Yeast Immuno-fluorescence

Formaldehyde fixing followed by zymolyase treatment

1. Grow a 5 mL culture in YPD or selective media overnight.

2. Dilute culture to OD600 0.2 in 50mL of fresh YPD and grow to OD600 1.7.

3. Add 37% formaldehyde to a final concentration of 3.7% (1/10 of culture volume) directly to the culture and incubate with gentle shaking for 60 minutes at 30˚C.

4. Transfer cells to a 50 ml conical tube and pellet by centrifugation (3 minutes @ 2000 rpm).

5. Wash the cells twice with 40 ml of 100mM KPO4 pH 6.5 and once with 40ml of 100mM KPO4 pH 6.5/1.2 M sorbitol (P solution).

6. Re-suspend the cells in 10ml of P solution – at this point they can be stored at 4˚C for a couple of weeks.

7. Resuspend the cells and take a 1ml aliquot into an eppi. Add 2ul of b-ME and incubate at 30˚C for 10 minutes. Add 50 ul of 10mgr/ml zymolyase solution and incubate for 40 minutes at 30˚C or until digestion is sufficient. Check digestion by phase contrast microscopy: cells should be a dark, translucent gray. Bright cells are insufficiently digested; pale gray cells with little if any internal structure (ghosts) are over-digested. After digestion of the cell walls treat the yeast delicately to avoid lysing them.

8. Wash the cells three times with 2ml of P solution.

9. Resuspend the cells in 1ml of P solution with 0.5% NP-40 and incubate at 30˚C for 10 minutes to permeabilize the cells (no shaking or mixing). When the cells are pelleted after this step they will be difficult to see! Wash twice with 1ml of P solution and resuspend the cells in 0.5ml of PBS-BSA (1% BSA – filtered sterilized).

10. Place 50 ul of the cell suspension into each well of a poly-lysine coated slide. Let the cells settle for 20 minutes, then aspirate excess and let the slides dry at 60˚C for 30 minutes.

11. Re-hydrate and block the cells by adding 50 ul of PBS-BSA and incubating at RT for 15 minutes. Repeat once.

12. Add 50 ul of the 1st antibody diluted in PBS-BSA. Leave incubating o/n at 4˚C.

13. Wash the cells 5 times with 50 ul of PBS-BSA. Incubate 5’ at RT with 50 ul of PBS-BSA and then wash again three times.

14. Add 50 ul of the secondary antibody diluted 1:1000 in PBS-BSA. Incubate at RT for 2 hours in the dark.

15. Wash the cells 5 times with 50 ul of PBS-BSA. Incubate 5’ at RT with 50 ul of PBS-BSA (in the dark) and then wash again three times.

16. Wash cells once with 50 ul of PBS-BSA with Hoechst to stain the DNA (1ul of 10 mg/mL stock solution of Hoechst into 1mL of PBS-BSA).

17. Wash the cells 3 times with 50 uul of PBS-BSA and 3 times with 50 uul of PBS.

18. Add a small drop of mounting media to each well and put on the cover slip. Remove excess of mounting media and seal with clear nail polish.


100mM KPO4 pH 6.5 (0.1L) 10mL of 1M KPO4 pH 6.5

100mM KPO4 pH 6.5 1.2 M sorbitol (0.1L) 10mL of 1M KPO4 pH 6.5 21.87 grams of sorbitol

Zymolyase:10 mg/ml in100mM KPO4 pH 6.5 1.2 M sorbitol (1mL) 10 mg of zymolyase in 1 mL of in100mM KPO4 pH 6.5 with1.2 M sorbitol. Vortex and let it sit on the bench for 5 minutes, then spin in micro-centrifuge at top speed for 5 minutes and use clear supernatant.

1% BSA in PBS (50mL) 0.5 g of BSA in 50 ml PBS buffer for 10 mg/ml final concentration. Filter and stored at 4ºC.

Mounting media (10mL) 0.5% p-phenylenediamine (50 mg) 20mM Tris pH 8 (200 ulL of 1M Tris pH 8) 90% glycerol (9 mL) Dissolved the p-phenylenediamine in 1mL of 200mM Tris pH 8, and then add the glycerol. Store at –20˚C in 1mL aliquots.

Slide Preparation

Prepare Teflon-masked slides by adding 10 ulL of 1 mg/mL polylyside solution onto each well. Incubate 1 minute, wash 3 times with water and allow slides to air-dry for 10 minutes before use.