1. Fix the embryos in 4% paraformaldehyde in PBS either for 2 h at room temperature or overnight at 4°C.
2. Wash the embryos in PBS containing 1% Triton X-100 three times for 30 min each.
3. Wash the embryos once in 1X terminal transferase buffer, 2.5 mM CoCl2, 1% (v/ v) Triton X-100.
4. Incubate the embryo for 3 h at 37°C in 1X terminal transferase buffer, 2.5 mM CoCl2, 0.5 |/mL terminal transferase, 10 mM dUTP (2:1 dUTP:dUTP-biotin), 1% (v/v) Triton X-100.
5. Wash the embryo three times in PBS, 1% (v/v) Triton X-100 for 30 min.
6. Incubate overnight at 4°C with rocking in an appropriate streptavidin conjugate, such as strepevidin-fluorescein or streptevidin-horseradish peroxidase.
7. Wash the embryos three times in PBS, 1% (v/v) Triton X-100 for 1 h each. Then proceed either according to step 8 or steps 9-12 as appropriate.
8. In the case of streptavidin-fluorescein detection, mount the specimens under 90% (v/v) glycerol, 1X PBS containing the antiquenching agent DABCO at 2.5% (v/v), and view under fluorescent optics.
9. Wash the embryos twice in PBS for 30 min each.
10. Incubate the embryos in DAB in PBS (5 mg/10 mL PBS) for between 1 and 3 h, depending on the stage of the embryo at 4°C with rocking in the dark.
11. Exchange the above solution for the same, but this time add hydrogen peroxide (60% or 200 vol), 10 |L/10 mL DAB in PBS, and develop in the dark for 5-15 min.
12. When the background staining starts to appear, stop the reaction by washing several times with PBS.
13. Wash in PBS twice each for 30 min, and then mount the specimens under a glass coverslip in 90% glycerol/10% PBS and view under bright-field optics.
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