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

The overall goal of this chromatin spread preparation is to chronologically visualize the dynamic localization patterns of chromatin-bound proteins and chromosome morphology of oocytes throughout mammalian oogenesis. This method can help answer key questions in the field of female reproductive biology, such as homologous chromosome pairing, synapsis, DNA repair, and meiotic chromosome segregation. The main advantage of this technique is that it allows for clear immunolabeling and visualization of proteins associated with chromosomes and assessment of oocyte ploidy.

Oogenesis is error-prone, and chromosome missegregation often results in genetic disease. This method can be used to improve our understanding of maternal causes of aneuploidy and infertility. After euthanizing a female mouse that is 14 to 19 days post-coitum, make a V-shaped opening into the abdominopelvic cavity.

Then remove the maternal uterine horn. Next separate the embryos from the placenta, and sacrifice the embryos by decapitation. Then transfer them to 35-millimeter Petri dish containing three milliliters of PBS at 37 degrees Celsius.

Then move one pup to a separate dish also loaded with warm PBS. After euthanizing the fetus, make a cut along the ventral midline of the posterior half, along the anterior half below the forelimbs, and directly above the hindlimbs and tail. Next open the embryo’s abdomen using 3.5-inch scissors.

Using fine-tipped forceps, displace or remove the liver and loops of bowel, exposing the gonads. Now determine if the gonads are male or female gonads. The ovaries are located immediately below and behind the kidney, toward the posterior wall of the peritoneal cavity.

Remove both ovaries using a pair of fine-tipped forceps, and transfer them into a small container containing PBS that is maintained at 37 degrees Celsius. Proceed quickly to collect the ovaries from all of the fetuses. Next in a new small container, completely immerse each separate set of ovaries in half a milliliter of freshly made hypo-extraction buffer.

Let them incubate at 25 degrees Celsius for at least 15 minutes, but not more than 30 minutes. During the incubation, use a hydrophobic barrier PAP pen to draw two 22 by 22 millimeter squares on a clean glass slide. Prepare one slide for each set of ovaries.

When the incubation is complete, add a 50-microliter drop of 100 millimolar sucrose to a clean slide, and transfer one pair of ovaries into the drop. Now use two 27-gauge needles to tease the ovaries and release the cells therein. With forceps, remove and discard the large pieces of ovary.

Then carefully pipette the sucrose solution to disperse the released cells. Next load each square on a slide with 40 microliters of 1%PFA with 0.2%Triton X-100. Spread the solution around the square using a pipette tip or by rocking the slide back and forth.

Then for each pool of sucrose solution with cells, load two fixative puddles with 20 microliters of suspended cells so that each slide has cells from one set of ovaries. For either MI or MII oocyte collections, first prepare and euthanize the donor mouse. Then make a large V-shaped opening in the abdominopelvic cavity, and use forceps to displace the intestines to locate each uterine horn.

Find the ovaries proximal to the rib cage. Hold the oviduct with fine forceps, and cut the fat superior to the ovary using dissection scissors. While still holding the oviduct, use fine forceps to release the ovary from the bursa, and transfer the ovaries into a collection dish containing medium maintained at 37 degrees Celsius.

To collect GV-stage oocytes, use a one-milliliter syringe with a 27-gauge needle to release the cumulus-oocyte complexes by manually puncturing the large antral follicles. To collect MII stage oocytes, tear a hole in the ampulla of the oviduct to release them rather than simply making a puncture. Now collect the oocytes using a mouth-operated glass pipette or capillary, or use a hand-operated micrometer syringe.

Some oocytes may be surrounded with cumulus cells, others not. For the MI oocytes, prepare fresh hyaluronidase in MEM-alpha/BSA to denude the oocytes of the surrounding cumulus cells. For each collection of oocytes, treat them in an incubator with 2.5 milliliters of prepared hyaluronidase for three minutes.

Then transfer the MI oocytes to freshly made warm MEM-alpha/BSA medium. There, use a mouth-operated glass pipette to provide the shearing force needed to completely detach the cumulus cells. In general, the success of this procedure requires the mastery of oocyte collection and manipulation using mouth-operated glass pipettes or capillaries.

Allow the oocytes to recover in the incubator. Meanwhile, into a heated nine-well glass plate, load 300 microliters of warm Tyrode’s solution. Load two wells with 300 microliters of warm MEM-alpha/BSA, and load one well with 300 microliters of warm Waymouth’s medium.

To remove the zona pellucida, transfer five to 10 MI or MII-stage oocytes into the bath of Tyrode’s solution for just 30 to 45 seconds while watching them under a dissection microscope. When the zona pellucida is visibly dissolved, immediately transfer the oocytes to the medium using a pipette pre-wet with MEM-alpha/BSA medium. Then transfer the oocytes to a second bath of warmed medium.

After the second bath, transfer the oocytes to warm Waymouth’s medium, and let them recover in an incubator for 30 minutes. To prepare for a chromatin spread, use a PAP pen to draw an 11 by 44 millimeter rectangle on a glass slide. Next coat the slide with a thin layer of 1%PFA with 0.2%Triton X-100 in water, and then rock the slide back and forth to spread the solution.

Then tap the slide to remove the excess. Now siphon between five and 10 oocytes with a minimal amount of medium. Then transfer the oocytes to the prepared slide by simultaneously dragging a pipette tip through the fixative solution and dropping the oocytes onto the slide from about one centimeter above.

The oocytes should burst immediately, spreading chromatin on the slide. Now incubate slides at room temperature in a closed humidified chamber. Let them settle overnight so the chromatin adheres to the slide.

The described embryonic chromatin collection was used to quantitatively analyze crossover formation in oocytes harvested from C57 black/6J embryos. Prophase chromatin spread preparations were immunolabeled to detect SYCP3, SYCP1, and MLH1, and stained with DAPI. Pachytene-stage chromosomes had 27 MLH1 foci distributed along 20 fully assembled SC structures.

When the spread preparation was suboptimal and individual chromosomes were indistinguishable, the preparation was not included in the assessment. In good preparations, SMC6 was enriched at the pericentromeric heterochromatin region and along the chromosome arms, and the MLH1 foci distributed along the SC at the pachytene stage. The number of MLH1-positive crossovers from 15 good wild-type preparations was about 24.

The oocyte collection method was used to assess chromatin morphology in oocytes following meiotic resumption. Protein localization, as well as ploidy, was easily assessed, exposing potential causes of chromosome segregation errors. Topoisomerase II-alpha could be seen along the chromosome arms and the pericentromeric heterochromatin.

In MII oocytes, paired sister chromatids could be distinguished by chromosome and centromere morphology. Once mastered, these techniques without or after culturing can be completed within one or two hours per mouse. Following this procedure, other complementary methods, like whole cell oocyte immunolabeling and live cell imaging can be performed to assess homologous chromosomes and sister chromatids within the context of the cell.

Collectively, chromatin spread methods described here can be easily applied to assess mammalian oogenesis and the sexually dimorphic features of chromosome and protein dynamics during meiosis.