Team:UCSF/Protocols

Cloning Overview

We cloned our protein of interest into the display plasmid (pCTCON2) by either using the restriction sites NotI-HF and BamHI-HF or by using homologous recombination. In the case of using restriction sites, we would first PCR the desired insert protein. We would then restriction digest the PCR product and the plasmid pCTCON2, separately of course. We would then phosphatase the pCTCON2. After we restriction digested both, we would PCR purify both the PCR product and the plasmid. We would then set up a ligation reaction. After we ligated the PCR product and plasmid, we would transform it into E. Coli Mach I cells. If successful, we would miniprep and transform into EBY100 Yeast cells.

In the case of using homologous recombination, we would make primers with the homologous region. We would PCR the desired protein and the plasmid with the correct primers. Afterwards, we would PCR purify and then transform it into EBY100 Yeast cells.

Gal Induction and Microscopy

The growing of yeast cultures for experiments is a two day process. First, cultures are grown in S-Raff media (5ml) overnight. 24 hours prior to microscopy, the yeast are induced with galactose by taking yeast from S-Raff and placing them in S-Gal(1%) media, with a final OD of 0.001 for both tube cultures and plated cultures. For experiments using more than one strain of yeast growing in one culture, they are added together at the induction stage, with an OD of 0.001 for each strain.

Microscopy was our main method of obtaining data. Microscopy was done using 96-well plates, with tube-grown cultures being plated prior to microscopy at an OD of 0.2. Wells containing tube-grown yeast were coated with Con-a (to help yeast stick to the plate) 30 minutes prior to observation, and then spun down for 1 minute at 3000 RPM. The wells were then observed at (60)x magnification by differential interference contrast (DIC) microscopy, with fluorescence also being visualized in some experiments using yeast with GFP or mCherry.