Metaphase spreads of human oocytes

This protocol outlines the generation of metaphase spreads of a single oocyte at either the immature metaphase I (MI) or mature metaphase II (MII) stage. This protocol utilizes formaldehyde xation and therefore allows for further immunouorescence analysis. These spreads can also be used to identify aneuploidy and other large-scale structural chromosomal abnormalities. However, this assay is prone to false positives for whole chromosome aneuploidy losses, due to either xation failure or excessive spreading leading to an inability to nd all the chromosomes. Additionally, success rates can be low using human oocytes compared to similar protocols for mouse oocytes due to diculty bursting the oocyte membrane and providing sucient dispersion of the chromosomes for individual analysis. The protocol duration is two days, however, immunouorescence staining will add to the total time before nal analysis.


Introduction
Metaphase spreads are an excellent way of xing the DNA to allow for direct visualization of compacted chromosomes using many different downstream methods, including immuno uorescence and uorescence in situ hybridization. This method works for immature MI oocytes and mature MII oocytes from both in vivo and in vitro matured large antral follicles collected during in vitro fertilization (IVF) procedures, as well as with in vitro matured small antral follicles (Gruhn et al, Science 2019). This protocol can also be used for mouse oocytes (as described in Sankar 2020 Solutions & preparation of materials: 1. 1% formaldehyde solution is made freshly from paraformaldehyde (PFA). Note that PFA is a polymer of formaldehyde that must be hydrolyzed to be functional. This is done by heating the aqueous solution (60°C) and increasing the pH with NaOH. a. Add 0.25 g of PFA to 22.5 ml of double-distilled water in a 50 ml conic tube. Add one drop of 1 M NaOH. Incubate in 60°C water bath for at least 20 minutes. Invert every 5 minutes or so to ensure all PFA is dissolved.
b. Allow solution to cool to room temperature before adjusting the pH to 9.2 using 50mM boric acid (usually takes about 0.5 ml).
c. Add 35 µl of Triton X-100 (0.15% nal v/v). Make sure is completely dissolved before use. Triton X-100 is a nonionic detergent that bursts the nuclear envelope. This is done concomitantly with the formaldehyde xation step. 4. Prepare a humid chamber with hot water before beginning protocol.
5. Etch and clean microscope slides. Use a diamond pen to draw a circle in the middle on the underside of the slide where you will drop the oocyte. Use ethanol or IMS to wipe the slides to remove grease using standard laboratory tissue wipes. Procedure: The following steps are carried out under the stereomicroscope on a heated stage.
6. Place the slides in a Coplin jar containing in 1% formaldehyde solution to coat the slide. 11. While looking under the stereomicroscope, drop the oocyte onto the formaldehyde-soaked slide within the etched circle, transferring the least amount of wash solution possible. Watch for the oocyte to visibly burst when it hits the slide. The oocyte will look to atten and 'crack open', followed by a visible dispersion of the cytoplasm. Once completely burst the oocyte will be nearly invisible on the slide.
12. Taking care to keep the slide absolutely at, transfer it to the pre-warmed humid chamber (37 °C) for gradual drying for a minimum of two hours at room temperature.
13. After slow drying, crack the lid of the humid chamber and leave until slides are fully dry at room temperature (approx. 20 minutes).
14. Move slides to a Coplin jar containing a 1% Photo o solution (500 µL in 50 mL of water) and wash for 2 minutes.
15. Remove slides and air dry. Slides may dry standing vertically against any surface to allow excess liquid to run to the end of the slide or lying at on paper towels. Leave at room temperature on the bench until completely dry (approx. 10-20 min).

Troubleshooting
Step 1: The nal 1% formaldehyde solution can be stored at 4°C. However, the PFA will eventually fall out of solution and change the nal concentration; therefore, discard after two weeks to avoid suboptimal xation.
Step 2: Once a Tyrode's solution aliquot has been thawed, refreezing and thawing will cause the e ciency of the solution to decrease. Make small enough aliquots (~500 µl) from the stock bottle to allow for limited refreezing.
Step 6: Low humidity can impact the speed of formaldehyde evaporation and can decrease the quality of the spreads. If you work in a dry environment use a humidi er in the room.
Step 7: The zona will usually slough off with mechanical movement of the oocyte. However, sometimes the Tyrode's will simply thin the zona until it is not visible, but will still stop the oocyte from properly bursting on the slide. In this case, for MII oocytes it is easiest to see any residual zona by looking at where the zona is pulled away from the oocyte by the polar body, or simply by the act of the polar body separating from the oocyte. With MI oocytes this is much more di cult; therefore, leave the oocyte in the Tyrode's solution for approx. 10 extra seconds if no visible zona has separated from the oocyte. When spreading oocytes expect to have relatively low e ciency. Our rates from spreading to analysis was slightly more than 25%.
Step 8: If the oocyte is left in Tyrode's for too long, it will either burst in the Tyrode's or immediately upon transfer to the sodium citrate wash drops. To limit time in Tyrode's, the oocyte can be moved to the sodium citrate drops before the zona has completely dissociated from the oocyte. The oocyte should then be pipetted up and down to completely remove the zona by the time the cell is in the nal wash drop. At this point the oocytes will be extremely sticky, and therefore, when in the sodium citrate drops try to continuously pipette the oocyte so that it does not settle onto the bottom of the dish.
Step 9: At this stage hold the oocyte in your pipette before retrieving your slide to again avoid the oocyte settling, and sticking, to the bottom of your dish. Minimize the amount of liquid transferred to limit the dilution of the formaldehyde solution coating the slide. If too much liquid is transferred, this can lead to a poor burst and poor xation.
Step 10: When removing excess formaldehyde from the slide, only a quick dab on the end of the slide is needed. However, it is also important to wipe the back of the slide. If the back is not wiped, the slide wiill suction onto the stage and makes it di cult to quickly move the slide to the humid chamber without tipping it.
Step 11: It is important to limit the amount of media that is transferred, as it will dilute the formaldehyde solution on the slide and impact xation. If the oocyte does not burst, agitate the liquid surrounding the cell with the pipette tip (careful to not aspirate any liquid) until you see the cell start to lyse. However, do not agitate too much as it may cause the chromosomes to spread excessively on the slide and cause unintended chromosomal loss.
Step 12: Slides should be slow dried in the humid chamber for a minimum of 2 hours. We have also found no difference if the slides are left to dry overnight. However, try to not leave the slides for more than 24 hours as the quality of protein staining may decrease.
Step 14: The Photo o wash is to remove any excess debris from the slide. It is best to complete this step prior to storage, however, the Photo o wash can also be completed after thawing prior to staining.
Time Taken

Anticipated Results
For a successful metaphase spread, the chromosomes must a x to the slide and separate enough to allow for individual identi cation. Due to variation in each individual oocyte (e.g. MI vs. MII, thickness of the zona, quality and fragility of the cytoplasmic membrane, etc.) the outcomes will greatly vary and can decrease the effectiveness of the spreading technique to approximately 25% of samples producing an analyzable spread.