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Antibody Staining of Whole Mount Drosophila Embryos

Whole mount staining means the staining of small pieces of tissue (normally embryos) without sectioning. It is very similar to immunocytochemistry (ICC) or staining of cryo-sections. The difference is that the sample being stained is much larger and thicker than a normal section on a slide. Thus, incubations for fixative, blocking buffer, antibody, wash buffer, permeabilization and substrate color development will need to be much longer to allow for permeabilization right into the center of the sample. Creative Biolabs here provides the protocol of antibody staining of whole mount Drosophila embryos. The protocol need to be optimized for your experiments.

Reagents

Methods

  1. Prepare fixation mix in a 1.5ml tube: 400µl PBS, 100µl 4% formaldehyde, 500µl n-heptane.
  2. Note: For the detection of some extracellular antigens, add 0.02% SDS or to fix the embryos in 500µl picric acid / 500µl n-heptane. Vortex the fixation mix at highest speed for 1min.

  3. Remove yeast. Dechorionate embryos by covering the apple-juice plate with 100% bleach for 2min. Slight rotations of the apple-juice plate will bring the dechorionated embryos to the surface. Close the narrow opening of a funnel with a nylon-mesh. Attach mesh by wrapping a rubber band around the funnel.
  4. Wash embryos into the funnel by squirting deionized water over the plate. Wash embryos in funnel by squirting water along the funnel three times.
  5. Remove rubber band and mesh. Pick up the embryos with a very fine paint brush. Dechorionated embryos will adhere to the brush. Dip brush with embryos into the fixation mix and whisk the brush.
  6. Fix embryos for 15min on a shaker with gentle rotation.
  7. Remove about 80% of the lower phase of the fixation mix with the pipette to stop fixtion.
  8. Add 1ml of methanol and vortex on highest speed for about 1min. This step will remove the extraembryonic membrane.
  9. After vortexing for 2 mins, the majority of the embryos will be at the bottom of the cap. If not, all embryos sink to the bottom of the cap, remove as much of the liquid as possible and refill the cap with methanol. Do not vortex again. After most of the embryos are at the bottom of the cap, remove all liquid and wash embryos three times with methanol.

  10. Rehydrate the embryos with PBT for three times. Incubate in PBT containing 20% NCS on shaker for at least 10min.
  11. Add primary antibody diluted in PBT containing 10% NCS and 0.02% sodium azide. Incubations can last from 2h at RT (high affinity antibodies) to overnight at 4°C (low affinity antibodies).
  12. Note: Overnight incubation at 4°C aides the perfusion of the antibody.

  13. Wash procedure:
    • Three rinses with PBT, 10min incubation with PBT containing 30% NCS.
    • Three rinses with PBT, 10min incubation with PBT containing 30% NCS.
    • Three rinses with PBT.
  14. Add secondary antibody diluted in PBT containing 10% NCS for 2h at RT.
  15. Repeat step 10.
  16. Non-fluorescent detection: use Diaminobenzidine assay (normal signal) or Alkaline Phosphatase assay (weak signals). Develop Alkaline phosphatase signal in the dark and take an aliquot of embryos out of the cap with a 1ml pipette to follow the staining. Postfix embryos with 4% formaldehyde / PBT for 10min to stabilize Alkaline Phosphatase signal.
  17. Incubate embryos in 50% glycerol / PBS until they sink to the bottom of the cap (about 10min). Replace the 50% glycerol/PBS with 70% glycerol/ PBS wait again until embryos are at the bottom of the cap (about 1h). Replace 70% glycerol/ PBS with 90% glycerol/PBS and allow embryos to sink to the bottom of the cap. Embryos can be stored in 90% glycerol / PBS at 4°C for at least three years.

Tips

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