1. Use 15 ml conical Corning tubes. Harvest cells (10 ml at 2 x 105 cells/ml).
2. Wash in 10 mM Tris pH 7.4, 10 mls, 3000 rpm, 5 min
3. Concentrate cell to <0.5 ml.
4. Extract 3 minutes in 0.5% Triton X-100, 1 mM Taxol in PHEM (use 3 ml).
5. Spin 2 minutes at 455-500 x g (~2000 rpm).
Cells should be in a loose suspension—not a true pellet. Remove
supernatant with a Pasteur pipette.
6. Wash in 3 ml of PHEM, 1 mM taxol by centrifugation as above, concentrate to 0.5 ml. Shake gently to resuspend.
7. Add 3 ml of 2% paraformaldehyde in PHEM, swirl gently, fix 30 min. to 1 hr.
8. Wash in PBS (3 ml) 1x.
9. Resuspend in 0.5 ml PBS. Fixed cells can be stored at 4°C for
up to a month (even longer) without decrease in quality.
II. Antibody Incubation and Washes
Method A with drying
1.Take 50 ml of fixed cells and wash with 0.5 ml of PBS.
Mark the non-cell side of poly-L-lysine coated coverslips with nail
polish or model paint so you can identify each coverslip.
2. Drop ~50 µl cells in PBS onto the coverslip and let
air dry (takes some time can be done at 30-37oC). Prepare two cover slips
for each sample.
3. Once dry, align each set of two coverslips back to back (cells
side outside), insert into small (Coplin) staining jars with PBS, incubate
5 min. at room temp.
4. Blot excess liquid by touching edge of coverslip to a kimwipe.
5. Incubate coverslips in 3% BSA/PBS/0.1% Tween, 3 x 5
min. in a jar.
6. Place a piece of parafilm in a plastic box with a layer of
wet paper towels (moist chamber). Drop 50 µl of antibody solution
(For the 6-11 B1 antibody use 10 ml of monoclonal antibody + 40 ml of PBS+BSA+Tween)
onto parafilm, drain PBS off coverslips, lay coverslip with cells
face down on top of drop.Incubate at room temperature for 1-2 hr or overnight
in a cold room (better o/n).
7. Wash 3 X PBS/Tween/BSA, 5 min. at room temp.
9. Dilute the secondary antibody according to the manufacturer’s recommended
concentration into blocking solution and incubate with cells for 1 hour
in the dark at room temp in the moist chamber as above (e.g. 1 ml of antibody
per 100 ml of PBS-BSA-Tween).
10.Wash coverslips in PBS, 3 times, 5 min., room temp.
11. To stain DNA with DAPI, add 1 µl of 0.1 mg/ml stock into
10 ml PBS/Tween during the first wash.
13. Mount on a slide with DABCO antibleaching agent (6-8 ml of DABCO
solution, remove excess before sealing).
14. Seal coverslip edges with fingernail polish.
Method B (in suspension)
All steps in 15 ml conical polysteryne tubes.
1. Take 50 ml of fixed cell suspension, wash with 2 mls of PBS for 10 min. Concentrate by centrifugation (use as low speed as possible, on our centrifuge 2000 rpms).
2. Wash, PBS-3% BSA-0.1% Tween for 10 min, 3 times.
3. Concentrate cells in 100 ml of PBS/Tween/BSA, add primary antibodies (e.g. 20 ml of 6-11 B1). Incubate with rocking at 4oC overnight.
4. Wash 3 times with PBS-BSA-Tween.
5. Incubate with the secondary antibody (usually 1:100, 100 ml total volume, rocking, 1 hr at room temperature)
6. Wash 3 times with PBS. Add Dapi to the first wash if needed (see above). Concentrate cells in 100 ml, add 10 ml of DABCO, store at 4oC in dark.
7. To mount cells, take 4 ml of cell suspension and combine with equal volume of DABCO on a microscopic slide, cover with coverslip, seal edges with a nail polish.
Method C
ETOH/Triton fixation method (preserves natural shape of cilia) but gives increased fluorescence of cytosol
1. Handle cells as in the suspension method, wash with tris and concentrate down to 0.5 ml.
2. Add 4.5 mls of ice-cold fixative (prepare as follows: 5 mls of PHEM, 5 mls of 100% ETOH, 50 ml of triton X-100). Fix for 2’ , centrifuge at 2K for 3’
3. Remove supernatant, add 3 mls of 2% paraformaldehyde in PHEM, fix for 30’.
4. wash 1x with PBS and resuspend in 0.5 ml. Cell can be stored at 4oC
5. Stain cells as in the suspension protocol.
III Coating coverslips with poly-L-lysine
1.Wear gloves and handle coverslips with forceps.
2.Drop one at a time into 70% ethanol; let sit for 10 min.
3. Pour off 70% ethanol and add 95% ethanol. Remove and repeat with
99% ethanol. Lay out on 3MM paper to dry, about 30 min.
4. When dry, immerse in poly-L-lysine stock solution (Sigma cat# 8920)
diluted 1:10 with water (final concentration is 0.01%). Leave in lysine
solution at least 10 min. (coverslips can be stored in lysine solution
at 4°C for several months).
5. Remove coverslips with forceps, rinse in water and let dry on filter
paper.
Solutions:
M.W. For 1 liter:
PHEM 60 mM PIPES, pH 6.9 302.40 18.144 g
25 mM HEPES 238.30 5.96
g
10 mM EGTA 380.35 3.85 g
2 mM MgCl2x 6 H2O 230.30 0.41
g
(Add all components and adjust pH to 6.9 using 10 M NaOH).
2% Paraformaldehyde in PHEM:
To prepare paraformaldehyde, heat 10 ml of PHEM (in a small flask in the microwave, then add 0.2 g para-formaldehyde crystals to flask in the hood. Heat gently on stir plate if necessary. There should not be any crystals visible, but you don’t want the solution to boil).Prepare this solution first, as it takes about half an hour to dissolve.
PBS modified (phosphate buffered saline) modified, pH 7.2 (adjust pH
with NaOH).
M.W. 1
liter
130 mM NaCl 58.44 7.58 g
2 mM KCl 74.56
0.15 g
8 mM Na2HPO4x 7H2O 268.07
2.14 g
2 mM KH2PO4
136.09 0.272 g
10 mM EGTA 380.35
3.805 g
2 mM MgCl2x 6H2O 203.30
0.41 g
Extraction buffer:
15 mls of PHEM pH. 6.9
75 ml Triton X-100
7.5 ml leupeptin stock
10 ml E-64 stock
15 ml chymostatin stock
15 ml antipain stock
1.5 ml 10 mM taxol
(inhibitors and taxol optional but may help)
DABCO: 1,4-diazobicyclo-[2,2,2]-octane, dissolved at 100 mg/ml in 90%
glycerol in 1x PBS.
Note:
10% normal goat serum can be used as an additional blocking agent
if background is a problem.
References:
For method with drying - Gaertig, personal communication.
For method in suspension - Gaertig J., Cruz M.A., Bowen J., Gu
L., Pennock D.G. and Gorovsky M.A. (1995). Acetylation of lysine 40 in
a-tubulin is not essential in Tetrahymena thermophila. J. Cell Biol. 129,
1301-1310.