapitation according to IACUC guidelines. Place decapitated embryos or pups in pre-warmed PBS prior to further dissection.
4. Dissect one embryo or pup at a time in a separate 35 mm petri dish containing 3 mL of pre-warmed PBS at 37 °C. Proceed by cutting along the ventral midline of the posterior half of the embryo, along the anterior half below the forelimbs and directly above the hind-limbs and tail as outlined in Figure 2A–B.
5. Open the abdomen using 3.5 inch surgical scissors. Using fine-tipped forceps displace or remove the liver and loops of bowel, exposing the ovaries in a new 35 mm petri dish containing 3 mL of pre-warmed PBS at 37 °C (see guide in Figure 2B). The ovaries are located immediately below and behind the kidney towards the posterior wall of the peritoneal cavity (Figure 2B–C). A guide for differentiating between male and female gonads is provided in Figure 2D.
   NOTE: For optimal conditions, rapidly dissect out the ovaries after pregnant female is sacrificed.
6. Remove both ovaries from each female fetus using a pair of fine-tipped forceps under a dissection scope and place in a watch glasses or separate wells of a multi-well plate containing 0.5 to 1.0 mL of pre-warmed PBS and maintain at 37 °C.
7. Place each pair of ovaries in 0.5 mL hypo-extraction buffer (17 mM trisodium citrate dihydrate, 50 mM sucrose, 5 mM EDTA, 0.5 mM DTT, 30 mM Tris-HCl, protease inhibitor, pH 8.2) in a clean watch glass or a small well of a 9-well plate, making sure to immerse the ovaries completely. Incubate for at least 15 min, but no more than 30 min.
   NOTE: Make hypo-extraction buffer fresh and use within 2 h of DTT addition.
8. During incubation, using a hydrophobic barrier PAP pen, draw two 22 x 22 mm² squares on a clean glass 25 x 75 mm², 1 mm thick microscope slide as shown in Figure 3A.
   NOTE: Slides can be prepared ahead of time.
9. Pipette 45–50 µL of 100 mM sucrose onto a clean slide and transfer one pair of ovaries to the drop.
10. Using the sharp ends of two 27 G needles, tease the ovaries apart to release cells into the sucrose solution. With forceps, remove large pieces of ovary and carefully pipette sucrose solution to disperse cells.
11. Place 40 µL of fixative solution (1% paraformaldehyde (PFA), 0.2% detergent (see Table of Materials), pH 9.2) into each square of the prepared slide. With a 200 µl pipette tip, spread the fixative solution evenly over the slide surface.
12. Pipette 20 µL of the sucrose mix onto the each square of the slide containing the fixative.
    NOTE: This equates to using one pair of ovaries per slide.
13. Incubate the slides in a closed humid chamber overnight at room temperature.
    NOTE: The overnight incubation can range around (12-15 hrs) at room temperature.
14. Open the chamber lid and allow the slides to air-dry completely.
15. Wash slides in a Coplin jar containing 50 mL of 0.4% wetting agent-PBS solution (see Table of Materials) for 2 min, air dry, and proceed to immunolabelling protocol (step 6).
    NOTE: It may be possible to store the slides at -80 °C for later use, but this varies depending on the protein of interest. For optimal results proceed immediately to the immunolabelling protocol (step 6).

2. Metaphase I Oocyte Collection

1. To maximize the number of antral follicles isolated from each mouse, intraperitoneally inject sexually mature virgin female mice with 5 IU of pregnant mare's serum gonadotropin (PMSG, also known as equine chorionic gonadotropin (eCG)).
   NOTE: For optimal oocyte yield, mice should be 1 to 3 months of age, as the number of oocytes harvested will decrease with age. To prepare PMSG, dissolve bottle of 5,000 IU in 100 mL of sterile PBS (5 U/0.1 mL) Store in single-use aliquots of 600 µL at -20 °C.
2. After 44–48 h, prepare collection medium, minimum essential medium alpha (MEMα) medium supplemented with 5% fetal bovine serum (FBS) and 3 mg/mL bovine serum albumin (BSA; MEMα/BSA/FBS). Sterilize media through a 0.2 µm pore filter. Decant 2.5 mL of MEMα/BSA/FBS medium into one glass dish per mouse and warm to 37 °C in a 5% CO₂ incubator.
   NOTE: Other collection and culture media that are commercially available (M2 and M16) are also commonly used.
   Caution: We recommend removing culture dishes from the 5% CO₂ incubator one at a time for limited duration to minimize ambient air exposure during manipulation steps.
3. Sacrifice the mice via cervical dislocation or CO₂ asphyxiation according to IACUC guidelines.
4. Open the abdominopelvic cavity using sterile scissors, making a large V-shaped opening. Using forceps, displace intestines towards the head of the mouse. Locate each uterine horn; the ovaries are present proximal to the rib cage. Hold the oviduct with fine forceps and cut the fat superior to the ovary using dissection scissors (Figure 4). Continue to hold the oviduct, and with another set of fine forceps release the ovary from the bursa and place into collection dish containing MEMα/BSA/FBS medium.
   NOTE: As in step 1, it is imperative to keep the ovaries at 37 °C and maintain at 5% CO₂. A fliptop incubator hooked up to 5% CO₂ mix can be used to allow for optimal maintenance of temperature and CO₂ levels. In addition, a temperature regulated glass stage should also be used to maintain temperature during oocyte manipulation.
5. Using a 1 mL syringe with a 27-gauge needle (or similar size), release cumulus oocyte complexes by manually puncturing the large antral follicles.
6. While observing through a dissection microscope, collect GV-staged oocytes using a mouth-operated glass pipette or capillary, or a hand-operated micrometer-syringe (Figure 5). Only collect oocytes that are released from antral follicles. GV oocytes have a diameter of approximately 90 µm. GV oocytes may be surrounded by 2-3 layers of granulosa cells, and have a total diameter of around 200 µm.
7. Culture oocytes in a clean watch glass or a 9-well plate containing MEMα/BSA/FBS medium for 6 h to reach metaphase I at 37 °C in a 5% CO₂ incubator.
8. Proceed to step 4.

3. Metaphase II (MII) Oocyte Collection

1. To maximize oocytes isolated from each mouse, intraperitoneally inject sexually mature virgin female mice with 5 IU of PMSG. After 44–48 h, intraperitoneally inject with 5 IU of human chorionic gonadotropin (hCG).
   NOTE: For optimal oocyte yield, mice should be 1 to 3 months of age, as the number of oocytes harvested will decrease with age. To prepare hCG, dissolve a bottle of 10,000 U in 200 mL PBS (5 U/0.1 mL). Store in single-use aliquots of 600 µL at -20 °C.
2. After 12-14 hrs, prepare MEMα/BSA/FBS collection medium, as described in step 2.2, above.
3. Sacrifice the mice via cervical dislocation or CO₂ asphyxiation according to IACUC guidelines.
4. Open the abdominopelvic cavity using sterile scissors, making a large V-shaped opening. Using forceps, displace intestines towards the head of the mouse. Locate each uterine horn, the ovaries are present proximal to the rib cage. Hold the oviduct with fine forceps and cut the fat superior to the ovary using dissection scissors (Figure 4). Then remove the ovary and oviduct and place into collection dish containing MEMα/BSA/FBS medium.
   NOTE: As in steps 1 and 2, it is imperative to keep the ovaries at 37 °C and maintain at 5% CO₂. A fliptop incubator hooked up to 5% CO₂ mix can be used to allow for optimal maintenance of temperature and CO₂ levels. In addition, a temperature regulated glass stage should also be used to maintain temperature during oocyte manipulation.
5. Using a 1 mL syringe with a 27 G (or similar size) or sharp forceps, tear a hole in the ampulla of the oviduct to release the MII oocytes.
   NOTE: Be careful not to damage the oocytes in the ampulla. The ampulla will appear swollen and translucent such that the oocytes are visible (Figure 4).
6. Harvest MII oocytes from the ampulla of the oviduct into a new dish with collection medium using a mouth-operated glass pipette or capillary, or a hand-operated micrometer-syringe (Figure 5).
7. Proceed to step 4.

4. Oocyte Denuding and Zona Pellucida Removal

1. Prepare 300 IU/mL of hyaluronidase in MEMα medium supplemented with 3 mg/mL BSA to denude oocytes of surrounding cumulus cells (2.5 mL/mouse) in a watch glass, and keep at 37 °C, 5% CO₂.
   NOTE: Hyaluronidase efficacy sharply decreases after 1 h of preparation. For MII oocytes, hyaluronidase treatment is not required.
2. Expose oocyte-cumulus cell complexes to 300 IU/mL of hyaluronidase in MEMα/BSA for 3 min to denude oocytes of surrounding cumulus cells.
   Caution: Do not exceed 3 min exposure to hyaluronidase, as it will damage the oocytes.
3. To wash the oocytes, transfer oocytes to a fresh warmed MEMα/BSA medium dish. Using a small mouth-operated glass pipette or capillary (slightly larger than the diameter of the oocyte, which is approximately 90 µm), pipette the complexes up and down to completely detach the cumulus cells. Allow the oocytes to recover in the incubator while preparing for the next step.
4. Warm acidic Tyrode's solution, MEMα/BSA, and Waymouth's media to 37 °C (500 µL/mouse). Place 300 µL in each well of small 9-well glass plates.
5. To remove the zona pellucida (ZP) transfer 5–10 oocytes into Tyrode's solution. Expose the oocytes for only 30-45 s to the solution.
   NOTE: Too little exposure to the solution will not remove the ZP completely, which will prevent the oocytes from bursting and chromosomes from spreading. Too much exposure will damage and kill the oocyte. Watching the ZP dissolve under the dissection microscope can aid in optimizing exposure time.
   Caution: Using fresh Tyrode's solution is critical, as its function declines with age. Use a fresh well of Tyrode's solution for each group of oocytes treated to ensure uniform digestion of the ZP.
6. Immediately after ZP removal, wash the oocytes by transferring into warmed MEMα/BSA/FBS medium. Repeat this wash step.
   NOTE: Be careful when transferring, as oocytes will easily stick together and to the glass pipette or capillary following removal of the ZP. Pre-wetting the glass pipette or capillary in Waymouth's medium will minimize oocytes sticking together.
7. Transfer and allow the oocytes to recover for 30 minutes in Waymouth's medium at 37 °C in a 5% CO₂ incubator.
8. Proceed to step 5.

5. MI and MII Oocyte Chromatin Spreads

1. Using a PAP pen, draw a 11 x 44 mm² rectangle on the glass slide as shown in Figure 6A.
2. Coat the slide with a thin layer of fixative (1% paraformaldehyde, 0.2% Triton-X 100 in H₂O, pH 9.2) by pipetting 100 µL of fixative onto the slide and rocking the slide back and forth. Tap the slide to get rid of excess fixative.
3. Pick up between 5-10 oocytes with a small mouth pipette needle with as little media as possible and drag the needle in a line across the fixative-coated slide while depositing the oocytes from 1 cm above the slide into the fixative solution. Perform this step using a dissection microscope to ensure that the oocytes have been deposited and that they have burst.
   NOTE: The oocytes should burst immediately on the slide. The height at which the oocytes are deposited impact the degree of chromatin spreading. See representative results in Figure 6 for further details.
4. Incubate slides at room temperature in a closed humidified chamber overnight to allow the chromatin to adhere.
5. Allow the slides to air-dry completely and rinse slides in a Coplin jar containing 50 mL of 0.4% wetting agent-H₂O, pH 8.0.
6. Proceed to step 6.

6. Immunolabelling

1. Prepare the antibody dilution buffer (ADB) described in Table 1.
2. Prepare two Coplin jars containing 50 mL of wash buffer (WB) solution (10% ADB diluted in PBS), and one Coplin jar containing 50 mL WB and 0.05% detergent solution (e.g., add 250 µL of 10% detergent to 50 mL WB).
3. Wash the slides that were prepared in section 1 and 5 of this protocol for 10 min in one Coplin jar containing 50 mL of WB.
   Caution: Do not let the slides dry at any point during immunolabelling.
4. Wash the slides for 10 min in the Coplin jar containing 50 mL WB and 0.05% detergent solution. Then, wash the slides in the remaining Coplin jar containing 50 mL of WB solution for 10 min.
5. Tap off excess liquid and cover slide with 100 µL of selected primary antibodies diluted in ADB. Incubate at 4 °C overnight in a closed humid chamber.
   NOTE: Incubation can be shortened to 2 to 3 hrs at 37 °C. Use smaller volumes of antibody (e.g., 50 µL) by covering the slide with a coverslip or parafilm.
6. Rinse slides in a Coplin jar containing 50 mL of 0.2% wetting agent-PBS solution, pH 8.0.
7. Repeat steps 6.2 to 6.5.
8. Tap off excess liquid and cover slide with 100 µL of selected secondary antibodies diluted in ADB. Incubate slides 1 to 2 hrs at 37 °C in a closed humid chamber.
9. Wash slides 2 times for 10 min in Coplin jars containing 50 mL of 0.2% wetting agent-PBS solution, pH 8.0.
10. Wash slides 2 times for 5 min in Coplin jars containing 50 mL of 0.2% wetting agent-H₂O solution, pH 8.0.

7. Mounting slides

1. For prophase chromatin spreads prepared in step 2, add 100 µL of mounting medium containing 4', 6-diamidino-2-phenylindole (DAPI, 1.5µg/mL). For MI and MII chromatin spreads prepared in step 5, add 50 µL of mounting medium containing DAPI (1.5µg/mL). Gently blot away excess liquid.
2. Place a 22 mm x 60 mm coverslip on top and seal with clear nail polish. Store in a slide box at 4 °C or -20 °C until assessment via fluorescence microscopy.
   NOTE: Tap lightly on cover slip to rid of excess mounting medium before sealing with nail polish.