Generating Primary Cells from D. innubila Dissected Ovaries

This process generally followed the same method as the previous ovary primary cell generation with some differences.

  • Isolated out 100 females into empty vials, 5 flies per vial about an hour before starting so the would hopefully poop out much in their guts
    • Females were age “4”
  • Prepared:
    • a small beaker with room temp Schneider’s medium
    • a dish with 70% ethanol
    • a dish with DI-water
    • a 15mL tube with room temp Schneider’s medium and a transfer pipette
  • Used microscope slides and dissecting forceps
  • Steps of each dissection:
    • Put a drop of medium on the microscope stage
    • Placed a blank glass slide on the water drop
    • Added a drop of medium to one side of the slide
    • Put one vial of flies to sleep on the CO2 pad
    • Took a fly with forceps and dipped it into 70% ethanol, and then into the DI water
    • Placed the fly on the droplet of ringers on the slide
    • Use the forceps to try to pull off some tergets on the abdomen of the fly while holding the thorax with the other forcep
    • Tried as best as possible to squeeze out the ovaries and not break the gut
      • This was tough, many flies just fell apart as I dissected, or had small ovaries that I had to dig through the gut to get to
    • Picked up the isolated ovaries and put time into the beaker of Schneider’s medium
    • Dipped the forceps in 70% ethanol
  • This was repeated for ~ 75 flies, I’m not sure how many I got through but it wasn’t all 100. This took me about 2 hours at least
  • Then the beaker with the ovaries was taken to the cell culture hood in 4012
  • There I had fly extract, mushroom extract, and mediums warmed to room temperature
  • Put a mesh 100um cell strainer in an autoclaved flask and poured the ovaries in through the mesh. To get any stuck on the beaker I used DI water to wash them into the strainer
  • Squirted 70% ethanol over the strained ovaries for about 1 minute to try to sterilize them
  • Washed the ovaries out into a 50mL conical with 7mL of 10% FBS Schneider’s medium
  • Put that liquid into a 10mL glass tube and let the ovaries settle
  • This time they didn’t fully settle on their own, so I centrifuged them at 400rpm for 3 min (still not all settled but it was still easy to pipette out the liquid)
  • Removed all the liquid from the 10mL tube
  • Washed the ovaries again with 7mL of 0% FBS Schneider’s medium and centrifuged again
  • Removed all the liquid from the 10mL tube
  • Added 3mL of 10% FBS Schneider’s medium
  • Transferred the liquid and ovaries to the 7mL dounce homogenizer
  • Homogenized the ovaries as thoroughly as possible
  • Added 5mL of 10% FBS medium to the dounce and pipette mixed
  • Added another 100um mesh cell strainer over a new 50mL conical and pipetted the homogenate through the strainer
  • Then I released the strained cells into a new 50mL conical with 5mL 10% FBS medium
  • This left me with a conical with filtered cell homogenate (from the top of the filter) and the flow through cell homogenate
  • I put the filtered cell homogenate in 1 T25 flask with 10% fly extract and 10% FBS Schenieder’s medium, 10mL total volume
  • I split the flow through cell homogenate in 2 T25 flasks, each with 10% fly homogenat and 10% FBS Schneider’s medium at 10mL, and one also had 4% mushroom extract
  • All flasks were put in the 23C incubator