Protocol for methanol fixation for immunofluorescence

This protocol is recommended for proteins of the post-synaptic density (PSD)

Methanol fixation works by denaturing and precipitating proteins, and as such it is a quick method. For most antibodies/proteins it takes only 2 - 5 minutes. This procedure leads to an unmasking of the proteins in the PSD.

Solutions needed:

MES: 100 mM MES, pH 6.9; 1 mM EGTA; 1 mM MgCl₂. Store at 4°C
Methanol fix (100 ml): 10 ml MES; 90 ml methanol. Store at -20°C
PBS, pH 7.4 (0.1 % Tween-20 can be added for the washing steps)
3 % BSA in PBS, pH 7.4 (0.1 % Tween-20 can be added for the washing steps)

Procedure:

Rinse coverslips in 70 % ethanol and flame.
Let cells grow on coverslips to desired density. Note that some cell lines require poly-lysine coated cover slips for proper adhesion.
Transfer coverslips to 6 well plate(s) and wash twice with PBS.
Pour some -20°C Methanol fix carefully onto the plated cells and leave the dish on the bench
for 5 min.
Wash the cells repeatedly with PBS. There is no need for a permeabilization step following this fixation.
Block the cells with 3 % BSA in PBS.
Put a piece of parafilm on wet Whatman paper and apply 200 µl of primary antibody solution in BSA/PBS. Appropriate dilution must be determined experimentally.
Put coverslips upside down on antibody-solution and incubate for 1-2 h at RT.
Transfer slips to 6 well plate and wash twice with PBS.
Incubate for 10 min with 3 % BSA in PBS.
Repeat steps 10-13 with secondary antibody (1 : 1000 dilution). Avoid bright light when working with the secondary antibody to minimize photo bleeching of the fluorescent dye.
Wash twice with PBS.
Add a drop of Mowiol or DAKO fluorescent mounting medium on slide and mount dry coverslip.
Incubate for 30 min at RT.
Incubate over night at 4°C.
Incubate for 30 min at RT.
Seal with nail polish and microscope.