Last updated 8/97 by DAS.
Required solutions are given below.
1. Cells should be plated in 35 mm dishes containing coverslips. Fix cells when the cell density is not too high so the cells will not be crammed together. Usually plating at 1-2x105 cells per 35mm dish will be OK for next day use.
2. Prepare fix just before use: I usually use 2% paraformaldehyde in PBS but for some cells I use a different buffer. Use PBS unless indicated.
For 10 ml of fix (adjust volumes as needed):
3. Remove media from dishes and rinse cells once in ~1ml of HBSS or PBS (room temperature).
4. Remove HBSS and add 1ml of 2% paraformaldehyde in PBS. Incubate at room temperature for 15 min.
5. Remove fix and quench excess aldehyde by incubation in 1mg/ml sodium borohydride prepared just before use in PBS, pH 8. Incubate in this solution for 15 min at RT.
6. Permeabilize cells by incubation in 0.1-0.5% Triton X-100 in PBS for 10 min at RT.
7. Block: Incubate fixed and permeabilized cells in blocking solution for 30 min.
8. Prepare primary antibody solutions diluted in blocking buffer. Need about 150ml/coverslip. Microfuge the diluted antibody solutions for 15 min in the cold room.
9. Transfer coverslips to a box containing moist filter paper covered with parafilm. Mark a grid on the parafilm to help keep track of the coverslips. Lay the coverslips cell-side-up on the parafilm. Only transfer a few at at time so they don't dry out.
10. Apply antibody solution to each coverslip. Incubate in primary antibody for 2 hours at RT or overnight at 4°C.
11. Transfer coverslips back into their dishes and wash them 3 times with TTBS, 15 min/wash.
12. Prepare appropriate secondary fluorescent-labeled antibody solutions and apply to coverslips as above. Dilute antibodies in the blocking solution. Incubate coverslips for 2 hours at RT in a dark cabinet.
13. Wash coverslips as above in TTBS.
14. Mount the coverslips onto glass slides (can put two on each slide) using mounting media that contains an anti-bleaching agent (sometime I buy this from Molecular Probes but if there is none, prepare n-propylgallate solution as described). Aspirate excess mounting liquid from the slide and seal coverslip in place with clear nail polish (Revlon is best--$$ but worth it).
15. Look at on scope.
10 % paraformaldehyde stock (for preparation of 10 ml)
Weigh out 1g of paraformaldehyde (special EM grade). Add to 10 ml H2O in a 15ml capped tube and shake vigorously. The PFA will not go in to solution yet. Heat in 65°C water bath. Add 1-2 drops from a yellow Pipetman tip of 1M NaOH to the suspension and shake vigorously. The PFA will begin to dissolve. Continue heating until most of it is dissolved. Any remaining undissolved stuff can be filtered out using a 0.22 m disposable filter.
Blocking Solution
TTBS (Tris-buffered salt solution with Tween) is 0.3 M NaCl, 20 mM TrisCl, pH 8.0, 0.1% (v/v) Tween-20 and 0.01% NaN3
n-propylgallate/glycerol mounting medium
Best if made fresh each time, but you can store it in dark at 4°C for about 1 week. It gets yellow colored as it ages, so don't use it if it is a weird color.