Yeast Immunofluorescence

Courtesy of Sue Biggins

Cell Fixation

  1. Grow cells to mid-log phase (O.D.(600) = 1, or 2E7 cells/ml), usually about 5 ml of culture per sample to accomodate the loss of cells through all of the wash steps, however you can use fewer if you are limited for cells. Make sure you grow an isogenic strain that lacks the antigen of interest, if available, as a control.

  2. To fix cells, add formaldehyde to 3.7% final concentration, and incubate on roller drum at 23oC. You should do a fixation time course the first time you localize a protein, such as 10, 20, 40, 60 minutes. For tubulin, fix cells for 1 hour. For HA-tagged proteins, cells should be fixed for as short a time as possible. Fixation time is also strain dependent. Some cells require longer fixation or they become extremely fragile and sensitive ot osmotic shocks. Excessive fixation may result in uniform staining.

  3. Harvest cells by centrifugation and wash them twice in 5 ml 0.1 M phosphate buffer.

  4. Wash cells once in 5 ml 1.2 M sorbitol.

  5. Wash cells once in 5 ml phosphate buffer.

  6. Read step 7 before continuing! Resuspend washed cells in 1 ml 1.2 M sorbitol and add 5 ul beta-mercaptoethanol and 10 ul 5 mg/ml zymolyase 100T in 1.2 M sorbitol. To make the zymolyase stock solution, dissolve 50 mg zymolyase in 10 ml 1.2 M sorbitol. Spin in microcentrifuge at 14000 rpm for 5 minutes to pellet insolubles. You can freeze the zymolyase solution and use it multiple times successsfully.

  7. Put the cells on a roller drum at 23oC or 30oC. Spheroplasting times can differ substantially, so carefully monitor spheroplasting every 15 minutes or so. You can do this either by.

    1. Phase contrast microscopy. Under the microscope, the cells should be a dark transluscent gray. Bright (refractile) cells are insufficiently digested, whereas ghosts (pale gray cells with little, if any, internal structure) have been overdigested.

    2. Sensitivity to SDS lysis. Put an aliquot of cells (usually 20ul is enough) into a 1 ml aliquot of 1% SDS and read the O.D.(600). Be sure to take a zero time point, and the initial reading should be at least 0.1 in order to watch the decline in O.D.(600) readings as the cells are spheroplasted. Let the O.D.(600) drop 80-90% of the initial reading. It is not necessary to monitor every sample during the time course, but you should do a couple. Alpha factor arrested cells spheroplast much more quickly than others, so a sample treated with alpha factor should be monitored.

  8. Harvest cells by centrifugation and wash once with 5 ml 1.2 M sorbitol. BE VERY GENTLE ONCE CELLS ARE SPHEROPLASTED.

  9. Resuspend cells in 1 ml or less of 1.2 M Sorbitol. Spheroplasted cells can be stored at 4oC for a few days.

Slide Preparation and Antibody Staining

  1. Make sure you have a coplin jar containing 100% methanol at -20oC prepared well ahead of time.

  2. Prepare a solution of 0.1% polylysine (w/v 400,000mw) by diluting a 1% stock solution in distilled H2O. The stock solution should be stored at -20oC.

  3. Place 15 ul of polylysine in each well of the immunofluorescence slide. Let sit for 1-3 minutes. Wash three times with distilled H2O, and air dry befor proceeding.

  4. Place 15 ul of cells into each well and let settle for 15 minutes. Make sure you have a well for all of the controls (no secondary antibody control, Ab dilution series controls, etc) as well as the experimental samples.

  5. Aspirate the supernatant from the cells and immediately plunge the slide into the cold methanol. To keep the methanol cold, you can put it on dry ice in an ice bucket. Incubate for 6 minutes.

  6. Remove slide and submerge in room temperature acetone for 30 seconds.

  7. Quickly air dry slide and immediately add 40 ul of block solution. Don't let cells dry out at any pint from now on until they are mounted.

  8. While the cells are blocking, prepare primary antibody solutions. Dilute antibody with blocking solution. Do a series of three-fold dilutions the first time you are localizing a protein and/or using an antibody. Ten fold dilutions can miss the optimal concentration between high background and no staining. Use only affinity purified polyclonal antibodies or monoclonal antibodies as most rabbits and mice have high reactivity to yeast proteins. If possible, use an isogenic control that lacks the antigen of interest. Spin the antibody in the microcentrifuge for 10 minutes at 14,000 rpm to pellet insolubles.

  9. Remove blocking solution and antibody solution to cells. Put slides in a humid chamber. You can wet some paper towels and place them in the bottom of a tupperware dish Incubate at room temperature overnight. Shorter incubations (45-120 minutes) are sometimes okay, but overnight seems to be the best.

  10. Wash cells by aspirating off primary antibody, adding blocking solution, and incubating for 15 minutes. Wash at least four times in this manner.

  11. Add diluted secondary antibody. For most, a 1:500 dilution is good, but this should also be titrated when done for the first time. Microcentrifuge secondary antibody at 14,000 rpm for 10 minutes prior to use to pellet insolubles. When you are double staining use the FITC antibody for the weaker signal and the Rhodamine for the stronger signal because your eyes tend to see FITC better.

  12. Incubate at room temperature for 1-2 hours in the dark in a humid chamber to minimize bleaching of the fluorochrome.

  13. Wash cells as in step 10.

  14. Wash cells twice with phosphate buffer prior to DAPI staining. To stain with DAPI, prepare a 1 ug/ml DAPI-phosphate buffer solution from a 1 mg/ml DAPI stock solution. Add DAPI to cells and incubate for 5 minutes. Wash once with phosphate buffer, then aspirate wells and air dry before mounting.

  15. Mount cells by adding a dollop of mounting media to the end of the slide a putting a coverslip on top. gently push on the coverslip to make sure the media covers the entire slide and all air is expelled. Blot the excess media with a paper towel, and seal the edges with nail polish.

  16. Wash the slide with distilled H2O and dry with a kimwipe. Slides can be stored for several weeks at 4oC in the dark.


10X PHOSPHATE BUFFER

TO MAKE 100 ml: 

  • 83.4 ml 1 M K2HPO4 stock solution
  • 16.6 ml 1 M KH2PO4 stock solution
pH should be 7.5.

BLOCK SOLUTION

TO MAKE 10 ml: 

  • 1 ml 100 mg/ml BSA stock solution
  • 500 ul 10X Phosphate buffer stock solution
  • 300 ul 5 M NaCl stock solution
  • 1 mg NaN3
pH should be 7.5. Block solution can be kept in fridge for extended periods of time.

MOUNTING MEDIA

TO MAKE 100 ml: 

  • 100 mg phenylenediamine
  • 10 ml phosphate buffer
  • distilled H2O to 100 ml
Dissolve phenylenediamine in phosphate buffer. bring the pH to 9.0. Then add H2O. Store mounting media at -20oC in the dark (wrap bottle with foil). Old mounting media turns brown and autofluorescent on the rhodamine filter.