Preparation of Fixed Drosophila Oocytes for Immunostaining: A High-Throughput Method to Fix and Remove the Outer Membrane

This protocol is excerpted from Radford and McKim, Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes, J. Vis. Exp. (2016).

NOTE: Procedures are performed at room temperature unless otherwise noted. Temperature-controlled incubators are used to maintain temperatures for fly rearing and crosses unless otherwise noted.

1. Preparations

1. Prepare Flies.
   1. Prometaphase-enriched oocyte collections.
      1. Clear adult flies from healthy, young stock or cross cultures. Age bottles for two days at 25 °C.
         NOTE: Generally, two healthy bottles will suffice, although more may be needed for some cross cultures.
      2. After two days, collect ~100 to 300 females (who are 0 to 2 days old at this point) from the bottles. The females do not need to be virgins. Add a dab of yeast paste to the side of a vial and place 30 females and 10 to 15 males each into the yeasted vials. Age vials for two days at 25 °C.
   2. Metaphase-enriched Oocyte Collections.
      1. Collect ~100 to 300 females from stock or cross cultures. Add a dab of yeast paste to the side of a vial and place 30 females each (with no males added) into the yeasted vials. Age vials for three to five days at 25 °C.
2. Prepare Solutions.
   NOTE: Solutions may be stored indefinitely at room temperature, unless otherwise noted.
   1. Prepare Modified Robb's Buffer (5x): 500 mM HEPES, 500 mM sucrose, 275 mM sodium acetate, 200 mM potassium acetate, 50 mM glucose, 6 mM magnesium chloride, and 5 mM calcium chloride. Use 10 N 11:8 sodium hydroxide:potassium hydroxide to bring pH to 7.4. Sterilize by filtration; do not autoclave. Store at -20 °C. Thaw as needed to prepare ~200 ml of 1x Robb's per oocyte prep.
   2. Fixation Solutions.
      1. Option #1: Prepare formaldehyde/heptane fixation. Prepare Fixation Buffer: 1x phosphate-buffered saline (PBS) plus 150 mM sucrose. To use, make fresh with 687.5 µl Fixation Buffer and 312.5 µl 16% formaldehyde per oocyte prep.
         CAUTION: Wear gloves while using formaldehyde solutions in a fume hood. Dispose of waste according to institutional guidelines.
      2. Option #2: Prepare formaldehyde/cacodylate fixation. Prepare Fix Mix: 250 mM sucrose, 100 mM potassium acetate (pH 7.5), 25 mM sodium acetate (pH 7.0), and 25 mM EGTA (pH 8.0). To use, make fresh with 400 µl Fix Mix, 100 µl potassium cacodylate (1 M, pH 7.2), and 500 µl 16% formaldehyde per oocyte prep.
         CAUTION: Potassium cacodylate contains Arsenic.
   3. Prepare PBS/Triton X-100: 1x PBS plus 1% or 0.05% Triton X-100. Store at 4 °C.
   4. Prepare PBS-Tween 20-Bovine Serum Albumin (BSA) (PTB): 1x PBS, 0.5% BSA (w/v), and 0.1% Tween-20. May be stored at 4 °C for one week.

2. Collection of Late-stage Drosophila Oocytes

1. As Drosophila oocytes with membranes intact will stick to plastic and glass, pre-coat the inside of one 5 ml tube and one Pasteur pipet per oocyte prep with PTB.
2. Anesthetize all ~100 to 300 yeasted flies with carbon dioxide and add to a blender containing ~100 ml 1x Robb's Buffer. Pulse three times (~1 sec each). Keep oocytes in Robb's for <20 min to avoid activation.
   NOTE: Alternatively, oocytes may be hand-dissected from females. The advantage of this method is that it requires less females. However, care must be taken to limit exposure to carbon dioxide to only a few minutes to avoid artifacts associated with hypoxia.
3. Filter through large mesh (~1,500 µm) into 250 ml beaker to remove large body parts. If many intact abdomens remain on mesh, re-grind material using additional Robb's Buffer, and filter again. Let settle ~2 min, then aspirate off top layer, removing as many of the large body parts as possible.
4. Filter through small mesh (~300 µm) into a 250 ml beaker. Rinse remaining oocytes out of first beaker using additional Robb's and coated Pasteur pipet. Let settle ~3 min; oocytes will settle out. Aspirate off all but ~10 ml.
5. Pour as much of the 10 ml as will fit into coated 5 ml tube. Let settle, remove liquid, and repeat with remainder. Rinse remaining oocytes out of beaker using additional Robb's and coated Pasteur pipet. Let settle in 5 ml tube for ~3-5 min.

3. Drug Treatments (Optional)

1. Coat a second 5 ml tube with PTB for each oocyte prep. Add appropriate solvent (control) or drug to 1 ml Robb's each for each oocyte prep (Table 1).
2. Split oocytes into second coated 5 ml tube. Let settle, remove liquid, and add 1 ml Robb's plus solvent into one tube and 1 ml Robb's plus drug into second tube. Nutate for appropriate amount of time for drug treatment (Table 1). Let settle.

4. Fixation

1. Aspirate off all liquid and immediately add 1 ml Fix.
2. Option #1: Formaldehyde/heptane fixation (Fixation Buffer plus 5% formaldehyde).
   1. Fix for 2.5 min on a nutator. Add 1 ml heptane and vortex 1 min. Let settle ~1 min.
   2. Remove all liquid, and then add 1 ml 1x PBS. Vortex 30 sec. Let settle ~1 min.
   3. Remove all liquid, and then fill tube with 1x PBS.
      NOTE: Oocytes may be used immediately or kept on the nutator for several hours at room temperature.
3. Option #2: Formaldehyde/cacodylate fixation (Fix Mix plus 8% formaldehyde and 100 mM cacodylate).
   1. Fix for 6 min on a nutator. Let settle 2 min. Remove liquid, and then fill tube with 1x PBS.
      NOTE: Oocytes may be used immediately or kept on the nutator for several hours at room temperature.

5. Removing Membranes ("Rolling")

1. Using coated Pasteur pipet, add ~500 to 1,000 oocytes to the frosted (sand-blasted) part of a glass slide. Remove all body parts and extraneous material using forceps. Do not let oocytes dry out; add 1x PBS as necessary.
2. Place a coverslip on top of the oocytes and gently "roll" oocytes until all membranes are removed (dragging the edge of the coverslip across the oocytes works best). Periodically check progress under the microscope, adding more 1x PBS as necessary. Take care as too much pressure will destroy the oocytes.