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.

Reagents:

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.