Immunostaining Drosophila Embryos

Using the multiwell plate:

Day 1:

  1. Remove the yeast from the grape plate and squirt PBT into the grape plate. Gently loosen embryos from grape plates using a paintbrush and wash extensively with PBT. (Note: PBT = PBS + 0.1% Triton X.)
  2. Using a plastic pipette, remove embryos from grape plate and place embryos in one well of mesh-bottomed plate place (aka “the staining plate”) in the big funnel. Be sure to mark genotypes and positions on lid of plate!
  3. Once all the embryos have been placed in the mesh bottomed wells, wash the embryos thoroughly with PBT from a squirt bottle. Pour some 50% bleach into square petri dish, place the staining plate in this square petri dish, and shake slowly for 6 minutes. (Note: To make new 50% bleach, mix 100mL of 100% Bleach with 100mL 1x PBS in a glass bottle.)
  4. Remove the staining plate from the bleach and place back in the big funnel. Wash the embryos thoroughly with PBT using a squirt bottle, and add about 20 mL of PBT to a square petri dish. Place the plate with the embryos in this petri dish with PBT until you’re ready for the next step.
  5. Make 10 mL of 4% formaldehyde by adding 1 mL of “10x Formaldehyde” to 9 mL of PBS. Add 15 mL of heptane to two square petri dishes in the fume hood. Dry off the staining plate with a paper towel, and place the staining plate in one of the squre dishes of heptane. Add 500uL of formaldehyde to each well (even if they don’t have embryos). Fix for 22 minutes with shaking.
  6. Blot the staining plate dry on a paper towel and place the staining plate in the new dish with the heptane.
  7. ***Add 1 mL of methanol to three empty wells. Then, add 800 uL of methanol to a well with embryos – sucking the embryos up and down in tip vigorously. I usually suck the embyros up and down about 15 times. This should make a milky slurry (“cracking” embryos)
  8. After all the embryos have been cracked, lift the staining plate out of the petri dish and blot the plate on a paper towel. Place the plate in the large funnel INSIDE THE HOOD and rinse the embryos and the plate thoroughly in methanol (using a squirt bottle).
  9. Place the plate with embryos into a new square Petri dish with 100% methanol (5 minutes or so).
  10. Remove the staining plate from the dish containing methanol, blot it dry with paper towels, place it in the large funnel OUTSIDE THE HOOD and rinse with PBT.
  11. Place the staining plate into PBT in a square petri dish. Using a large-mouthed pipette, drop some PBT onto each well of embryos. Using a P1000, transfer the embryos from your wells into a LABELED eppendorf tube (you may have to let them settle a couple times if you have LOTS of embryos in each well)
  12. Wash the embryos 3 x 15 minutes in PBT. Rotate on nutator when washing. While they’re in their last wash, make up 1 mL of 5% HIGS (heat-inactivated goat serum) per eppendorf tube. Note: To make 1 mL of 5% HIGS, add 50 uL of 100% HIGS (fridge) to 950 uL of PBT.
  13. Block for 1 hour in 500 uL of the 5% HIGS you just made. Save the remaining 5% HIGS for diluting your antibodies. Rotate on nutator when blocking.
  14. Make 500 uL of your primary antibody solution in 5% HIGS for each tube.

For example: Use 1D4 at a 1:5 dilution and use anti-beta-gal at a 1:5000 dilution

So, for 1 mL of primary antibody solution (enough for two tubes) you would need:

800 uL 5% HIGS

200 uL 1D4 antibody

0.2 uL anti beta-gal antibody (or use 2 uL of a 1:10 dilution)

15.Incubate overnight on the nutator in the cold room.

Day 2:

  1. Wash off primary 6 x 15 minutes in PBT. Do the washes on the nutator at room temperature.

Note: SAVE THE PRIMARY ANTIBODY!!! You can RE-USE it 2-3x!!!

Note: While they’re on their last wash, make up 1 mL of 5% HIGS per eppendorf tube.

  1. Block in 5% HIGS for 1 hour on the nutator at room temperature.
  2. Add secondary antibody diluted in 5% HIGS in PBT overnight.

Note: Use HRP-conjugated goat anti-Rabbit and HRP-conjugated goat anti-Mouse 1:500

So, for 1 mL of secondary antibody solution (enough for two tubes) you would need:

996 uL 5% HIGS

2 uL HRP-conjugated goat anti-Rabbit

2 uL HRP-conjugated goat anti-Mouse

Day 3:

  1. Wash off secondary antibody 6 x 15 minutes in PBT.
  2. Wash embryos 1 x 5 minutes in PBS.
  3. To a 15 mL tube, add 5 mL of deionized water (from the Di tap), plus 2 drops of buffer, 4 drops of DAB and 2 drops of H2O2 from the Vectastain Peroxidase substrate kit in the door of the refrigerator. WEAR GLOVES! DAB is toxic. Mix well.
  4. Remove the PBS from the embryos and add 500 uL of the DAB reaction mix to each tube.
  5. Let the reaction go for about 20 minutes or until embryos LOOK GOOD. You can put a few embryos on a microscope slide to make sure they look well-stained. You want to see nice dark-staining nervous systems (not just dark beta-Gal staining!)
  6. Remove the DAB reaction mix from your embryos using a P1000 and put it back in the 15 mL tube. Also, leave the P1000 tip in the 15mL tube. Wash your embryos in PBS 1 x 5 minutes.
  7. Add 1 mL of 70% glycerol in PBS to each eppendorf tube and let your embryos sink overnight. They are now stable for LONG-TERM use. Like, years.

Important: To inactivate the DAB, add 5 mL of bleach to the 15mL tube containing the used DAB solution. Let it sit overnight, then dump it down the sink with water running.

To make 10x PBS:

Weigh out:

5.12 g NaH2PO4

23.9g Na2HPO4

204.5 g NaCl

Dissolve in 1.5L of MilliQ water

Bring pH to 7.2 and add MilliQ water to 2L

To make 10x PBT:

Do as above but add 1% Triton (1 mL / 100mL = 1% = 20 mL / 2L)