1. Early in the day, make up 400 mL fresh 4% paraformaldehyde in PBS, and store on ice until required. Also ensure that you have 3.5 L of DEPC-treated water for dilution of salines, etc. Use designated RNase-free glassware and racks for all of the following procedures, wear gloves throughout, and use DEPC-treated water and solutions.
2. Dewax slides in xylene twice each for 10 min in a fume cupboard. For this and all subsequent steps, use 350 mL of each solution for each incubation; sufficient to cover the slides completely. Slides are processed in metal racks holding up to 25 individual slides in appropriately sized staining troughs. Do not agitate the slides during incubations, since it may cause sections to detach.
3. Place in 100% ethanol for 2 min to remove most of the xylene. Meanwhile make up 1.6 L 1X PBS and 800 mL 1X saline.
4. Rehydrate sections by "dipping" through the following graded alcohol series 100% ethanol twice, 95% ethanol, 80% ethanol, 70% ethanol, 50% ethanol, and 30% ethanol. (Keep and reuse ethanols up to 10 times).
5. Place in 1X saline for 5 min and then in 1X PBS for 5 min. Discard used saline and PBS.
6. Place in the freshly made, cold 4% paraformaldehyde in PBS for 20 min in the fume hood. At the end of this, do not discard the fix, but keep for step 11 of Subheading 3.6.
7. Meanwhile for 25 slides, prepare 15 mL of DEPC-treated TE, pH 8.0, mixed with 20 ||L of proteinase K (20 mg/mL). Also wash sufficient cover slips of appropriate size in ethanol and leave to air-dry in a rack covered in foil.
8. Place slides in 1X PBS twice each for 5 min, and then lay flat on a sheet of foil on the bench with the sections uppermost.
9. Cover sections with 300 ||L or more of proteinase K (sufficient to cover the sections), and incubate for exactly 7 min at room temperature.
10. Tip off the proteinase K, and place slides in a rack in fresh 1X PBS for 5 min.
11. Return the slides to the cold paraformaldehyde (Subheading 3.6., step 6) for 5 min in the fume hood.
12. Meanwhile, fill two troughs with DEPC water, and place in the fume hood. To the second (acetylation bath), add 6 mL triethanolamine and add 1 mL acetic anhydride, and mix with a small stir bar on a stirrer.
13. Dip slides briefly into the DEPC water, and then place in the acetylation bath for 10 min with stirring. (A small stir bar or "flea" will fit beneath standard slide racks and allow stirring in the presence of the latter. This step acetylates slides and sections, and helps prevent nonspecific binding of probe).
14. Place slides in fresh 1X PBS for 5 min and then fresh 1X saline for 5 min.
15. Dehydrate by "dipping" through the same ethanol series used in Subheading 3.6., step 4 starting with 30% ethanol and increasing to the two 100% ethanols.
16. Place slides section side up to air-dry on foil. Add probe to them the same day.
17. Dilute probe in hybridization buffer to give 105 dpm/|L/Kb.
18. Mix probe by vortexing and briefly spin in a centrifuge. Heat to 80°C for 3 min to remove secondary structures owing to internal hybridization of the probe. Allow to cool briefly on ice and then mix again.
19. Place 15-30 pL of probe solution on slide adjacent to sections, spread over sections with a piece of parafilm, and cover with a clean cover slip (Subheading 3.6., step 7) taking care not to introduce any bubbles.
20. Slides are collected on their sides in a rack that is then place inside a suitable box with a tightly sealed lid. Include a tissue soaked in 5X SSC, 50% formamide to prevent dehydration. Seal the box with suitable tape, and hybridize at 55°C overnight in an oven.
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