Yeast Transformation I

Method

Grow a 25 to 50 ml culture of yeast to log phase, OD₆₀₀ of 0.6 to 2.0. You need about 10-20 ml of culture for each transformation.

OD is fairly flexible, but log phase will give the highest transformation efficiency. If you are transforming a supercoiled plasmid – say from a maxi-prep – then even yeast scraped off a fresh plate will work ('fresh' means just grown, not stored in the refrigerator yet).

If you are trying to do a knock-out or have a small amount of DNA or impure DNA, then pay special attention to getting the yeast at the correct density. In this lab, we usually grow a small overnight culture (2-5 ml) and then seed a larger culture in the morning. Use 1-2 ml culture for each 50 ml media seeded. Let grow at least 4 hours.
Spin culture down in a 50-ml tube. Discard supernatant.
Wash yeast pellet in 10 ml of Li-TE. Spin. Discard supernatant.
Resuspend pellet in Li-TE. Use 100 µl per 10-ml culture. For example, use 1 ml for 100 ml of original culture.
Put 5 µl of DNA to be transformed and 5 µl of carrier DNA (calf thymus or salmon sperm DNA, 10 mg/ml) into a sterile microfuge tube. Add 100 µl of yeast and tap to mix.
Let sit at room temperature for 5 minutes.
Add 280 µl of 40% PEG in Li-TE to the transformation and invert to mix.
Incubate 45 minutes at 30°C.
Add 45 µl of DMSO, mix by inversion and heat shock for 5 minutes at 42°C.
Pellet yeast with short microfuge spin.
Aspirate supernatant. Change to a new sterile tip for each sample.
Resuspend yeast in 1 ml of sterile H₂O. Spin and discard supernatant as in step 11.
Resuspend yeast pellet in 0.5 ml of sterile H₂O and plate on appropriate selective media.
Incubate 2-4 days at 30°C.