• Dechorionate directly on apple juice/agar plates with 50-60% chlorox; embryos will float to the surface after 2-3'.
• Collect embryos on a Millipore device with a nitex filter; rinse well with 0.1% Triton X-100 from a squirt bottle.
• Transfer embryos to 3:1 heptane:fix with a paint brush and fix for 20-40 min. Fix in a glass scintillation vial using 6.1ml HEPES Fix Mix + 0.75ml 37% formaldehyde + heptane to more-or-less fill the vial. For very small collections, stab vials may be used. Try to avoid having more than a monolayer of embryos (tilt vial to increase surface area).
• Devitillenize with a methanol shock. Remove the aqueous (bottom = fix) phase completely (use a drawn out pipet) as well as one pipetful of heptane. Fill the vial with MeOH and vortex vigorously for 20-30". If much fix remains, fewer embryos are released from their vitelline membranes. Very young embryos are hard to get out of the membrane in any case. If you need these, try the additional steps in the Mithchinson and Sedat paper.
• Allow the phases to separate. Aspirate off the upper phase, embryos at the interface and, finally, most of the lower phase.
• Transfer embryos to a microfuge tube using a blue tip with the end cut off. Rinse several times with MeOH. The embryos may be stored at this stage @ 4 degrees C for several months. Note that some antigens are methanol-sensitive and for these this type of storage is obviously not possible.
• Rinse embryos several times in a 30 min to 2 hr period with TBS.
• Add primary antibody in TBS and rotate @ 4 degrees C ON.
• Wash with TBS+N 2h @ RT. Do two quick washes and at least two more 30' washes.
• Add secondary antibody in TBS and rotate 2-4h @ RT.
• Wash 1X in TBS and 3X in PBT.
• For HRP-conjugated 2o: mix 2ml PBT + 0.5mg DAB + 0.5ml H2O2, use immediately (staining takes 1-10'). Notes on the use of other conjugated secondaries below.
• For mounting, dehydrate the embryos in EtOH, pipet onto a slide and cover with a coverslip smeared with GMM. The slides are immediately placed in a 55-60 degrees C oven or on a slide warmer overnight.
  

• Biotinylated secondary antibodies can be preadsorbed against embryos, although my current view is that this is not necessary. I use Tago G anti- Ms, Rb and Rt.
• I use HRP-avidin and AP-avidin from BRL @ 1:2000 and 1:1000, respectively. Add avidin-enzyme conjugate in PBT, rotate 30'.
• Wash 3X in PBT with a final wash in AP buffer if using AP-conjugate.. The AP staining conditions are as described in the BRL handout that comes with the conjugate. Although the AP stain is more sensitive than HRP, the background is also higher and the embryos will eventually turn dark purple throughout. There is also a segmental background pattern on older (germ band retracted) embryos.
• For fluorescence, tubes are wrapped in foil after addition of the conjugated 2o. Subsequent washes are in PBT, and the embryos mounted in 1:1 0.2M sodium bicarbonate containing 5 mg/ml p-phenylenediamine (antibleaching agent- Sigma P-6001):glycerol. Make the sodium bicarb (pH 8.5) fresh each time, and wrap in foil during the time when the phenylenediamine is dissolving.

HEPES Fix Mix
0.1M Hepes, pH 6.9
2mM MgSO4
1mM EGTA

TBS
150mM NaCl = 30ml 5M NaCl
50mM Tris-HCl, pH 7.5 = 50ml 1M Tris-HCl
0.1% BSA = 10ml 10% BSA H2O to 1L

0.1% Triton X-100
1ml 100% Triton X-100
0.05% Thimerosal = 0.5g Thimerosal (Hg-containing poison!)

TBS + N
TBS + 2% appropriate serum (e.g., goat for goat secondary)

PBT
PBS + 0.1% Tween-20

10X PBS
1.3M NaCl = 76.0 g NaCl
0.07M Na2HPO4 = 9.94g Na2HPO4 (MW=142)
0.03M NaH2PO4 = 3.6 g NaH2PO4 (MW=120)
H2O to 1L

GMM is Canadian balsam:methyl salicylate (‰20:1)