FIXING AND STAINING CELLS – A QUICK METHOD.
Require:
1) 3.7% formaldehyde in PBS or 4% parafomaldehyde or methanol:acetone
(1:1)
2) 0.1% Triton X-100 in PBS
3) 10% FCS, 0.5% BSA in PBS
Method:
1) Aspirate medium from cells and fix in 3.7% formaldehyde or 4%
paraformaldehyde or methanol:acetone IN THE FUME HOOD for 15
min.
2) Remove fixing solution and pour down fume hood sink. Wash and
permeabilise the cells 3 X 5 min. with 2ml of 0.1% Triton X-100 in PBS.
(NB I think you can miss this step if you have fixed your cells in
methanol:acetone as they will have been permeabilised at the fixing stage
however when staining centrosomes I have found that the results are
cleaner if this step is included).
3) Transfer coverslips to 24 well plate lid in a humid black box.
DO NOT ALLOW TO DRY OUT.
4) Block with 100ul of 10% FCS, 0.5% BSA in PBS for 30 min.
5) Remove blocking solution and replace with the primary
antibody/antibodies at 1/100 dilution in 10% FCS, 0.5% BSA in PBS.
Incubate for 1 hour (at least) at room temperature.
6) Wash 3 X 5 min. with 100ul 10%FCS, 0.5% BSA in PBS.
7) Incubate with 100ul of the secondary antibody/antibodies at 1/100
dilution in 10% FCS, 0.5% BSA in PBS for 40 min. At this point add
propidium iodide 1/100 dilution of 1mg/ml stock. KEEP IN THE DARK AS
MUCH AS POSSIBLE FROM THIS STAGE ONWARDS.
8) Wash 2 X 5 min. with 10% FCS, 0.5% BSA in PBS followed by 1 X 5 min.
wash in 0.1% Triton X-100.
9) Mount on glass slide with vectashield or something similar.