TITLE: Free-floating Mouse Brain Immunohistochemistry
AUTHORS: Jonathan Breiter

[DESCRIPTION]
This protocol enables immunohistochemical staining of murine tissue with superior penetration of the tissue by the reagents due to the free-floating approach.

In this new version, the last step contains a supplemental video with extra context and tips, as part of the ASAP Protocol Particulars, featuring conversations with protocol authors

[STEPS]
SECTION: Tissue Preparation
1. Remove PFA-fixed tissue from storage solution and add to mesh bottom netwell insert inside of 12 well plate that is filled with 0.22 μm filtered 1x PBS. 

SECTION: Buffer Preparation
3. Per well of tissue make a minimum of 3.5 mL of blocking solution, consider this is needed for blocking, primary and secondary antibody incubation. All reagents should be 0.22 μm filtered:

1x PBS
5% Normal Goat Serum
2.5% Bovine Serum Albumin
0.2% Triton-X

Make in excess and keep on ice. Keep at 4 degrees celcius overnight. 4 °C 

SECTION: Tissue Preparation
2. Place the 12 well plate with the tissue on a horizontal shaker and wash tissue 3x 5 min at approx. 150 rpm, moving the netwell insert to the next well down after each 5 min period to immerse the tissue in new 1x PBS. Room temperature 

SECTION: Primary Antibody Incubation
4. In a new 12 well plate, add 2-3 mL per well of blocking solution and transfer the washed tissue sections inside their netwell inserts into these wells. Incubate for 1h - 2.5 h  on horizontal shaker at approx 150 rpm. Room temperature 

SECTION: Primary Antibody Incubation
5. Meanwhile, dilute primary antibodies in blocking solution to appropriate concentrations and keep on ice .

E.g.
- Mouse monoclonal anti pS129 (81A) (ab184674) @ 1:750 
SECTION: Primary Antibody Incubation
6. Add approx 250 ul of primary antibody solution to the approriate number of wells of a 24 well plate for the number of brain sections. 
SECTION: Primary Antibody Incubation
7. When blocking is finished, move brain tissue sections from the netwell inserts to their appropriate primary antibody well using a fine paintbrush, being careful not to destroy the tisssue. 
SECTION: Primary Antibody Incubation
8. Incubate on horizontal shaker at approx. 150 rpm overnight. 4 °C 

SECTION: Secondary Antibody Incubation
9. Prepare a new 12 well plate with clean netwell inserts and fill all wells with 0.22 μm filtered 1x PBS.
SECTION: Secondary Antibody Incubation
10. Transfer brain tissue sections from 24 well plate into netwell inserts inside 12 well plate and wash 4x 10 minutes on a horizontal shaker at approx. 150 rpm. Room temperature 

SECTION: Secondary Antibody Incubation
11. Meanwhile, make appropriate dilutions of secondary antibody in blocking solution. Keep away from light and keep on ice .

E.g.
- Donkey anti-Mouse Alexa Fluor 568 (A10037) @ 1:500
SECTION: Secondary Antibody Incubation
12. Add approx 250 ul of secondary antibody solution to the approriate number of wells of a 24 well plate for the number of brain sections. 
SECTION: Secondary Antibody Incubation
13. When washing step is finished, move brain tissue sections from the netwell inserts to their appropriate secondary antibody well using a fine paintbrush, being careful not to destroy the tisssue. 

Keep the well plates covered from now to avoid bleaching of fluorophores. 
SECTION: Secondary Antibody Incubation
14. Incubate 24 well plate on a horizontal shaker at room temperature at approx. 150 rpm for 1 h - 2.5 h. Room temperature 

SECTION: Secondary Antibody Incubation
15. Transfer brain tissue sections from 24 well plate into netwell inserts inside 12 well plate and wash 4x 10 minutes on a horizontal shaker at room temperature at approx. 150 rpm. Room temperature 

SECTION: Microscope slide preparation &amp; Imaging
16. Plasma-clean microscope cover slips in Argon plasma for 15 minutes. 
SECTION: Microscope slide preparation &amp; Imaging
17. Using a fine paintbrush, transfer brain tissue sections from 1x PBS onto a microscope slide, using excess 1x PBS to mount multiple sections next to each other and making sure they are not folded over. 
SECTION: Microscope slide preparation &amp; Imaging
18. Leave sections on microscope slides to dry out in the dark (they will turn white). 
SECTION: Microscope slide preparation &amp; Imaging
19. Add approx. 150 uL of mounting media on top of each tissue section and apply the plasma-cleaned cover slip on top of the tissue, closing the slide. 
SECTION: Microscope slide preparation &amp; Imaging
20. Leave the mounting media to dry in the dark.
SECTION: Microscope slide preparation &amp; Imaging
21. Seal the edges of the coverslip with nail varnish and let dry for approx. 30 minutes in the dark.
SECTION: Microscope slide preparation &amp; Imaging
22. Immediately take finished microscope slides to imaging, avoiding unecessary light exposure. 
SECTION: ASAP Protocol Particulars: context and tips
23.