This is the immunofluorescence protocol I use:

• Obtain your antibodies
• Remove paraffin using the following washes:

·      Xylene-3min

·      Xylene-3min

·      100%Alcohol-3min

·      100%Alcohol-3min

·      95% Alcohol-3min

·      70%Alchocol-3min

·      water-3min

·      PBST (Phosphate Buffer Saline+Triton)-10 min

• Incubate for 30 minutes in 5% normal goat serum in PBST (Phosphate Buffer Saline Triton-X)
• Apply primary antibody at desired dilution in PGB (PB pH7.4, normal serum, Triton-X,BSA) overnight at 4C
• Negative control in place without primary antibody (only PGB)

• Continue protocol the following day
• Remove samples from 4C and wash three times in PBST for 10 min each
• Prepare secondary antibody AF-488 Goat anti Mouse or Rabbit (Mol Probes) 1/200 in PBG and incubate for 2 hrs at RT
• Wash twice with PBST for 10 min each
• Wash in Hoersct solution for 10 min and then a final PBST wash for 10 min
• Coverslip in Aqueous based mounting medium (Dako Flouromount) and store at 4C in aluminum foil
• Image as desired

Best of Luck!

Thank you very much!!! Really Appreciated!!!

For bright field : instead of fluophore labeled secondary abs you can use biotinylated secondaris 1:500 for 90 min and the detect them with ABC-Peroxidase 1:1000 for 90 min (both from Vector Labs). Visualisation of the reaction by DAB-staining for 10 min, dipends of the dilution of the primary abs. After dehydration in ascending alcohols, clearing in Xylene you can mount the sections in Depex or a similar xylene soluble mounting medium.

Thanks!! I will consider it...