Non-fluorescent protein/RNA double-labeling

1. Collect embryos, dechorionate and fix in a (1:1) mixture of heptane:fix. Fix is 5% formaldehyde, 0.05M EGTA in 1X PBS. pH 7.0. Devitellinize embryos by shaking in an equal mixture of methanol:heptane (embryos will sink to the bottom) and wash the embryos well with methanol. Transfer the embryos to a 0.5 ml eppendorf tube.

2. Wash the embryos with 3% hydrogen peroxide in methanol for 15 min. Rinse well in methanol.

3. Wash 3 X 30 minutes with PBTH (filtered 1 X PBS, 0.1% Tween-20, 50 ug/ml heparin, and 250 ug/ml tRNA). You can also include 1% BSA providing you know it is ultra pure/RNase-free. With our reagents, we find we get similar results without it.

4. Incubate with the primary antibody (diluted appropriately in PBTH) overnight at 4C. For extra RNase protection, include 2- 5 Units of RNasin/250 ul of antibody mixture.

5. Wash at room temperature with several changes of PBTH for at least 1-2 hr.

6. Incubate with biotinylated secondary antibody (diluted in PBTH) for 1.5 hours at room temperature (again add RNase inhibitor).

7. Wash with several changes of PBTH for 60 minutes. For detection of the biotinylated secondary antibody, we generally use the Vector Lab ABC amplification system. Mix the reagents per manufacturer's instructions 30 min. prior to use.

8. Incubate embryos with the pre-incubated ABC mixture for 30 minutes at room temperature.

9. Wash for 15-30 min. with 3-4 changes of PBTH. Develop the signal with 20 ug/ml Diaminobenzidine, 0.03% hydrogen peroxide in PBTH. This will produce a brown signal. Stop the reaction with several rinses of PBTH.

In situ HYBRIDIZATION :

1. Fix the embryos for 20 minutes with 5% formaldehyde in PBT (PBT= filtered 1 X PBS with 0.1% Tween-20) at room temperature.

2. Wash 3 X 5 minutes with PBT.

3. Incubate the embryos with 50 ug/ml of Proteinase K in PBT for 2-3 minutes (time varies with different Prot. K preps and needs to be optimized).

4. Stop the Proteinase K reaction with two washes of 2 mg/ml glycine in PBT. Each wash should be 2 -3 minutes. Then rinse the embryos twice with PBT.

5. Fix the embryos again for 20 minutes with 5% formaldehyde in PBT at room temperature.

6. Wash the embryos 5 times with PBT.

7. Rinse the embryos with PBT:Hybridization buffer (1:1), and then with hybridization buffer alone. Hybridization buffer for DNA probes is: 5 X SSC, 100 ug/ml sonicated, boiled salmon sperm DNA, 100 ug/ml tRNA, 50 ug/ml heparin, and 0.1% Tween-20 (for RNA probes and hybridization conditions, see for ex. our protocol on FISH whole-mount staining).

8. Pre-hybridize the embryos in hybridization buffer for at least one hour at the appropriate temperature (48 C for DNA probes). Longer pre-hybridizations of up to 3 hours seem to improve the signal to noise ratio.

9. Add your Digoxygenin-labeled probe to the hybridization buffer and incubate overnight (12 to 16 hours) at the appropriate temperature. Use about 100 ul hybridization buffer per 50 ul settled embryos.

10. Carry out the following post-hybridization washes at the hybridization temperature for at least 20 minutes each:

(i) hybridization buffer alone

(ii) PBT:hybridization buffer (1:1)

(iii) PBT alone for at least 5 washes

11. Incubate with anti-Digoxygenin antibody (Boerhinger Mannheim) conjugated to alkaline phosphatase (diluted 1:2000 in PBT) for 2 hours at room temperature.

12. Wash with PBT 4 times for 20 minutes each wash. Rinse with alkaline phosphatase staining buffer (100mM NaCl, 50 mM MgCl2, 100 mM Tris pH 9.5, 1 mM Levamisol, and 0.1% Tween 20).

13. Develop the colour reaction using NBT and BCIP in the alkaline phosphatase staining buffer as described by the manufacturer (Boehringer Mannheim).

14. Stop the reaction with several washes of PBT and dehydrate with an ethanol series (in PBT).

15. Rehydrate the embryos in a reverse ethanol series (in PBT), and then rinse several times with PBT.

16. Mount in 70% glycerol and 30% 0.1 M Tris pH 8.0.