Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Summary

Time-lapse microscopy is a valuable tool for studying meiosis in budding yeast. This protocol describes a method that combines cell-cycle synchronization, time-lapse microscopy, and conditional depletion of a target protein to demonstrate how to study the function of a specific protein during meiotic chromosome segregation.

Abstract

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often not seen by imaging fixed cells. Budding yeast has proved to be an important model organism to study meiotic chromosome segregation because many meiotic genes are highly conserved. Time-lapse microscopy of meiosis in budding yeast allows the monitoring of different meiotic mutants to show how the mutation disrupts meiotic processes. However, many proteins function at multiple points in meiosis. The use of loss-of-function or meiotic null mutants can therefore disrupt an early process, blocking or disturbing the later process and making it difficult to determine the phenotypes associated with each individual role. To circumvent this challenge, this protocol describes how the proteins can be conditionally depleted from the nucleus at specific stages of meiosis while monitoring meiotic events using time-lapse microscopy. Specifically, this protocol describes how the cells are synchronized in prophase I, how the anchor away technique is used to deplete proteins from the nucleus at specific meiotic stages, and how time-lapse imaging is used to monitor meiotic chromosome segregation. As an example of the usefulness of the technique, the kinetochore protein Ctf19 was depleted from the nucleus at different time points during meiosis, and the number of chromatin masses was analyzed at the end of meiosis II. Overall, this protocol can be adapted to deplete different nuclear proteins from the nucleus while monitoring the meiotic divisions.

Introduction

Time-lapse fluorescence microcopy is a valuable tool for studying the dynamics of meiotic chromosome segregation in budding yeast^1,2. Budding yeast cells can be induced to undergo meiosis through starvation of key nutrients³. During meiosis, cells undergo one round of chromosome segregation followed by two divisions to create four meiotic products that are packaged into spores (Figure 1). Individual cells can be visualized throughout each stage of meiosis, which generates spatial and temporal data that can be easily missed by fixed-cell imaging. This protocol shows how combining time-lapse fluorescence microscopy with two previously established methods, the inducible NDT80 system (NDT80-in) and the anchor away technique, can be used to study the function of specific proteins in distinct meiotic stages.

The NDT80-in system is a powerful tool for meiotic cell cycle synchronization that relies on the inducible expression of the middle meiosis transcription factor NDT80^4,5. NDT80 expression is required for prophase I exit^6,7. With the NDT80-in system, NDT80 is under the control of the GAL1-10 promoter in cells expressing the Gal4 transcription factor fused to an estrogen receptor (Gal4-ER)^4,5. Because Gal4-ER only enters the nucleus when bound to β-estradiol, NDT80-in cells arrest in prophase I in the absence of β-estradiol, which allows the synchronization of cells in prophase I (Figure 1). β-estradiol addition promotes the translocation of the Gal4-ER transcription factor into the nucleus, where it binds GAL1-10 to drive expression of NDT80, leading to synchronous entry into the meiotic divisions. Although time-lapse microscopy can be performed without synchronization, the advantage of using synchronization is the ability to add an inhibitor or a drug while cells are at a specific stage of meiosis.

The anchor away technique is an inducible system by which a protein can be depleted from the nucleus with the addition of rapamycin⁸. This technique is ideal for studying nuclear proteins during cell division in budding yeast because yeast cells undergo closed mitosis and meiosis, in which the nuclear envelope does not break down. Furthermore, this technique is very useful for proteins that have multiple functions throughout meiosis. Unlike for deletions, mutant alleles, or meiotic null alleles, the removal of a target protein from the nucleus at a specific stage does not compromise target protein activity at earlier stages, allowing for a more accurate interpretation of results. The anchor away system utilizes the shuttling of ribosomal subunits between the nucleus and cytoplasm that occurs upon ribosomal maturation⁸. To deplete the target protein from the nucleus, the target protein is tagged with the FKBP12-rapamycin-binding domain (FRB) in a strain in which the ribosomal subunit Rpl13A is tagged with FKBP12. Without rapamycin, FRB and FKBP12 do not interact, and the FRB-tagged protein remains in the nucleus. Upon rapamycin addition, the rapamycin forms a stable complex with FKBP12 and FRB, and the complex is shuttled out of the nucleus due to the interaction with Rpl13A (Figure 1). To prevent cell death upon rapamycin addition, cells harbor the tor1-1 mutation of the TOR1 gene. Additionally, these cells contain fpr1Δ, a null allele of the S. cerevisiae FKBP12 protein, which prevents endogenous Fpr1 from out-competing Rpl13A-FKBP12 for FRB and rapamycin binding. The anchor away background mutations, tor1-1 and fpr1Δ, do not affect meiotic timings or chromosome segregation².

To demonstrate the usefulness of this technique, the kinetochore protein Ctf19 was depleted at different timepoints throughout meiosis. Ctf19 is a component of the kinetochore that is dispensable in mitosis but required for proper chromosome segregation in meiosis^{9,10,11,12,13}. In meiosis, the kinetochore is shed in prophase I, and Ctf19 is important for kinetochore re-assembly^9,14. For this protocol, cells with the NDT80-in system were synchronized, and the anchor away technique was used to deplete the target protein Ctf19 from the nucleus before and after the release from prophase I, and after meiosis I chromosome segregation (Figure 1). This protocol can be adapted to deplete other proteins of interest at any stage of meiosis and mitosis.

Protocol

1. Preparation of necessary materials

1. Prepare reagents for the growth and sporulation of yeast cells.
   NOTE: If budding yeast strains are ade2- and trp1-, supplement all media in steps 1.1.1-1.1.3 with a final concentration of 0.01% adenine and 0.01% tryptophan from 1% stocks. If sterilizing media by autoclave, add these amino acids only after the reagents have been autoclaved and allowed to cool to room temperature.
   1. For vegetative growth, prepare 2x synthetic complete + dextrose medium (2XSC) by dissolving 6.7 g of yeast nitrogen base without amino acids, 2 g of complete amino acid mix, and 20 g of dextrose in 500 mL of water. Sterilize the mixture by autoclaving for 20 min at 121 °C or filtering through a 0.2 µm filter.
   2. For the first step of sporulation, prepare 2x synthetic complete + acetate medium (2XSCA) by dissolving 6.7 g of yeast nitrogen base without amino acids, 2 g of complete amino acid mix, and 20 g of potassium acetate in 500 mL of water. Sterilize the mixture by autoclaving for 20 min at 121 °C or filtering through a 0.2 µm filter.
   3. For the final step of sporulation, prepare 1% potassium acetate (1% KAc) by dissolving 5 g of KAc in 500 mL of water. Sterilize the mixture by autoclaving for 20 min at 121 °C or filtering through a 0.2 µm filter.
2. Prepare drugs and reagents for microscopy, synchronization, and anchor away.
   1. For adhering cells to the coverslip during time-lapse imaging, make 1 mg/mL of concanavalin A (ConA) in 1x PBS. Filter sterilize using a 0.2 µm filter and store small (~10 μL) aliquots at -20 °C.
   2. To use the NDT80-in system, make 2 mL of 1 mM β-estradiol dissolved in ethanol. Filter sterilize using a 0.2 µm filter and store aliquots (~500 μL) at -20 °C.
   3. For the anchor away system, make 1 mg/mL of rapamycin dissolved in DMSO. Filter sterilize using a 0.2 µm filter and store small (~10 μL) aliquots at -20 °C.
      NOTE: Prepare small aliquots of rapamycin to avoid multiple freeze-thaw cycles of the drug.
3. Generate yeast strains containing the NDT80-in system (P_GAL1,10-NDT80/P_GAL1,10-NDT80; GAL4-ER/ GAL4-ER) and a target protein tagged with FRB in the anchor away genetic background (tor1-1/tor1-1; fpr1Δ/ fpr1Δ; Rpl13A-FKBP12/Rpl13A-FKBP12)^4,8. Ctf19-FRB was used for this study. Additionally, one copy of histone protein Htb2 was tagged with mCherry to allow for monitoring of meiotic progression and chromatin segregation.
4. Prepare a chamber for time-lapse imaging
   1. Start preparing the chamber 6 h-24 h prior to imaging (Figure 2). Cut an 18 mm x 18 mm chamber from the pipette tip box insert using a heated scalpel or other sharp blade.
   2. Use a plastic pipette tip to spread a thin layer of silicone sealant around the bottom edges of the chamber that will adhere to the coverslip. Add enough sealant so that the edges of the chamber are completely covered (see Figure 2).
   3. Adhere the chamber to a 24 mm x 50 mm coverslip by gently placing the chamber, sealant-side-down, onto the coverslip. Make sure that there are no gaps in the sealant so that the chamber does not leak.
   4. Spread 8-10 µL of ConA onto the coverslip. Dispense ConA onto the middle of the coverslip and use a pipette tip to spread the ConA into a thin layer such that it covers most of the coverslip that is surrounded by the chamber.
      ​NOTE: Plastic chambers can be reused indefinitely. After time-lapse imaging is completed, remove the chamber from the coverslip using a razor, clean the silicone sealant from the chamber, and keep the chambers submerged in 95% ethanol for future use.

2. Sporulation of yeast cells

1. Perform the following steps for starvation of yeast cells to induce the meiotic program.
   NOTE: These steps are for the sporulation of the W303 strain of yeast. Other strains may require different protocols¹⁵. Sporulation efficiency is highly variable between lab strains, with W303 exhibiting a sporulation efficiency of ~60%^16,17,18.
   1. Take a single colony of the appropriate diploid yeast strain from a plate and inoculate 2 mL of 2XSC and let grow at 30 °C on a roller drum for 12-24 h to saturation.
   2. Dilute the saturated culture into 2XSCA by adding 80 µL of the culture from step 2.1.1 to 2 mL of 2XSCA. Let it grow at 30 °C on a roller drum for 12-16 h. Do not leave in 2XSCA for longer than 16 h because cells can become sick or auto-fluorescent.
   3. Perform two washes by spinning down the culture at 800 x g at room temperature (25 °C) for 1 min, discarding the liquid, and resuspending the pellet in 2 mL of sterile distilled water.
   4. After the second wash, remove the liquid and resuspend the pellet in 2 mL of 1% KAc and let it grow at 25 °C on a roller drum for 8-12 h. Cells that have entered meiosis will arrest in pachytene of prophase I.
2. Prophase I release system
   1. Add β-estradiol directly to the sporulation culture to a final concentration of 1 μM and vortex the culture tube quickly. The β-estradiol will release cells from prophase I.

3. Depletion of target protein from the nucleus using the anchor away technique

1. Add rapamycin to the cells to deplete the FRB-tagged protein from the nucleus at a particular stage.
   1. To deplete proteins from the nucleus at prophase I exit, add rapamycin to a final concentration of 1 μg/mL to the sporulation culture tube at the same time as β-estradiol addition.
   2. To deplete proteins from the nucleus at a particular stage of meiosis, monitor the cell cycle stage starting at 60 min after β-estradiol addition. Every 20 min, pipette 5 µL of culture onto a 24 mm x 40 mm coverslip and cover the cells with an 18 mm x 18 mm coverslip. Keep cultures spinning on the roller drum at 25 °C between time points.
   3. Image the cells using the A594/mCherry filter and the 60x objective of a fluorescence microscope. Set the percent transmittance to 2% and the exposure time to 250 ms. The presence of one, two, or four DNA masses will indicate the meiotic stage at which the cells have progressed. Add rapamycin to a final concentration of 1 μg/mL at the meiotic stage of interest.
2. Keep cells spinning on the roller drum at 25 °C until they are ready for imaging. Nuclear depletion of a target protein occurs 30-45 min after rapamycin addition^2,8.

4. Time-lapse fluorescence microscopy

1. Make an agar pad that will be used to create a monolayer of cells for imaging (see step 4.2).
   1. Cut off and discard the cap and bottom 1/3 of a 1.5 mL microfuge tube to create a cylinder. The cylinder will serve as a mold for the agar pad. Make two cylinders to have an extra in case the agar does not polymerize properly in the first one.
   2. Place the cut microfuge tube cylinder on a clean glass slide with the top of the tube upside down sitting on the slide.
   3. Make 6 mL of a 5% agar solution (use 1% KAc as solvent) in a 50 mL beaker and microwave it until the agar is fully dissolved.
      NOTE: The agar will easily boil over in the microwave; watch the agar and start and stop the microwave when the agar starts to boil and swirl the beaker. Start and stop the microwave several times to fully dissolve the agar. The agar solution is made in excess of the amount needed to ensure that the agar dissolves fully.
   4. Cut off the tip of the pipet to make a larger opening and pipet ~500 µL of melted agar into each microfuge tube cylinder. Let it sit at room temperature until the agar solidifies (~10-12 min).
2. Preparing yeast cells for imaging
   1. Spin down 200 µL of the sporulation culture from step 3.2 at 800 x g for 2 min in 1 mL microcentrifuge tubes. Remove and discard 180 µL of the supernatant. Resuspend the pellet in the remaining supernatant by swirling and flicking the tube.
   2. Pipet 6 µL of the concentrated cells onto the coverslip in the middle of the chamber made in step 1.4.
   3. Hold the cylinder with the agar pad made in step 4.1 and carefully slide it off the glass slide. Make sure the agar is completely flat at the bottom and using the bottom of a pipette tip, apply a slight amount of pressure to the microfuge mold such that the agar pad is pushed out slightly above the boundary of the tube.
   4. Invert the mold such that the agar pad is facing down toward the chamber.
   5. Using forceps, gently place the agar pad (still in the microfuge tube mold) on top of the cells. Use a pipette tip to gently slide the agar pad around the chamber 10-20 times to create a monolayer of cells on the coverslip.
   6. Leave the agar pad in the chamber for 12-15 min. This step will allow the cells to adhere to the ConA on the coverslip.
   7. Transfer 2 mL of sporulation culture from step 3.2 to two microcentrifuge tubes and spin at 15,700 x g for 2 min. Transfer the supernatant into clean microfuge tubes and spin again at 15,700 x g for 2 min. Transfer the supernatant to clean microcentrifuge tubes to be used in the next step.
      NOTE: It is important to use the pre-conditioned KAc supernatant with the β-estradiol and rapamycin to ensure efficient sporulation and continued depletion of the protein of interest.
   8. After the agar pad has sat in the chamber for 12-15 min, float and remove the agar pad before imaging. To do so, add 2 mL of the supernatant from step 4.2.7 dropwise to the chamber. Once the liquid has reached the top of the chamber, the agar pad will most likely float.
      NOTE: If the agar pad does not float automatically, wait 1-2 min. If the agar pad still does not float, remove it gently with forceps. Ideally, the agar pad would float on its own because removing it prior to floating could lead to removal of the cells.
   9. After the agar pad has floated, remove it gently with forceps and discard it. Place a 24 mm x 50 mm coverslip on the top of the chamber to prevent evaporation during imaging.
3. Setting up a movie on the microscope
   NOTE: The instructions below are for an inverted microscope fitted with a slide holder that accommodates the 24 mm x 50 mm coverslip (See Table of Materials for microscope, camera, and software details). A 60x oil-immersion objective is used for image acquisition. This protocol may need to be altered when using other microscopes or other slide holders. The exact steps for executing step 4.3.2 through step 4.3.16 will vary depending on the microscope and imaging software used. See section 4.4 for instructions for a different microscope.
   1. Fit the coverslip inside the slide holder. Adhere the molding clay to the side of the coverslip to keep it secure in the slide holder.
   2. Open the image acquisition software. Use course and fine adjustment knobs to focus the cells using DIC or brightfield.
   3. On the main menu of the image acquisition software, click on File > Acquire (Resolve 3D). Three windows will pop-up.
   4. In the window named Resolve 3D, click on the Erlenmeyer Flask icon. This will open a window titled Design/Run Experiment, which contains the controls for setting up an experiment to set up a time-lapse movie.
   5. Under the Design tab, navigate to the tab labeled Sectioning. Select the box next to Z Sectioning. Set the z stacks as follows: Optical section spacing = 1 µm, Number of optical sections = 5, Sample thickness = 5.0 µm.
   6. Under the Channels tab, click on the + icon such that one channel option appears. Select the appropriate channel. For this experiment, we select A594, which is used to image mCherry. Select the box next to Reference Image and set the Z position to the middle of the sample from the drop-down menu.
   7. Select a value from the dropdown menu adjacent to %T and Exp. to set percent transmittance and exposure time, respectively. For this experiment, 2% transmittance and 250 ms exposure time are used in the A594 channel, and 10% transmittance and 500 ms exposure time are used for brightfield.
      NOTE: The exposure time and percent transmittance will vary with different microscopes. To avoid overexposure, use the lowest percent transmittance and shortest exposure time possible that allows for proper visualization of the protein of interest.
   8. Under the Time-lapse tab, select the box next to Time-lapse. In the table that appears, enter 10 under the min column in the time-lapse row and 10 under the hours column in the total time row. This will run a time course taking images every 10 min for 10 h.
   9. Select the box next to Maintain Focus with Ultimate Focus to prevent stage drift during the movie. Under the Points tab, select the box next to Visit Point List.
   10. In the main menu, click on View > Point List. A window called point list will pop up. Move the stage to an area of the chamber that shows a monolayer of cells. Click on Point Mark in the point list window.
   11. Move the stage to select 25-30 points without any overlap to avoid overexposing the cells. Each field will be imaged during each time course.
   12. In the point list window, select Calibrate All to set ultimate focus for each point. Under the Design tab in the Desgin/Run Experiment window, enter the range of point values (obtained from the point list window) in the box next to Visit Point List. Under the Run tab, save the file to the appropriate destination on the computer. On the Design/Run Experiment window, select the Play button (green triangle icon) to start the movie.
4. Optional method: Setting up a movie on a microscope
   NOTE: These instructions are for an inverted microscope fitted with a slide holder that can accommodate a 24 mm x 50 mm coverslip. See Table of Materials for specifications about the microscope, camera, and imaging software used. A 60x oil-immersion objective is used for image acquisition.
   1. Fit the coverslip inside the slide holder. Open the image acquisition software. Use course and fine adjustment knobs to focus cells using DIC or brightfield. Right click in the open window of the software. A drop-down menu will appear.
   2. Click on Acquisition Controls > Acquisition. Click on Application > Define/Run Experiment. This will open a window labeled ND Acquisition.
   3. In the window that opens, check the box next to XY. Move the stage to the desired location and click the box under Point Name to select that location as a point. Repeat this until 25-30 points are selected without any overlap to avoid overexposing the cells. Each field will be imaged during each time course.
   4. Set percent transmittance for each fluorescent channel. Because the strains produce Htb2-mCherry, the mCherry filter is used here. Under the Acquisition window that opened in step 4.3.3, navigate to the 555 nm button. Set the percent transmittance to 5% by adjusting the sliding scale under the 555 nm button until it reads 5%.
   5. Set the exposure time for each fluorescent channel. Under the Acquisition window, select 200 ms from the Exposure drop-down menu.
      NOTE: The exposure time and percent transmittance will vary with different microscopes. To avoid overexposure, use the lowest percent transmittance and shortest exposure time possible that allows for proper visualization of the protein of interest.
   6. Click on the box next to Z in the ND Acquisition window. Set five z-stacks of the yeast cells that are 1.2 µM apart.
   7. Click on the box next to λ. Select the appropriate channels to be used for the experiment. For DIC, ensure that Home is selected from the drop-down menu under Z pos. For mCherry, ensure that All is selected from the drop-down menu under Z pos. This ensures that, only a single section (in the middle of the z-stack) is taken for DIC.
   8. Select the box next to Time in the ND Acquisition window. Select the box under Phase. In the drop-down menu under Interval, select 10 min. Under Duration, select 10 hour (s). This will run time course such that images are taken every 10 min for 10 h.

5. Analysis of chromatin segregation

1. Open the Fiji software. Open one field of view at a time: for each field, open the DIC and mCherry channels.
2. Take the maximum intensity projection of the mCherry channel to obtain a single image: click on Image > Stacks > Z Project, select Max Intensity from the dropdown menu.
3. Merge the DIC and mCherry channels in one image: click on Image > Color > Merge Channels.
4. Follow a single cell through meiosis. After meiosis II completion, record the number of DNA masses.

Results

To monitor chromatin segregation, histone protein Htb2 was tagged with mCherry. In prophase I, the chromatin appears as a single Htb2 mass. After homologous chromosomes segregate in the first meiotic division, the chromatin appears as two distinct masses (Figure 3A). After the sister chromatids segregate, the chromatin appears as four masses. If some chromosomes fail to attach to spindle microtubules, additional masses can be seen after meiosis I or meiosis II.

Th...

Discussion

This protocol combines the NDT80-in system to synchronize cells, the anchor away technique to deplete proteins from the nucleus, and fluorescence time-lapse microscopy to image budding yeast cells during meiosis. The NDT80-in system is a method for meiotic cell cycle synchronization that utilizes a prophase I arrest and release^4,8. Although individual cells will vary slightly in the amount of time spent in each of the subsequent meiotic stages, ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
β-estradiol	Millipore Sigma	E8875	Make 1mM stocks in 95% EtOH
0.22 uM Threaded Bottle-top Filter	Millipore Sigma	S2GPT02RE
100% EtOH	Fisher Scientific	22-032-601
10X PBS	Fisher Scientific	BP399500	Dilute 1:10 to use as solvent for ConA
24 mm x 50 mm coverslip No. 1.5	VWR North American	48393241
25 mm x 75 mm microscope slides	VWR North American	48300-026
Adenine hemisulfate salt	Millipore Sigma	A9126	To supplement SC, SCA, and 1% Kac
Bacto Agar	BD	214030
Concanavialin A	Mllipore Sigma	C2010	Make as 1mg/mL in 1X PBS
CoolSNAP HQ2 CCD camera	Photometrics		Used in Section 4.3
D-glucose	Fisher Scientific	D16-10
Difco Yeast Nitrogen Base w/o Amino Acids	BD	291920
Dimethyl sulfoxide (DMSO)	Millipore Sigma	D5879
Eclipse Ti2 inverted-objective micrscope	Nikon		Used in Section 4.4
Fiji	NIH		Download from https://fiji.sc/
GE Personal DeltaVision Microscope	Applied Precision		Used in Section 4.3
L-Tryptophan	Millipore Sigma	T0254	To supplement SC, SCA, and 1% Kac
Modeling Clay	Crayola	2302880000	To secure coverslip in slide holder
NIS-Elements AR 5.30.04 Imaging Software	Nikon		Used in Section 4.4
ORCA-Fustion BT Camera	Hamamatsu	C15440-20UP	Used in Section 4.4
Plastic pipette tip holder	Dot Scientific	LTS1000-HR	Cut a 4 square x 4 square section of the rack portion of this product.
Pottassium Acetate	Fisher Scientific	BP264
Rapamycin	Fisher Scientific	BP29631	Make 1mg/mL stocks in DMSO
Silicone Sealant	Aqueon	100165001	Also known as aquarium glue.
SoftWorx7.0.0  Imaging Software	Applied Precision		Used in Section 4.3
Synthetic Complete Mixture (Kaiser)	Formedium	DSCK2500
Type N immersion oil	Nikon	MXA22166