Live and Fixed Cell Imaging of Yeast

For live-cell imaging, yeast were grown in liquid culture to mid-log phase (0.5–0.8 OD) and were prepared by adhering 200 µl of cells to 4-well glass-bottom dishes (CellVis) coated with concanavalin-A submerged with proper media for cell survival. For fixed imaging, yeast were grown in liquid culture to mid-log phase and 4% paraformaldehyde was added to the culture for 10 min. Cells then followed appropriate protocol (see Actin staining) and then were adhered to coverslips, again by using concanavalin-A. Coverslips were then mounted using soft mounting media (Invitrogen) and secured by using clear nail polish. Some fixed cells were also imaged using agarose pads, where 4 µl of fixed cells were added to a 1% agarose pad and media was allowed to diffuse into the agarose (Pringle et al. 1989 (link); Shin et al. 2018 (link)). All fluorescent microscopy was conducted on an inverted microscope (Leica DMI6000B) which had a spinning disk confocal unit (Yokogawa CSU-X1) with 100× 1.45 NA objective (Leica) with either a Evolve 512Delta EMCCD or with a Flash 4.0v2 CMOS camera. DIC imaging was conducted on this microscope as well as on a Leica De-Convolution Microscope (DMi8). Slidebook 6.0 or FIJI software were used for image analysis.