Developmental Biology
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Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries
Chapters
Summary May 2nd, 2018
Here, we present a protocol to quantify follicles in cultured ovaries of young mice without serial sectioning. Using whole organ immunofluorescence and tissue clearing, physical sectioning is replaced with optical sectioning. This method of sample preparation and visualization maintains organ integrity and facilitates automated quantification of specific cells.
Transcript
The overall goal of this procedure is to obtain images of cultured whole ovaries that are compatible with automated follicle quantification. This method can help answer key questions in the reproductive biology field such as how different conditions interfere with ovarian fitness. The main advantage of this technique is that it is less labor intensive and maintains the organ's three-dimensional architecture.
Though this method has been optimized for cultured mouse ovaries at postnatal day five, it can also be applied to other ages and tissues such as embryonic gonads or young mice testes. Start with a clean work surface. Wipe with ten percent bleach and allow the bleach to remain on the work area surface for at least five minutes.
After five minutes, remove excess bleach with clean paper towels and 70 percent ethanol. Clean the dissecting microscope thoroughly with 70 percent ethanol and place a dry paper towel on the stage. Wrap clean forceps and scissors with a paper towel dampened with 70 percent ethanol.
Then make 50 milliliters of ovary culture media in a conical tube and warm in a 37 degrees Celsius water bath. After the medium has warmed, prepare plates and inserts by first adding one milliliter of ovary culture medium to 35 millimeter tissue culture plates. Then add approximately 0.55 milliliters of medium to the wells of a 24-well tissue culture plate and place an insert into each well containing medium.
Make sure that the membranes touch the media. Place the 35 millimeter plates and the 24-well carrier plate into a 37 degrees Celsius, five percent carbon dioxide and atmospheric oxygen incubator. Then place a hot plate next to the dissection scope and set the temperature to 37 degrees Celsius.
If the tissue culture incubator is not close to the dissection area, place one of the 35 millimeter tissue culture dishes containing ovary culture media on the hot plate. Then begin the dissection by spraying the freshly euthanized female with 70 percent ethanol and place on top of the dry paper towel under the dissection microscope. After making an incision in the abdominal cavity, extrude the intestines and locate the ovaries.
Extract the ovaries and place them into the culture medium in the 35 millimeter dish. Once the ovaries have been dissected, increase the magnification of the dissection microscope to better visualize ovaries and any attached tissue. With clean fine tip forceps, remove all non-ovarian tissue such as bursa and fatty tissue without damaging the ovaries.
Next, place each pair of ovaries onto a pre-soaked cell culture insert and adjust the volume of the medium to ensure that it is sufficient to keep the organ moist without completely submerging it. Place explanted organs in a 37 degrees Celsius, five percent carbon dioxide and atmospheric oxygen incubator. At the desired experimental time point, remove the ovary culture media from the plate and fix the cultured ovaries on the insert by covering the tissue with freshly prepared four percent PFA solution.
Seal the edges of the plate with paraffin film to avoid evaporation and place the samples at four degrees Celsius overnight. Rinse the tissue three times with 70 percent ethanol. Then store in scintillation vials filled with 70 percent ethanol at four degrees Celsius until further processing.
Begin staining by removing the 70 percent ethanol from the vials and washing the fixed tissue with PBS at room temperature with shaking for a minimum of four hours. After removing the PBS, add enough permeabilization solution to completely submerge the ovaries. Place on an orbital shaker and incubate at room temperature for four hours.
Replace the permeabilization solution with enough blocking solution to completely submerge the ovaries. Then incubate the sealed vials at room temperature for a minimum of 12 hours on an orbital shaker. After blocking, place the inserts into a 24-well plate and add about 750 microliters of primary antibody solution ensuring that ovaries are completely submerged.
Then place the sealed plate on the orbital shaker and incubate for four days at room temperature. After four days, remove the primary antibody and add a generous amount of washing solution. Wash the ovaries overnight.
The next day, replace with fresh washing solution and incubate on the orbital shaker for two hours or more. After the washes, add secondary antibody solution. Protect the samples from light and incubate on an orbital shaker at room temperature for two days.
After two days, remove the secondary antibody solution from the samples and add washing solution. Begin by preparing ScaleS0 clearing solution. Mix the solution by inversion and incubate at 50 degrees Celsius for 30 minutes.
De-gas the ScaleS0 clearing solution. Then remove the washing solution from the immunostained ovaries. Add the clearing solution.
Cover the plate with aluminum foil and return it to the orbital shaker. It is extremely important to be patient during the staining and clearing steps. Once the tissue has become transparent, use a fine tip scalpel to carefully remove the insert membranes containing the cultured ovaries.
Place the sample on a glass slide and gently cover the sample with a coverslip. Then proceed to image the samples on a microscope capable of optical sectioning. This image shows a postnatal day five ovary which has not been cleared.
Here, the ovary is shown one hour after the addition of clearing solution. The ovary will begin to become translucent within minutes of being submerged in clearing solution. Optical sectioning of cultured ovaries without clearing is possible, but cells deep within the tissue have a signal that is difficult to differentiate from background, which impedes proper oocyte quantification.
In contrast, this representative image shows a cleared sample in which oocyte quantification throughout the entire organ is possible. While attempting this procedure, it's important to remember that antibodies have to diffuse through the tissue;thus, thicker tissues will require longer incubation time. After watching this video, you should have a good understanding of how to culture, stain, and image whole ovaries.
The text describes the use of a free software, Fiji ImageJ, that can be used for simple follicle quantification. Don't forget that working with paraformaldehyde and sodium azide can be hazardous and precautions such as use of fume hoods and personal protective equipment should always be taken while performing this procedure.
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