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

1. Media Preparation^15,16

1. Stock solutions
   1. Prepare a 50x salt stock solution by adding 52.5 g of MgCl₂·6H₂0, 0.735 g of CaCl₂·2H₂0, 50 g of KCl, and 2 g of Na₂S0₄ in a bottle containing 1 L of distilled water. Mix the solution thoroughly and sterilize by filtration. Place at 4 °C for long-term storage.
   2. Make a 1,000x vitamin stock solution by mixing 1 g of pantothenic acid, 10 g of nicotinic acid, 10 g of inositol, and 10 mg of biotin in a bottle containing 1 L of distilled water. Mix the solution well and sterilize by filtration. Keep at 4 °C for long-term storage.
   3. Prepare a 10,000x mineral stock solution by adding 5 g of boric acid, 4 g of MnSO₄, 4 g of ZnSO₄·7H₂O, 2 g of FeCl₂·6H₂O, 1 g of KI, 0.4 g of molybdic acid, 0.4 g of CuSO₄·5H₂0, and 10 g of citric acid in a bottle containing 1 L of distilled water. Mix the solution thoroughly and sterilize by filtration. Place at 4 °C for long-term storage.
   4. Make individual nutrient stock solutions by adding 7.5 g of either adenine, leucine, histidine or lysine into a bottle containing 1 L of distilled water. Use only 3.75 g to make the uracil solution. Sterilize solutions by autoclaving.
      NOTE: Over time, uracil precipitates out of solution. To bring into the solution again, warm in the microwave at 60% power in 10 s increments or place in a 55 °C water bath for 20 min. Swirl the warm solution until all uracil clumps disappear.
2. Yeast extract plus supplements (YES)
   1. In a 2 L flask, add 1 L of distilled water, 5 g of yeast extract base, 30 g of glucose, and 225 mg each of adenine, uracil, L-histidine, L-leucine, and L-lysine. Add 20 g of agar to make the solid medium.
   2. Use a magnetic stir bar to completely dissolve ingredients into the solution. Agar will melt after the medium is sterilized by autoclaving.
      NOTE: For convenience, ready-to-use YES powder is also commercially available.
3. Edinburgh minimal medium (EMM) and pombe glutamate medium (PMG)
   1. In a 2 L flask, combine 1 L of distilled water with 3 g of potassium hydrogen phthalate, 2.2 g of Na₂HPO₄, 5 g of NH₄Cl, 20 g of glucose, 20 mL of salt stock solution, 1 mL of vitamin stock solution, and 0.1 mL of mineral stock solution. Replace NH₄Cl with 2.2 g of L-glutamic acid monosodium salt to prepare PMG. To make the solid medium, add 20 g of agar.
   2. Using a magnetic stir bar, dissolve ingredients thoroughly. Agar will melt after the medium is sterilized by autoclaving. Before use, add nutrient stock solutions as necessary. For every 1 L of medium, add 15 mL each of adenine, leucine, histidine, and lysine. Use 30 mL for uracil, since it is less concentrated than the other nutrient solutions.
      NOTE: For convenience, ready-to-use EMM and PMG powders are also commercially available.
4. Malt extract (ME)
   1. Use a 2 L flask to mix 1 L of distilled water with 30 g of malt extract and 225 mg each of adenine, uracil, histidine, and leucine. Adjust the pH to 5.5. Include 20 g of agar to make the solid medium.
   2. Dissolve components into the solution using a magnetic stir bar. Agar will melt after the medium is sterilized by autoclaving.
5. Sporulation agar with supplements (SPAS)
   1. In a 2 L flask, add 1 L of distilled water, 10 g of glucose, 1 g of KH₂PO₄, 1 mL of vitamin stock, 45 mg each of adenine, uracil, histidine, leucine, and lysine hydrochloride. Add 20 g of agar to make the solid medium.
   2. Use a magnetic stir bar to thoroughly combine ingredients. Agar will melt after the medium is sterilized by autoclaving.

2. Fission Yeast Culture^15,16

1. Use YES solid medium to wake up fission yeast strains from cryogenic preservation. Depending on the temperature requirements of each strain, incubate at either 25 °C or 32 °C for 3-5 days.
2. When colonies are visible, prepare a starter liquid culture. Pick cells from individual colonies and inoculate them into test tubes containing 3 mL YES liquid medium. Grow at the appropriate temperature to the mid- or late-log phase.
   NOTE: For proper aeration, use tubes or flasks with volumes at least 5x greater than the intended liquid culture volume. Shaking speeds between 150-220 rpm are customary for routine culture growth.
3. Dispense 250-500 µL of starter culture into a 50 mL flask containing 9.5-9.75 mL of either YES, EMM or PMG plus appropriate supplements. Allow cells to grow to the desired density and check under the microscope for the proper cell morphology and nutritional state.
   NOTE: To store awoken strains for up to a month, seal YES plates with paraffin tape and place at 4 °C.

3. Sample Preparation²

1. Microscope slide setup
   1. Add 2 g agarose in a 500 mL beaker containing 100 mL of either minimal medium plus supplements (mitosis) or liquid SPAS (meiosis). Warm the agarose solution in a microwave oven at 60% power in 10 s increments or place in a 55 °C water bath for 10 min. Swirl the solution to ensure efficient melting. Prepare agarose slide-making setup (Figure 1A) to ensure quick pad preparation and prevent molten agarose from solidifying prematurely.
   2. Allow the molten agarose to cool for 1 min at room temperature and dispense 50-100 µL spots on microscope slides using wide-bore pipette tips. Before the agarose cools down, place a microscope slide on the top to generate a spread pad of about 1.5-2 cm in diameter (Figure 1B-C). The width of the agarose pad is typically proportional to the length of imaging time. In other words, thicker pads withstand longer imaging periods.
   3. Use a hydrocarbon mixture as a sealant to ensure proper coverage of slides with thick agarose pads and to prevent cell death from solvent exposure at the coverslip edges. Mix equal parts (w/w) of petroleum jelly, lanolin, and paraffin in a 500 mL beaker. Heat ingredients on a hot plate at 120 °C for 5 min. As components melt, carefully swirl the molten solution to ensure appropriate mixing.
2. Sample setup
   1. For the examination of mitotic events, grow cells from starter cultures in either liquid EMM or PMG plus supplements to approximately mid-log phase (OD₅₉₅ of 0.4). Centrifuge 1 mL of the cell suspension at 1,375 x g for 1 min, remove the supernatant and resuspend the cell pellet in the minimal medium plus supplements to a final volume of 100 µL.
   2. To image meiotic events, grow cells from starter cultures in the minimal medium plus supplements to late-log phase (OD₅₉₅ of 0.7-1.0). Combine equal volumes of opposite mate-type cells (h^– and h⁺) in a 1 mL cell suspension. Centrifuge cells at 1,375 x g for 1 min and resuspend the pellet in liquid ME. Repeat this step thrice to ensure efficient nutrient removal.
   3. After the last ME wash, resuspend cells in 1 mL of ME and add it to a 50 mL flask containing 9 mL ME. Incubate for 12-16 h at 22-25 °C on minimal rotation speed (50-100 RPM). Abundant cell flocculation, which results in many round fission yeast clumps and indicates efficient mating.
   4. Take 1 mL sample of the mating culture and centrifuge at 1,375 x g for 1 min. Eliminate the supernatant but leave 250 µL in the tube to resuspend cells. Before placing cells on agarose pads, vortex vigorously for 5 s to disrupt clumps.
      NOTE: The remaining 250 µL ME used to resuspend cells contains mating factors that help cells enter meiosis.
   5. Place 20 µL 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 a lint-free paper towel for 2-3 s (Figure 1D). Set the slide pad-side up and gently place a glass coverslip, ensuring not to generate air bubbles (Figure 1F).
      NOTE: Eliminating excess medium with a paper towel is more time-efficient than gravity absorption, which is particularly critical in meiosis experiments.
   6. To create a cell monolayer, rotate the coverslip clockwise with the index finger for one (mitosis) or two (meiosis) full turns (Figure 1F). Ensure that the cell matter disperses across the agarose pad allowing for the better separation of single cells or asci. With the aid of a small wooden stick, dispense molten sealant along the edges of the coverslip to seal each agarose pad (Figure 1G).
   7. Once the agarose pad is sealed, place it on the microscope stage and let it equilibrate for 10-15 min at the appropriate imaging conditions (e.g., temperature) (Figure 1H) to allow air bubbles to dissipate and let any last-minute agarose shifts to occur. Begin imaging once the slide is efficiently equilibrated and do not remove it from the stage until data collection ends.
      NOTE: Control of temperature and humidity is determined by the microscope system employed and specific experimental requirements. If present, apply the temperature and humidity control to all imaging experiments. If absent, researchers must devise a way to prevent temperature fluctuations, especially during meiosis experiments.

4. Live-cell Imaging² and Processing

1. Use the 40x objective to find appropriate fields of view to the image. Switch to the 60x objective to begin data acquisition. Depending on the microscope system used, subsequent refocusing and sectioning on the sample at each time point will occur manually or through a microscope- or software-automated process.
   NOTE: The images presented in this protocol were collected using a deconvolution, fluorescence microscope equipped with Sedat, RFP and GFP filter sets, nano-motion stage, 60x NA 1.4 objective lens, and 12-bit CCD camera. Built-in mechanical shutters and motorized filter wheels allowed for the reduced excitation exposure and photobleaching of samples.
2. Use appropriate software to acquire and process images. To deconvolve images, employ manufacturer-provided optical transfer functions.
   NOTE: For details on the microscope equipment and software used in this protocol, refer to the Table of Materials.
3. To use a conventional fluorescence microscope to acquire live-cell images ensure that the microscope collects data digitally, has 40x or 60x objectives with numerical apertures (NA) greater than 1.2, and is able to perform optical sectioning.
   NOTE: Without these requirements, images will show reduced resolution, compromising the quality of microscopic measurements and qualitative observations.
4. Use free plug-in programs (e.g., Fiji) to minimize systematic blur errors arising from the contrast loss during the image acquisition^17,18,19,20 and to ensure proper quantitative analysis of the fluorescence intensity and of other temporal and dynamic events within the cell.
   NOTE: Other available commercial deconvolution programs can be found in the Table of Materials.
5. Choose the microscope filter sets that best match the fluorophores under observation. Ensure that the excitation and emission filters have the right band passes that allow for the specific detection of fluorescent proteins. For example, to detect CFP signal, an excitation and emission band pass of 430/25 and 470/30 is appropriate, but not for RFP fluorescence.
6. Use a minimal-effective-settings approach (MESA) when imaging fission yeast at multiple time points. In other words, avoid prolonged use of excitation wavelengths and employ the lowest excitation power and exposure time that generate acceptable, yet quantifiable and reproducible imaging data.
7. For prolonged imaging during mitosis or meiosis, limit the interval between acquisition time points to 5-10 min. Although 2% agarose pads can withstand 12 to 16 h imaging sessions, cell-shifting due to agarose evaporation is likely. Therefore, perform full mitosis or meiosis experiments in 4-6 h or 8-10 h windows, respectively.
8. Collect imaging data for 4-8 h consisting of at least 24-48 acquisition time points, respectively, one fluorescence protein, and a z-stack per time point and fluorescent channel comprised of 13 sections with 0.5-µm spacing using a 60x objective. This requires at least 0.5-1 GB of hard drive storage space. Thus, besides eliminating photobleaching and cell-shifting, create a workflow that considers computing capabilities^1,2,3.
   NOTE: These acquisition parameters are typically set and modified in the software that controls microscope functions and which collects and processes images.
9. To examine specific nuclear processes in closer detail, perform image acquisition every 5-10 s, as long as the emission does not decrease substantially after frequent exposure. Carry out a preliminary fluorescence intensity analysis over different time periods to determine the point after which the accuracy of the collected data is no longer reliable^1,3.
10. In the image-acquisition and image-processing software, use maximum intensity projection on the image z-stacks to observe all structures with high fluorescence density on a 2D plane regardless of their vertical location^1,2,3. This facilitates a rapid examination of nuclear processes occurring in different voxels. Collect a mid-focal bright-field image to generate a reference picture of the cells under observation.

5. ImageAnalysis^20,21

NOTE: Image analysis in this protocol is performed using Fiji. For other analysis programs and Fiji plug-ins see Table of Materials.

1. Upload a deconvolved image by selecting the Bio-Formats Importer feature under the Plugins menu. In the Import Options pop-up window, select Hyperstack, Default color mode, and check Autoscale before pressing the OK button.
2. 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 and prevents information loss.
3. Open an image that contains a scale bar generated during data processing by the microscope software. Determine the pixel length of the scale bar using the Straight Line tool to draw a parallel line of similar size and select Measure from the Analyze menu. Add a scale bar by setting the image pixel-to-µm ratio by selecting Set Scale from the Analyze menu.
   1. In the Set Scale pop-up window, enter the calculated Distance in pixels and Known distance in µm, set Pixel aspect ratio to 1.0, put micron as the Unit of length, and check the Global box before clicking the OK button.
4. Use the Set Measurements option under the Analyze menu to choose different parameters to quantify when selecting Measure. For the purposes of this protocol, select the following metrics: Area, Perimeter, Mean gray value, Median, Min & max gray value, Integrated density, Stack position, and Display label.
5. Depending on the precision of the collected data, enter more than 1 for the Decimal places option and check the Scientific notation box if necessary. After applying the Measure command, a Risultati pop-up window will show the values for each pertinent parameter. Save the results as a .csv file for future analysis in a statistics program.
   NOTE: It is not necessary to separate an image into its constituent fluorescent channels for determining the length, unless there is signal interference among the different colors in which case follow the step detailed below.
6. In the case of signal interference, select Color and Channels Tool from the Image menu and analyze each color separately. To measure the length of non-linear structures, right-click on the Line Tool and use the Segmented Line or Freehand Line option to trace the desired objects.
   1. For linear structures or to determine the distance between two points, use the Straight Line feature and select Measure from the Analyze menu. Values for length in µm will be automatically added to the parameters previously selected under the Set Measurements option.
7. For measuring changes in the signal size and fluorescence intensity, first create a region of interest (ROI) library, which increases quantification accuracy and facilitates speedy measurements across an image stack. Select Color and Channels Tool under the Image menu.
8. In the Channels pop-up window, check the color channel for which intensity or area will be measured. Choose the Adjust and Threshold features under the Image menu. Check the Dark background box, select the Default method (unless otherwise required by the collected data), and pick Red to overlay the signals of interest.
   NOTE: Measurement of protein stability, movement, and localization are closely dependent on the color threshold used for creating ROI libraries. It is important to choose the correct thresholding method for the type of fluorescent marker signal to be visualized^17,18.
9. 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. Repeat this process for each slice (time frame) in the image stack and store all ROIs by clicking on the Più button and selecting Save.
10. Open the zipped ROI folder to load the ROI manager and click on each ROI identifier on 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.
11. If cell shifting is not an issue and the focal plane remains the same for all slices of the stack, click on Multi measure in the ROI manager. In the pop-up window, select Measure all slices instead of One row per slice to iterate measurements across the image stack. Save results as a csv file and analyze measurements in a statistics program.
12. For multiple nuclear signals comprising the structure of interest, press the Shift key while selecting objects with the Wand tool to create a single ROI. Alternatively, create an ROI of constant size with the Oval tool to encompass all the signals of interest.
    NOTE: This oval approach may be necessary to measure and describe the signal behavior over an area where fluorescence signal changes in size and intensity over time. Signal intensity, in this case, is the mean signal density over a given area.
13. Qualitatively determine co-localization of two fluorescent signals by the change in color of the signal overlap region. However, as the co-localization zone narrows, it is more difficult to distinguish signal overlap. In this case, follow the steps detailed below.
14. Using the Channels Tool, as described in step 5.6, draw a straight line over each signal in question, and select the Plot Profile command under the Analyze menu.
    1. Repeat this step for all colors in question using the same drawn line.
    2. In the Plot of sample pop-up window that appears after each profiling step, press the List button to call up a Plot Values window, and save the measurements for each signal as a csv file.
    3. In a statistics program, create a graph that plots the Gray Value for each signal against its corresponding location (in µm) along the drawn line. Lack of overlap among signal profile plots generally indicates the lack of co-localization.
       NOTE: Use the Plot Profile tool on projected images to examine signal co-localization. This is only reliable if such overlap is also observed through the image z-stack.
15. To monitor the dynamic behavior of fluorescent proteins over time use the Multi Kymograph tool found under the Analyze menu. The resulting image shows the fluorescent signal movement through a series of condensed snapshots in each slice of the z-stack, which represents individual time points.
    1. Use the Channels Tool as described in step 5.6 to isolate the object of interest in the correct channel. Make sure the selected object or structure does not shift considerably through the z-stack. Place a straight line or rectangle over the object, select the Multi Kymograph tool from the Analyze menu, and enter the desired width of the line overlaying the object.
16. Create image panels that show nuclear dynamics over a specific time window using the Channels Tool as described in step 5.6 and by selecting the Stacks and Make Montage tools from the Image menu. In the Make Montage pop-up window, enter the number of Columns and Rows desired, set a Scale Factor of 1.0, choose the First and Last slices (time frames), and change Border width to at least 5 before pressing OK. It is possible to choose which images in the stack to show by changing Increments to suit the desired sequence.
    1. To generate a movie, upload the desired hyperstack, select Save as an avi file from the File menu, choose none for Compression, and enter a Frame rate of 2-8 fps. Select the Make Montage command while picking Composite in the Channels Tool to merge or separate all colors comprising the image.
       NOTE: Two avi files (Movie 1-2) are provided in this protocol. Use them in Fiji to try different features highlighted throughout step 5.
    2. To show a single, representative cell, use Transform and Rotate from the Image menu and enter degrees of rotation. Negative numbers rotate objects to the left, while positive values rotate objects to the right. Once the cell is in the proper orientation, draw a rectangle that encompasses the entire cell through the z-stack or during the time frames of interest and select Crop from the Image menu. Proceed as mentioned in step 5.16 and 5.16.1 to generate an image panel or movie.
17. Add a scale bar to the image panel or movie by selecting Tools and Scale bar from the Analyze menu. In the Scale Bar pop-up window, enter 5-15 for Width in microns for single cells or 100-250 for entire fields of view, choose white or black for Color, and pick an appropriate Location to place the scale bar.
    1. For movies, it is useful to show time progression during nuclear events. To show time change between frames, select Image, click on Stacks, use the Time Stamper tool, indicate the start frame in the time series, and select an appropriate location for the time display.