Hi everyone,

Great suggestions, thanks for sharing your thoughts. I have tried using fresh dehydrating alcohols, but still got the same results. I may try some of the different mounting media you suggested and a different source of DAB. I have not tried leaving out the differentiation step with acid alcohol. Is there a particular bluing reagent that works best?

The details of my protocol are as follows:

The sections were fixed in Modified Davidsons solution, paraffin embedded, re-hydrated, underwent sodium citrate heat-mediated antigen retrieval for 20 minutes, rinsed in PBS, incubated with 10% normal goat serum in PBS for 1 hour at room temperature, incubated with primary antibody in PBS overnight at 4 degrees C, rinsed in PBS, quenched for endogenous peroxidase in 3% hydrogen peroxidase for 15 minutes, rinsed in PBS, incubated with biotinylated secondary antibody in PBS for 1 hour at room temperature, rinsed in PBS, incubated with avidin and biotinylated horseradish peroxidase complex for 30 minutes, rinsed in PBS, developed with DAB (Invitrogen) for 1-2 minutes, counterstained with hematoxylin, rinsed in water, differentiated in acid alcohol, rinsed in water, blued in 0.2% ammonia water, rinsed in water, dehydrated in three changes of isopropanol, cleared through three changes of xylene, and mounted using Cytoseal XYL.

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Why does my DAB signal disappear so quickly?