Yeast IF with Methanol/Acetone Dehydration

 

This is the protocol to use with the Botstein anti-actin rabbit antibodies.

1. Grow cells at the appropriate temperature to 5x10E6 in 5 mls YPD. Add 0.5mls 37 formaldehyde (best grade) and incubate on the roller at same temp. for 10 min.
2. Spin down cells 2Kx3min. and resuspend in 5mls 40mMKPO4 pH6.5/500uM MgCl2 + 0.5 mls formaldehyde. Incubate 1 hr at 30¡C (or previous temp).
3. Wash the cells two times in the previous buffer (no formaldehyde) and once in the same buffer containing 1.2M sorbitol. Be very gentle with the cells! Resuspend in 0.5 mls sorbitol buffer. The cells can be stored at this point overnight at 4¡C.
4. Zymolyase treat the cells with 30 ul 10 mg/ml Zymolyase (100T) at 30¡C for anywhere between 10-30 min or more. Examine cells on a phase microscope. When cells are dark and mishapen, you have gone way too far, if bright and refractile they probably need to be incubated longer. If they look good but are dull grey they are juuuusssttt right. I suggest this be done as a time course as it is the single most variable part of the procedure and unfortunately it is the most critical as well.
5. Wash the cells once in sorbitol buffer and suspend in 100-500ul of the same. BE VERY VERY GENTLE! Place on ice.
6. Coat the wells of teflon coated slides with 0.1 polylysine (>400,000 MW) in water for 10 min at RT. Spin this solution for 10 min in a microfuge immedietly before use and stay away from the bottom. In general do high speed spins of all solutions that go on the wells right before they are used. Incubate the slides in a moist chamber.
7. Wash the wells 4-5 times with clean spun water and dry. These can be prepared in advance if kept dust free.
8. Spot 20ul cell suspension on the wells and incubate at RT for 10 min. I suggest you do each sample in duplicate. Aspirate off most of the liquid but not all (do not dry the slides) and plunge, this is important, plunge the slide into a coplin jar that is surrounded by dry ice for 6 min. then plunge into another coplin jar that contains acetone and is also surrounded by dry ice for 30sec. It is important that the jars and contents are down to temperature so place them on the dry ice an hour or two before this step. Change your methanol and acetone frequently and do not use sharpies to mark your slides.
9. When you remove the slide from the acetone immediatly place against a slanted, flat, warm, clean surface so that the acetone evaporates without the creation of condensation. I like to use the top of a warm water bath. It can help to wick away the excess acetone from the bottom edge with a kim wipe as the slide dries. This process should only take a few seconds and you should be wearing gloves as finger grease will make a mess of things. I also wash the talcum off the gloves.
10. Block the cells in PBS pH7.4/ 0.5 BSA/0.5 ovalbumin. High speed spin this solution. Sometimes the inclusion of 0.5 Tween 20 can help the specificity of the antibody and should be included in this block solution. Be fore warned that the tween reduces the hydrophobicity of the teflon and sample mixing can occur. I find that if you outline the wells with a sharpy after the acetone step you can prevent mixing. Incubate for 15 min at RT. The cells can be incubated for overnight at this step.
11. Incubate the cells in block containing antibody at the appropriate dilution (determine experimentally). A dilution series of 1:100 to 1:10,000 should cover it. Incubate at RT for 1 hr or longer, depending on the antibody. Sometimes an overnight incubation is helpful. Jon''s anti-actin Guinea pig antibodies from animal#1 are best at 1:6,000, #2 and #3 are best at 1:2,000.
12. Wash the cells 4 x 5 min with the block solution. Sometimes longer incubation times may lower the background but this is usually sufficient.
13. Incubate in secondary antibody conjugate diluted in block solution for 1 hr at RT. You may need to determine the concentration best for your secondary antibody as well. The Cappel anti-Guinea Pig-FITC is best at 1:800-1:1,000.
14. Wash as before. Aspirate most of last wash off the cells but do not dry and mount immediatly. Mount by slopping the mount solution over the slide (no bubbles) and gradually laying the cover slip down by the long axis. Lay paper towels over the slide and squeeze out the excess mount. While holding the slide down with the fingers of one hand, clean the slide with a kim wipe. Finally seal the edges of the slide with nail polish. I find that Sally Hansen''s "HARD AS NAILS" works best. Allow to dry and store at -20¡C until you are ready to view the results. Before you place the slide on the scope, completely clean any residual mount from the slide, this stuff is bad for the objectives.

Reagants

Zymolyase

Use the good stuff 100T. Dissolve in the sorbitol containing phosphate buffer described in the protocol, spin for 10 min in a microfuge, remove to a fresh tube and quick freeze on liquid nitrogen. Store at -80¡C. Do not repeatedly freeze and thaw, if you do always quick freeze.

 

Polylysine

Buy the good stuff from sigma, >400,000MW. Dissolve as a 1 stock solution in good water. Quick freeze in aliqoutes and store at -80¡C. Same applies as for zymolyase.

 

DAPI

Stock solution should be at 1 mg/ml in water and stored at -20¡C.

 

Mount

Dissolve 100 mg p-phenylenediamine in 10 mls PBS, adjust the pH to above 8.0 with 0.5 M Na Carbonate buffer (pH 9.0) and bring the volume to 100 mls with glycerol. Add Dapi to 50 ng/ml. Mix thoroughly and store at -20¡C. It turns brown when it is bad.