Yeast Immunofluorescence

Note this protocol is a modified version of the protocol from Mark Rose in the CSH Yeast Genetics Course Manual.

    1. Grow cells at the appropriate temperature to 5x10E6 in 5 mls YPD. Add 0.5mls 37% formaldehyde (best grade) for final concentration of 3.7% PFA and incubate on the roller at same temp. for 10 min.
    2. Spin down cells 2,000 rpm, 3min and resuspend in 5mls 40mMKPO4 pH6.5/500uM MgCl2 + 0.5 mls 3.7% 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 a 170µm thick (number 1.5) 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 onto the coverslips right before they are used. Incubate the coverslips on top of parafilm in a moist chamber.
    7. [notice]DO NOT use a multi-well teflon coated slide. Once a coverslip is in place, the teflon spacer will place the cells too far from the objective lens, introducing unnecessary aberration into the image. Instead use individual coverslips[/notice]
    8. Wash the coverslips 4-5 times with clean spun water and dry. These can be prepared in advance if kept dust free.
    9. Spot 50ul cell suspension on the coverslips (for 18mm diameter, more for bigger size – may need to titrate)  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 let the slides dry! Block the cells in PBS pH7.4/ 0.5% BSA/0.5% ovalbumin/0.5% Tween 20. High speed spin this solution. Be fore warned that the tween reduces the hydrophobicity, so less volume can fit onto the top of the coverslip before the surface tension no longer holds it there. Incubate for 15 min at RT. The cells can be incubated for overnight at this step.
  1. 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.
  2. Wash the cells 4 x 5 min with the block solution. Sometimes longer incubation times may lower the background but this is usually sufficient. Do not let the cells dry!
  3. 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.
  4. Wash as before. Aspirate most of last wash off the cells but do not dry and mount immediately.
  5. Stain with DAPI. Make a 50ng/ml solution of DAPI in ddH2O and incubate the coverslips in this solution for 2 minutes.
  6. Rinse in large volume (500ml beaker full) of ddH2O to remove unbound DAPI and traces of salts.
  7. Mount by placing about 5-10µL of mounting media on a clean slide and gradually laying the cover slip down to prevent any bubbles. Let the mass of the coverslip squeeze out any excess mounting media. Finally seal the edges of the slide with nail varnish. 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. Only mount 1 coverslip per slide.


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.
Buy good stuff from sigma, >400,000MW. Dissolve as a 1% stock solution in good water. Quick freeze in aliqoutes and store at -80°C.
Stock solution should be at 1 mg/ml in water and stored at -20°C.
Dissolve 500 mg p-phenylenediamine in 1 ml PBS, add 1 ml 200mM Tris-HCl pH8.8 to buffer and bring the volume to 10 mls with glycerol.  Mix thoroughly and store at -20°C. It turns brown when it is bad. Key to using PPD as anti fade is pH. Too low and your fluorescence will not be protected from bleaching.