Manipulation and Analysis of Cell Cycle-Dependent Processes in Budding Yeast

We study how dividing cells faithfully pass on their chromosomes during mitosis, focusing on the molecular machines and mechanisms that ensure accurate chromosome segregation. We synchronize cells to study molecular processes that change with the cell cycle. Without these methods, key changes would be hidden in an unsynchronized cell population.

Compared to other synchronization methods, alpha-factor arrest in BAR1 mutants provides a cleaner, reversible, G1 arrest, allowing us to track an entire yeast culture progressing synchronously through the cycle. Our work reveals dynamic protein localization and activity changes throughout the cell cycle, shedding light on key mitotic processes, like chromosome segregation and spindle maintenance. To begin, inoculate yeast into 25 milliliters of YPAD media, and incubate overnight to reach an optical density at 600 nanometers between 0.5 and 2.0.

Dilute the yeast cells to an optical density at 600 nanometers of 0.5. Add alpha-factor to the culture to reach a final concentration of one microgram per milliliter. At 2.5 to 3.5 hours after alpha-factor addition, count the percentage of non-budded shmooed cells under a microscope to assess the cell arrest.

Then, spin down the culture in a centrifuge at 3, 000g for three to five minutes at 23 degrees Celsius. Carefully pour off the supernatant to remove the alpha-factor. Resuspend the cell pellet in 25 milliliters of YPAD containing 1%dimethyl sulfoxide to wash the cells.

Transfer the cell pellet to a fresh tube and repeat the washes twice more for a total of three washes to remove any residual alpha-factor. Next, add YPAD to the washed cells to bring the final volume to 25 milliliters, and transfer the suspension into a new flask for further incubation or use. Collect the zero-minute time-point sample immediately after release and fix the sample.

At 60 minutes post-release, assess synchrony of the cell population under a light microscope. If desired, add alpha-factor again at 60 minutes after release to a final concentration of one microgram per milliliter to block progression into the next cell cycle. Continue collecting time-point samples every 15 minutes up to 180 minutes or for as long as needed.

To fix the yeast cells, centrifuge one milliliter of culture for one minute at maximum speed. Aspirate the supernatant completely. Then, resuspend the pellet in 500 microliters of fixative solution and incubate at 23 degrees Celsius for two to 15 minutes.

After centrifuging the fixed cells, aspirate the supernatant and resuspend the pellet in 500 microliters of 0.1-molar potassium phosphate buffer at pH 6.4. Before imaging, centrifuge the fixed cells at 23 degrees Celsius for one minute at maximum speed. Once the supernatant is aspirated, resuspend the pellet in 10 to 100 microliters of Triton, DAPI, and sorbitol solution.

Pipette approximately 0.8 microliters of the stained cell suspension directly onto the center of a clean coverslip. Use a pipette tip to gently spread the droplet into a circular area approximately one square centimeter in size. Place the coverslip onto a microscopy slide.

Using a Kimwipe, gently press around the edges of the coverslip to evenly distribute the sample. Then, seal the edges of the coverslip with nail polish to secure the sample. Take the prepared slide to a microscope equipped with a 60 times magnification, 1.42 numerical aperture oil immersion objective, and a red, green, blue, and far red laser and filter set.

Focus the microscope on the yeast cells adhered to the coverslip. Once focused, tune the exposure settings for each fluorescence channel to achieve a signal-to-noise ratio of at least three-to-one. Adjust the acquisition settings to collect 14 to 20 z-stack images with each slice spaced 0.2 micrometers apart, covering a total z-depth of two to four micrometers.

On microscopes equipped with post-processing modules, select Deconvolution and Quick Projection options for each z-stack image. Acquire z-stacks of the sample, capturing enough images to record approximately 100 to 200 yeast cells for each experimental condition or time point. For analysis, open the projected images in ImageJ software.

Adjust the brightness and contrast of each fluorescence channel to enhance visibility. To analyze cell cycle progression, count 100 cells for each time point and classify them as containing one or two nuclei. To calculate the percentage of cells showing Stu2-GFP puncta, standardize the green channel brightness settings across all images.

Count cells showing Stu2-GFP puncta that co-localize with Spc110-mCherry puncta as positive for kinetochore localization. Next, to quantify the intensity of protein puncta, use the freehand selection tool to outline the region of interest around the signal. Add the selection to the Region of Interest Manager by pressing T or choosing the option from the ROI Manager menu.

In the ROI Manager, click Measure to calculate puncta intensity. To visualize changes over time, plot the average intensity with 95%confidence intervals for each time point. Count approximately 100 cells per condition.

In G1-arrested cells, Stu2-GFP localized to a single spindle-pole proximal punctum with additional dispersed signal along cytoplasmic microtubules. At 60 minutes after release, Stu2-GFP localized as two puncta adjacent to duplicated spindle poles marked by Spc110-mCherry. By 90 minutes, during anaphase, Stu2-GFP appeared both near the spindle poles and along microtubules spanning the single axis.

Following mitotic exit, at 120 minutes, Stu2-GFP reappeared on astral microtubules in the cytoplasm. Quantification revealed that Stu2-GFP intensity near spindle poles increased during early mitosis, peaking before anaphase and declined thereafter. The percentage of binucleate cells peaked at approximately 90 minutes, indicating synchronized entry into anaphase.