Culture and Co-Culture of Mouse Ovaries and Ovarian Follicles

All animal work was performed in accordance with institutional guidelines under license by the UK Home Office (project license number PPL 60/1726 and 60/4026).

Note: House animals in accordance with UK legal requirements in a 14 hr light and 10 hr dark photoperiod. Conduct experiments on animals using wild type C57Bl6J mice, Tau-GFP mice that have ubiquitous expression of green fluorescent protein (GFP)⁸, and Thy1-YFP mice with occasional expression of yellow fluorescent protein (YFP) in a subset of neuronal cells⁹: both transgenic lines were bred on a C57Bl6J background.

1. Working Conditions and Preparation of Instruments

1. Perform all media preparation, tissue dissections, and culture work in a laminar flow hood to ensure sterility: this avoids the requirement of the addition of antibiotics to media.
2. Always allow media/culture plates/embryo dishes to equilibrate for at least 1 hr, in a 37 °C oven (dissection medium/embryo dishes) or 37 °C, 5% CO₂ incubator (culture medium and plates), prior to use.
3. Soak glass pipettes in 0.1% solution of bovine serum albumin (BSA) for around an hour, and then leave to dry. Pull pipettes in the flame of a Bunsen burner, bending the glass as you do so, to produce finely drawn curved glass pipettes. After the pipette is pulled, make a clean cut with a glass cutter. Oven sterilize at 160 °C for around 45 min.
   NOTE: A store of these glass pipettes will be needed to transfer ovaries and follicles.

2. Preparation of Dissection and Culture Media

1. Prepare Dissection Medium.
   1. Dissolve 3 mg/ml BSA in Leibowitz L15 medium and filter sterilize through a 0.2 µm pore, 25 mm diameter filter.
2. Neonatal Ovary Culture.
   1. To prepare medium for neonatal ovary culture, make 1 ml of medium for each ovary that will be cultured. Dissolve 3 mg/ml BSA in α-Minimal Essential Media (αMEM) and filter sterilize through a 0.2 µm pore, 13 mm diameter filter into a sterile tube.
   2. To prepare plates for neonatal ovary culture, add 1 ml of neonatal ovary culture medium into each well (one ovary per well) of a 24-well plate. Using sterile watchmaker forceps, place one polycarbonate membrane on top of the medium in each well, shiny surface up. UV sterilize membranes before use.
3. Follicle Culture.
   1. To prepare medium for follicle culture, supplement α-Minimal Essential Media with 1 IU/ml recombinant human follicle stimulating hormone, 5 μg/ml ascorbic acid and 5% v/v serum obtained from adult female mice. Filter sterilize through a 0.2 µm pore, 13 mm diameter filter into a sterile tube.
   2. Filter sterilize silicon oil through a 0.45 µm pore, 25 mm diameter filter, into a sterile tube.
   3. To prepare plates, place 30 μl droplets of follicle culture medium into each well of a non-tissue culture treated 96-well microtiter round-well plate, using only the top row of each plate (to allow follicles to be moved into new rows each day over the culture period). Carefully overlay medium with 70 μl of sterilized silicon oil, to prevent medium evaporation.
4. Follicle-ovary Co-culture.
   1. Prepare medium for follicle-ovary co-culture as for follicle culture in step 2.3.1 above, making up 1 ml of medium for each follicle-ovary co-culture being set up.
   2. To prepare plates, add 1 ml of medium into each well of a 24-well plate. Using sterile watchmaker forceps, place one Nucleopore membrane on top of the medium in each well, shiny surface up. UV sterilise membranes before use.

3. Neonatal Ovary Dissection and Culture

1. Place 1 ml of dissection medium into each sterile glass embryo dish.
2. Cull neonatal mouse pups aged between postnatal day 0 and 5, culling by decapitation according to UK Home Office regulations.
3. Grasp the skin covering the abdominal wall using a pair of fine dissection forceps and make a large incision in the skin and body wall. Pull open the incision so the entire abdomen is exposed. The bladder is usually engorged at this stage and can be punctured to make dissection easier.
4. Move the guts out of the way using watchmaker forceps. Follow the uterine horns from the bladder up to the kidney on each side. The ovary is located just below the kidney at the top of the uterus and will appear as a cloud-like structure under a dissecting microscope.
5. Grasp the ovary gently with watchmaker forceps, and use scissors to sever its attachment to the uterus. Transfer the pair of ovaries into embryo dishes containing pre-warmed dissection medium.
6. Carry out fine dissection of ovaries under a dissection microscope on a heated stage (37 °C). Use insulin needles to trim away the bursal sac and any excess material including the fallopian tube, until only the ovary remains.
7. Transfer each ovary into the well of a culture plate prepared in step 2.2.2 above, using a finely drawn curved glass pipette, one ovary on top of each membrane (see Figure 1A). Culture in a 37 °C, 5% CO₂ incubator.
8. Change the medium every second day. Use a pipette to exchange 50% of the medium in each well for pre-gassed fresh medium: place the pipette tip at the edge of the well of medium to avoid disturbing the membrane.
9. Maintain cultures for up to 6 days.
10. At the end of the culture, freeze medium and fix or freeze ovaries after a brief wash in PBS, as required.
    1. Fix ovaries in Bouin’s fixative for 90 min for histological analysis, or in 10% neutral buffered formalin for 90 min for immunohistological analysis. Snap freeze ovaries by placing tissue in a polypropylene tube, placing the tube in dry ice for 5-10 min before freezing at -70 °C for subsequent protein or mRNA extraction.

4. Follicle Dissection and Culture

1. Cull 19-23 day old female mice and isolate the ovaries. Wet the fur with 70% ethanol before making any incisions. Pinch the skin using blunt ended forceps and make a large incision in the abdomen with dissection scissors, penetrating through both the skin and the body wall.
2. Move the gut out of the way using a finer set of dissection instruments and locate the uterus. Follow the uterus up to the ovary which is located below the kidney on each side.
3. Gently grasping the ovary using a pair of watchmaker forceps, sever the ovaries attachment to the uterus with fine dissection scissors. Avoid collecting too much of the ovarian fat pad. Collect the ovaries and transfer into an embryo dishes containing pre-warmed dissection medium.
4. Remove the ovarian bursa using insulin needles to trim away the bursal sac and any excess material including the fallopian tube, until only the ovary remains. Roughly halve each ovary using insulin needles.
5. Carefully transfer each ovary-half into an individual watch glass containing 1 ml dissection medium. Cover each watch glass with a glass slide to prevent evaporation of medium and to ensure sterility.
6. Store watch glasses containing ovary halves in a 37 °C oven until needed, but carry out the next dissection step as soon as possible. Discard tissue stored in this way for more than an hour.
7. Transfer watch glass containing an ovary half to a 37 °C heated microscope stage in a laminar flow hood. Roughly dissect ovary into large pieces using two insulin needles, in order to identify late pre-antral follicles as follows: these will contain 2-3 layers of granulosa cells and have a diameter of around 180-200 µm.
8. Manually dissect out any identified follicles using one insulin needle and one 30 x 0.25 mm acupuncture needle that has been secured in a needle holder. Be careful to remove most of the surrounding stroma, but avoid damaging the basal lamina of follicles (see Figure 1B).
9. Use a finely drawn curved glass pipette to carefully transfer dissected follicles into a collecting watch glass containing pre-warmed dissection medium. Be careful to keep follicles within the thin caliber section of pipette to avoid losing follicles.
10. Measure follicles accurately using a calibrated eyepiece graticule fitted into a dissecting microscope.
11. Select follicles for culture only if they measure 190 ± 10 µm in diameter. Further select only healthy, spherical follicles for culture; these will be translucent, without dark atretic areas, and have an intact basal lamina, along with some attached thecal tissue: discard any follicles that do not fit this description. A yield of between 10-15 follicles per ovary is good.
12. Use a finely drawn curved glass pipette to transfer a single follicle into the well of a plate made up as in step 2.3.3 above. Carefully place the follicle in the bottom of the well (and not in the upper oil layer). Culture in a 37 °C, 5% CO₂ incubator.
13. Culture follicles for up to 6 days, moving follicles into a well containing fresh medium every day.
    1. Prepare the next row in the 96-well plate as in plate preparation in step 2.3.3 above. Return plate to the incubator for at least an hour, to allow equilibration. Transfer follicles into fresh wells of medium, using a finely drawn glass pipette.
    2. If at any point in the culture follicles are to be left for two days before moving into fresh medium, place each follicle in a minimum of 60 μl (rather than 30 μl) droplets of follicle culture medium.
14. To collect data on follicle growth, measure follicles daily, using an eyepiece graticule fitted into a dissecting microscope. The oil layer will distort measurements of follicle diameter, so work out the calibration coefficient for the set-up.
15. At the end of the culture, freeze medium and fix or freeze tissue for subsequent analysis, as in step 3.10 above.
16. Follicle-follicle Co-culture.
    1. Culture two follicles together, to investigate interactions between follicles. Culture as above, but place two follicles side-by-side, in contact, in a well.
    2. Place 100 μl droplets of follicle culture medium into each well of a non-tissue culture treated 96-well microtiter flat-well plate, overlaying medium with 100 μl of sterilized silicon oil. Do not transfer follicles into fresh wells as in step 4.13 above, but instead use a fine gel tip to change 50% of the medium every other day.
    3. In order to identify tissue origins within the co-culture, co-culture follicles each from a different genetic source, for example one from the ovary of a wild type mouse, and the other from the ovary of a mouse with ubiquitous expression of GFP. If using GFP or YFP tissue, minimize exposure of tissue to light as much as possible, during culture, and through the fixation/processing steps thereafter.
       NOTE: It is normal for the two follicles to grow together into a single, ‘two-follicle’ unit (see Figure 1C).

5. Follicle-Ovary Co-Cultures

1. Dissect ovaries and follicles out as in Sections 3 and 4 above.
2. Place one neonatal ovary on top of a membrane, in a plate prepared as in step 2.4.2 above. Carefully place a single follicle in contact with one pole of the neonatal ovary, using a finely drawn curved glass pipette. Culture in a 37 °C, 5% CO₂ incubator for up to 5 days.
3. In order to distinguish between the tissue origin within the co-culture, use ovaries and follicles from two different sources, for example one from a wild type mouse, and one from a mouse with ubiquitous expression of GFP.
4. Replace 500 μl of medium daily as in step 3.8 above, but using follicle culture medium. During co-culture, the follicle often becomes encapsulated by the ovary (Figure 1D).
5. At the end of the culture, freeze medium and fix or freeze tissue for subsequent analysis, as in step 3.10 above.

6. Fixation, Immunocytochemistry and Imaging of Cultured Tissue

1. At the end of culture, wash tissue in PBS and transfer to 100 μl (follicle) or 1 ml (ovary) of 10% neutral buffered formalin, and fix for 1 hr on ice. Move through 3 washes of 1x PBS at 4 °C.
2. For immunocytochemistry on follicles, transfer between wells of a 96-well microtiter round-well plate using a finely drawn curved glass pipette, with each well containing different washes or treatments.
3. For immunocytochemistry on ovaries (or ovary-follicle co-cultures), embed in 4% agarose gel and section at 50 µm using a vibrotome. Float sections in the wells of a 24-well plate to apply washes or treatments, removing with a pipette.
4. For imaging: transfer agarose sections to flat glass slides and mount with mounting medium; or transfer follicles (or follicle-follicle complexes) to cavity glass slides and mount with non-hardening mounting medium. Image specimens using a confocal microscope.