I'm not sure!

This is my first time working with IHC.

And here is my protocol:

IHC-Protocol:

All the tissue will be stained on net-wells.

1- Rinse tissue 3x10 min in 0.1M TBS (5x if in antifreeze previously).

2- Incubate in 0.3% H2O2 in PBS (peroxidase block) at room temperature for 20-30 minutes to block endogenous peroxidase activity.

3- Rinse 3x10 minutes in PBS and 0.3% Triton X-100 (PBST).

4- Incubate with 1.5% normal blocking serum (NBS) in PBS for 1h.

5- Rinse 3x10min in PBST.

6- Incubate in primary antibody (goat anti-CB1R), diluted 1:500 in 1.5% NBS/PBS overnight at 4°C.

 7- Rinse 3x10 minutes in PBST.

8- Sections will be incubated in biotinylated secondary antibody diluted in 1:200 for 1 hour.

9- Sections will be rinsed again 3x10 minutes in PBST.

10- Incubate in Avidin D-HRP complex reagent for 2 hours at RT on shaker (prepare 30 min before use), diluted in 1:300.

11- Rinse 3x10 minutes in PBS and 0.3% Triton X-100.12- Diaminobenzidine (DAB) solution will be made freshly before use to detect the color of antibody staining (3 minutes).

12- Diaminobenzidine (DAB) solution will be made freshly before use to detect the color of antibody staining (3 minutes).

13- After DAB, rinse for 2 minutes each for the first 2 rinses in PBS.

14- Rinse 3x10 minutes in PBS.

15- Mount tissue on the microscopic slide.

16- Allow mounted sections to dry overnight (up to 2 days) at RT (cover to protect from dust). 

17- Counterstain slides by immersing in Methyl Green for 1-2 min.

18- Wash with distilled water several times.

19- Dehydration: 4 changes of alcohol (95%, 95%, 100%, and 100%) for 2 minutes each.

20- Clear with xylene 2x2 minutes.

21- Mount and cover slip.

I think your primary antibody is not reacting with the antigen at the antigenic site. Ensure the right primary antibody is used. i will also suggest u try avidin-biotin complex(ABC).

You can modify the antigen retrieval step for proper exposure of the epitope.

Thank you both for your answers.

I have tried a new antibody and change the dilution as well. 

And as I mentioned before that I'm not staining the sections on the slides, instead I stained them first in net-wells (like a 12 wells plate), then I mount the tissues on the slide. The reasons for that is my sections thickness is 40uM.

I never worked with DAB and mounting of sections after staining, so it's just a guess:

For my experience it "kills" frozen sections/immuno-stains if you let them dry out once they've been wet and many antigens do not like organic solvents like alcohols. If I were you I would try to mount them on slides before staining no matter how thick they are. After IHC staining use a water-based, non-hardening mounting medium for microscopy, take your pictures and whatever you need. Now remove the cover slip again (possible with a buffer wash) and go on with the counter staining procedure.

Good luck!

Hi Ghadar Sindi,  your protocol sound good. I recommand the free floating method if you  like to stain 40 µm thick brain sections. Use 24well plates, take one well/section. 500 µl ab solution per well is suficient. What about your secondary ab, ddt you us a biotinylated anti-goat-ab?  As Leyira told you use a ABC- Kit (Vector Labs  is highly recommanded). If you prepare your DAB solution freshly don't forget to add H2O2 otherwise the DAB reaction does not work.  Make a dilution row to optimise the concentration of your primary ab. After you have mounted your sections leave them air dry and countrtstain them with Methylene Green or with Meyer's Hematoxyline as you have discribed  it. Hope it helps you. Best regards Ute

Thank you all, I will try all of your suggestions.

 Ute Neubacher: How long should it take to dry the tissues? I usually let them dry for 2 days! is that Okay!