Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live cell microscopy allows researchers to study fission yeast nuclear dynamics in mitosis and meiosis in real time. The strength of this method comes from studying nuclear processes in living cells under normal physiological conditions which eliminates the need for toxic fixatives and stains. This technique addresses questions related to protein timing, stability, and mobility that may not be amenable to genetic or biomechanical manipulation.

It is critical to pay close attention to the health and fitness of fission yeast cells before imaging. Always examine morphology and growth characteristics to ensure consistency of results across experiments. This protocol shows the relatively simple way to prepare microscope slides that when mastered correctly can enhance the consistency and reproducibility of prolonged live cell imaging.

To begin pick up cell matter from cryogenic wake up plate, Streak it onto YES plates, and incubate them at 25 degrees Celsius or 32 degrees Celsius depending on the yeast genotype to wake up fission yeast strains from cryogenic preservation. Then use a sterile loop to pick cells from individual colonies in the awakened fission yeast strains and inoculate them into test tubes containing three milliliters of YES liquid medium. Place the tubes in a shaker at 150 to 220 rpm to grow at 25 or 32 degrees Celsius overnight to late-log phase with a desired optical density reading of 0.7 to 1.0.

Transfer 10 microliters of the culture to a microscope slide. Put a coverslip on and place it under a microscope to check for proper cell morphology and nutritional state. To set up a microscope slide for mitosis or meiosis analysis first add two grams of agarose in the 500-milliliter flask containing 100 milliliters of either minimal medium plus supplements for mitosis or liquid sporulating medium for meiosis.

Warm the agarose solution in a microwave oven at 60%power in 10-second increments or place the beaker in a 55 degrees Celsius water bath for 10 minutes. Swirl the solution to ensure efficient melting. Set up two microscope slides on a pipette tip holder with the top slide resting on two stacks of lab tape at both ends as support making a cross shape.

Adjust the distance between the two slides to greater than two millimeters in order to form a thick pad in the following steps for prolonged imaging. After cooling the molten agarose for one minute at room temperature, remove the top slide and use a wide bore pipette tip to dispense 50 to 100 microliters on the bottom slide to make a spot. Before the agarose cools down place the top slide on top to generate a spread pad of about one and 1/2 to two centimeters in diameter.

Optimal live cell imaging is critical for subsequent data processing steps. Ensure to eliminate any air baubles in the molten agarose and create a thin pad for imaging periods longer than four hours. To examine mitotic events grow cells from starter cultures in either liquid EMM or PMG plus supplements overnight.

Transfer one milliliter of the culture into a cuvette, and measure on the spectrophotometer at a wavelength of 595 nanometers. The cell growth is considered mid-log phase when the optical density reaches 0.4. Then centrifuge one milliliter of the cell suspension at 1, 375 times g for one minute.

Remove the supernatant and resuspend the cell pellet in minimal medium plus supplements to a final volume of 100 microliters. To image meiotic events grow cells from starter cultures in minimal medium plus supplements overnight. Cell growth is considered late-log phase when the optical density at 595 nanometers is between 0.7 and one.

Next, from the late-log cultures obtain 500 microliters of each mate type strain, H negative and H positive, and combine to make a one milliliter cell suspension. Centrifuge the cells at 1, 375 times g for one minute. Remove the supernatant and resuspend the pellet in one milliliter of liquid maltose extract.

Repeat the maltose extract wash three more times to ensure efficient nutrient removal. After the last maltose extract wash resuspend the cells in one milliliter of maltose extract and transfer the mixture to a 50-milliliter flask containing nine milliliters of maltose extract. Incubate for 12 to 16 hours at 22 to 25 degrees Celsius on a minimal rotation speed between 50 and 100 rpm.

The appearance of many round fission yeast clumps resulting from abundant self-flocculation indicates efficient mating. Next, take a one-milliliter sample of the mating culture and centrifuge at 1, 375 times g for one minute. Remove 750 microliters of the supernatant and resuspend the cells in the remaining supernatant.

Vortex vigorously for five seconds to disrupt the clumps. Dispense 20 microliters of either a mitotic or meiotic cell suspension on a 2%agarose pad. Remove any excess medium by inverting the slide and putting it on the top of the lint-free paper towel for two to three seconds.

Flip the slide and gently place a glass coverslip on the top of the pad, ensuring not to generate air bubbles. To create a cell monolayer rotate the coverslip clockwise with the index finger for one full turn for mitosis cells or two full turns for meiosis cells. Ensure that the cell matter disperses across the agarose pad allowing for better separation of single cells or asci.

With the aid of a small wooden stick dispense molten hydrocarbon sealant along the edges of the coverslip to seal each agarose pad. Once the agarose pad is sealed place it on the microscope stage. Switch on the temperature chamber and set it to the desired temperature.

Place a plate with a wet paper towel into the chamber to control the humidity of the microscope system. Let it equilibrate for 10 to 15 minutes at the appropriate imaging conditions. This allows any remaining air bubbles to dissipate and any last-minute agarose shifts to occur.

Use the 40X objective to find appropriate fields of view to image. Switch to the 60X objective to begin data acquisition. In the software that controls microscope function choose the microscope filter sets that best match the fluorophores under observation.

Adjust the excitation wavelengths to match the fluorophores used, employ the lowest excitation power that produces a consistent signal, and use exposure times of four to eight hours to generate acceptable yet quantifiable and reproducible imaging data. Collect at least six acquisition time points every hour. In the image collection software select at least one fluorescence channel from which to acquire data and employ a Z-stock comprised of 13 sections with 0.5 micrometer spacing.

In the Fiji software upload a deconvolved image by selecting the Bio-Formats Importer feature under the plug-ins menu. In the Import Options pop-up window, select Hyperstack, Default Color mode, check Autoscale and press the OK button. Ensure the displayed window has the correct number of fluorescence channels and time frames by scrolling sideways on the respective bars at the bottom.

Save as a Tiff file which does not compress data. Then under the Analyze menu use the Set Measurements option to choose different parameters. For example:Area, Min max gray value, Integrated density, Mean gray value, Median, Stack position, Perimeter, and Display label.

Select Color and click on the Channels Tool. In the Channels pop-up window check the Color Channel for which intensity or area will be measured. Select Image, then Type and click on 32-bit.

Choose the Adjust and Threshold features under the Image menu. Check the Dark background box, select the Default method, and pick Red to overlay the signals of interest. Use the wand tool to highlight each structure of interest and press the letter T on the keyboard to add the selected ROI to the pop-up ROI manager window.

Next, open the zipped ROI folder to load the ROI manager and click on each ROI identifier in the left side panel. Select Measure from the Analyze menu and repeat this command on each ROI identifier to quantify the objects of interest in all slices of the image stack. Save the results as a CSV file for statistical analysis.

In this study if cells starve due to nutrient limitation or overgrowth, they will show excess vacuoles and decreased cell size. Logarithmic cells show active DNA replication and cell division, as well as pan-nuclear Tos4-GFP expression and Sad1-DsRed foci separation. Failure to mate, as is the case when cells are not sufficiently starved of nitrogen, will prevent them from entering meiosis.

A pair of fission cells undergoing karyogamy are shown. Robust flocculation of the mating cell suspension increases cell-to-cell interaction and thus indicates successful mating and efficient meiotic induction. Zygotic asci take multiple forms, including the zig-zag and banana cell shapes.

In mitosis as cells transition from metaphase to telophase, the first change involves a contraction of nuclear size in metaphase, while the second shows nucleus splitting during anaphase. Besides sharing some segregation dynamics with mitotic cells, meiotic cells exhibit nuclear oscillation during homologous recombination and further reduction of nucleosides at the end of anaphase II.It is the responsibility of researchers to ensure that experiment parameters can be reproduced across experiments, and that the collected data is fit for downstream analysis. Live cell imaging has allowed investigators to examine in close detail the mechanics of nuclear division in mitosis and meiosis.

As fluorescent tags and microscope capabilities improve the number and type of processes we can observe will increase.