Protocol for Immunoperoxidase staining of paraffin tissue sections

1. Clean slides with 95% ethanol
2. Treat with a solution containing 0.3% gelatin and 0.05% chromium poatassium sulfate
3. Cut tissue sections using microtome and apply to slides
4. Deparafinize in xylene for 10 minutes. Repeat twice for a total of three treatments
5. Hydrate sections through graded alcohols
6. Rinse three times with distilled water
7. Incubate for 10 minutes with 0.1% H₂O₂ in PBS to quench endogenous peroxidase activity
8. Rinse four times with PBS
9. Incubate slides for 20 minutes in 10% goat serum in PBS (suppresses non-specific binding of IgG)
10. Wash with PBS
11. Incubate with primary antibody for 1 hour at room temperature or overnight at 4^oC. Optimal antibody concentration is usually from 1-10ug/ml in PBS-BSA
12. Wash three times with PBS
13. Incubate with biotin-conjugated secondary antibody for 45 minutes. Again, optimal antibody concentration is usually from 1-10ug/ml in PBS-BSA
14. Wash three times with PBS
15. Incubate with streptavadin-HRP for 15 minutes. This step should be titrated as excess streptavadin-HRP can lead to high background
16. Wash with 0.5% triton x-100 in PBS for 30 seconds
17. Incubate with fresh DAB solution for 5 minutes. This step can be lengthened if additional sensitivity is necessary. It is a good idea to do a quick spot check here
18. Rinse with distilled water
19. Counterstain with hematoxylin
20. Dehydrate with alcholos and xylene
21. Mount permanent coverslip

Note: Incubations are to be at room temperature or a 4^oC humidified chamber. Antibody concentrations here may not hold for all sources. These concentrations should be titrated for each source.