Cover cells with or submerse constructs in 10% formalin (4% formaldehyde) for not longer than 24 hours.
Remove formalin or remove from formalin and store in dH2O until staining or dehydrating
10% formalin can be found in the histology lab
1. Briefly rinse cells with PBS at 37°C.
2. Fix samples with 4 % paraformaldehyde in PBS for 10 minutes at RT.
3. Wash samples 3x with PBS, for two minutes each time.
4. Permeabilize cells with 0.2% Triton X-100 diluted with PBS, for 5 minutes.
5. Rinse samples 4x with PBS, for two minutes each.
Cell monolayers
Paraformaldehyde Fixation
1. Prepare a fresh 4% solution of paraformaldehyde by heating in buffer (PBS, TRIS, HEPES, Veronal, etc.) to 55- 60 ° C. Add several drops of 1N NaOH and stir until dissolved. Allow cooling to room temperature before use.
2. Wash the coverslips once with PBS and then fix with PFA for 15min.
3. Wash the coverslips once with PBS and then permeabilise the cells with 0.2% Triton-X100 in PBS for 5min.
4. Wash the cells once with PBS and then quench in fresh 0.1% sodium borohydride in PBS for 5min.
5. Wash three times in PBS and proceed to staining protocol.
Staining
1. Dilute primary antibody as appropriate in 1% BSA in PBS. Centrifuge the diluted antibody for 5 min at 12,000 x g in a refrigerated microcentrifuge prior to use will remove aggregated material and reduce background.
2. Incubate the coverslip with a small volume of diluted primary antibody for 1h.
3. Dilute the secondary fluorescent antibody as appropriate in either PBS or 1% BSA in PBS, depending on the background staining. Again, centrifuging the diluted antibody for 20 min at 12,000 x g in a refrigerated microfuge will reduce background.
4. Incubate the coverslip with a small volume of diluted secondary antibody for 45min.
5. Wash coverslips three times with PBS.
Mounting
Mount in appropriate mounting media containing an anti-photobleaching reagent (glycerol, Mowiol, BABB etc. with 0.6% DABCO, p-phenylenediamine, etc). Coverslip, seal with nail polish and store in the dark at 4 degrees C.
DAPI
1. Equilibrate cells in PBS.
2. Dilute DAPI 1:300 to 1:1,000 with PBS.
3. Incubate cells for up to 5 minutes.
4. Wash 3x with PBS.