Protocol For Preparing Drosophila Eggs To Primary Cell Culture
This is the protocol starting from a plate already with eggs laid
Steps
Set up
- Make 50% bleach solution
- 10mL bleach
- 10mL 1X wash
- Take out Trypsin (0.25%) and media from the fridge to warm up
- Turn on blower in the tissue culture hood and turn the UV on for ~10 minutes
- Pour a small beaker with some 70% ethanol for washing forceps and paintbrush
- If needing to make new media, do this in the TC hood after the UV has finished:
- 1 50mL conical of 10% serum media
- 5mL of serum
- 44.5mL of 420 Ex-Cell medium
- 500ul of 100X antibiotics (Penicillin, Amphotericin B, and Streptomycin)
- 50ul of 50mg/mL Gentamicin
- 2 50mL conicals of media without serum
- 49.5mL 420 Ex-Cell medium
- 500ul of 100X antibiotics (Penicillin, Amphotericin B, and Streptomycin)
- 50ul of 50mg/mL Gentamicin
Fly Room
- If doing a second day of laying, bring the second apple juice and yeast plate to the fly room
- Set up CO2: unscrew tank if not on, plug in hose and open valve
- Tap fly cage onto the CO2 plate until all flies are asleep
- Take out red cap and plate and cover the plate immediately
- If doing a second day of laying:
- Put in new yeast plate to the red part and re-cap cage
- Rotate the cage horizontally until the flies wake up
- Set up the cage for overnight
- If flies are done:
- If not keeping flies, dump them out into the morgue
- If keeping flies, separate them out into 3 equal groups on the CO2 plate and add them to newly labeled vials
- Turn off all CO2
- Take plate covered back to 4012
Filtering: At the Lab Bench
- Pick off dead flies with tweezers and put in ethanol
- Set up the filter rig: autoclaved erlenmeyer flask, 100um filter, 400um filter, then the funnel on top
- Squirt water into the plate and begin mixing up the yeast with a paint brush, mix until as best you can get homogenous
- Tip over the plate into the funnel and squirt water to wash the liquid down
- Brush and rinse at least 2 more times to wash all the yeast and eggs out of the plate
- Take off the funnel and rinse the green filter with water
- Take of the green filter and rinse the yellow with 1X wash for close to 2 minutes
- Flip the yellow filter over into a new 50mL conical
- Serologically pipette 10mL 50% bleach into the filter to wash out the eggs into the conical below
- Use a siliconized pasteur pipette (spp) to transfer the 10mL to a siliconized 10mL tube
- Let sit for 3 minutes: this sterilized and de-chorionates the eggs
- Centrifuge for 3 minutes at 400rcf: program 2 in centrifuge by the incubators
Cell Culture Preparation: In the Tissue Culture Hood
- Make sure trypsin and media are at room temp
- Set up vacuum: attach tubing to the hood and to the vac in the chemical hood. Turn on the vac ~half a turn
- Use an spp and the vacuum to aspirate off the bleach from the 10mL tube, avoid the pellet
- Add 10mL trypsin to the 10mL tube and invert
- Centrifuge 3 min 400rcf
- Aspirate off the liquid with an spp
- Add 10mL trypsin and invert
- Centrifuge 3 min 400rcf
- Aspirate off the liquid with an spp
- Add 10mL trypsin and invert (3rd wash)
- Centrifuge 3 min 400rcf
- Aspirate off the liquid with an spp to about ~200ul
- Add back in 2mL fresh trypsin
- Unwrap dounce mortar and put in a separate rack
- Use an spp and a blub to pipette mix once and transfer all the liquid from the 10mL tube to the dounce
- Unwrap the pestle from the foil and homogenize for ~30 seconds, press down and twist, then release up, about 3 times. Liquid should become cloudy
- Lay the pestle back down on the clean autoclaved foil
- Let the dounce with the sample sit for 30 min in the hood
- Every ~5-10 minutes, take the pestle and put it in the dounce and lift up (no pressure) once just to mix the liquid
- After the incubation: use and spp to transfer the liquid from the dounce into a new siliconized 10mL glass tube
- Add 8mL 420 medium without serum to the 10mL tube and invert
- Centrifuge 800rcf for 3 minutes (program 1)
- Aspirate off the liquid with an spp
- Added 10mL 420 medium without serum and invert
- Centrifuged 800rcf for 3 minutes
- Aspirate off the liquid with an spp
- Added 10mL medium with serum to the 10mL tube
- Made two new 25cm2 flasks
- Label with fly line name, P1 primary, the date, and your initials
- Use a 2mL serological pipette (10mL does not fit inside the glass tube) to transfer the 10mL into a single tissue culture flask. Pipette carefully to avoid any bubbles or liquid on the sides of the flask
- Transfer 5mL from the first flask into the empty flask. Pipette mix a few times to mix the liquid and apply similar populations to each flask
- Cap and lay flasks on their sides, adjust liquid so that it covers the whole side of the flask
- Place flasks in the 23 degree incubator. Let them settle for a few hours before looking at under the scope
Cleanup
- Put all media and trypsin back into the cell culture fridge
- Put all used pasture pipettes in the glass disposal box
- Rinse out the plastic beaker for used pipettes with tap, then distilled water, then spray with ethanol and leave to dry in the TC room
- Rinse out the dounce mortar and pestle with tap then distilled water. Leave to dry for a few hours then wrap in foil for autoclaving. Wrap so that when unwrapping, the mortar comes out first, then the pestle
- Rinse out the 10mL tubes with tap, then distilled water. If there are eggs stuck on the sides, add a small droplet of detergent, shake up the tube to mix, then rinse many times. Let dry for a few hours then loosely cap, and tape for autoclaving
- Rinse out flask for filtering with tap, then distilled water, dry for a few hours, then foil for autoclaving
- Rinse out the beaker with ethanol with tap, then distilled water, dry for a few hours, then foil for autoclaving
- Clean up all wrappers of serological pipettes
- Turn off and uh-plug the vacuum setup
- Spray a paper towel with ethanol and wipe down the TC hood workspace
- Autoclave everything so that it is ready for the next day