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.
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.
Time-lapse fluorescence microcopy is a valuable tool for studying the dynamics of meiotic chromosome segregation in budding yeast1,2. 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 NDT804,5. NDT80 expression is required for prophase I exit6,7. 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 meiosis9,10,11,12,13. In meiosis, the kinetochore is shed in prophase I, and Ctf19 is important for kinetochore re-assembly9,14. 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.
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1. Preparation of necessary materials
- 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.
- 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.
- 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.
- 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.
- Prepare drugs and reagents for microscopy, synchronization, and anchor away.
- 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.
- 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.
- 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.
- Generate yeast strains containing the NDT80-in system (PGAL1,10-NDT80/PGAL1,10-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.
- Prepare a chamber for time-lapse imaging
- 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.
- 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).
- 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.
- 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
- 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.
- 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.
- 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.
- 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.
- 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.
- Prophase I release system
- 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
- Add rapamycin to the cells to deplete the FRB-tagged protein from the nucleus at a particular stage.
- 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.
- 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.
- 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.
- 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 addition2,8.
4. Time-lapse fluorescence microscopy
- Make an agar pad that will be used to create a monolayer of cells for imaging (see step 4.2).
- 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.
- Place the cut microfuge tube cylinder on a clean glass slide with the top of the tube upside down sitting on the slide.
- 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.
- 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).
- Preparing yeast cells for imaging
- 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.
- Pipet 6 µL of the concentrated cells onto the coverslip in the middle of the chamber made in step 1.4.
- 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.
- Invert the mold such that the agar pad is facing down toward the chamber.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- Open the image acquisition software. Use course and fine adjustment knobs to focus the cells using DIC or brightfield.
- On the main menu of the image acquisition software, click on File > Acquire (Resolve 3D). Three windows will pop-up.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- 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.
- Click on Acquisition Controls > Acquisition. Click on Application > Define/Run Experiment. This will open a window labeled ND Acquisition.
- 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.
- 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%.
- 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.
- 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.
- 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.
- 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
- Open the Fiji software. Open one field of view at a time: for each field, open the DIC and mCherry channels.
- 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.
- Merge the DIC and mCherry channels in one image: click on Image > Color > Merge Channels.
- Follow a single cell through meiosis. After meiosis II completion, record the number of DNA masses.
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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.
The method described above was used to study the role of the kinetochore component Ctf19 in ensuring proper chromosome segregation in budding yeast meiosis. Anchor away yeast strains expressing Ctf19 tagged with FRB were synchronized in prophase I using the NDT80-in system (tor1-1; fpr1Δ; RPL13A-FKBP12; CTF19-FRB; GAL1-10 promoted NDT80; GAL4-ER). As a control, an NDT80-in strain with the anchor away strain background but without the FRB-tagged protein was used (tor1-1; fpr1Δ; RPL13A-FKBP12; GAL1-10 promoted NDT80; GAL4-ER). Cells were released from the prophase I arrest by the addition of β-estradiol. Rapamycin was added to deplete Ctf19-FRB from the nucleus either at release (before kinetochore reassembly), 45 min after release (after kinetochore reassembly but before meiosis I), or 3 h after release (just before meiosis II) (Figure 3A-F). Following the addition of rapamycin, time-lapse fluorescence microscopy was performed, and images were acquired every 10 min for the remaining duration of meiosis.
After imaging, the open-access software Fiji was used to open the images and perform analysis for figures showing time progression (Figure 3A-F). The number of DNA masses after meiosis II were counted in at least 100 cells that were undergoing meiosis per condition. In wildtype cells with the anchor away background, there are typically four DNA masses at the end of meiosis II, representing the four products of meiosis (Figure 3A). In a small fraction of the cells, only three masses are visible after meiosis II (Figure 3B,G). However, it is likely that three masses are the result of a cell that has four products of meiosis but two of these masses appear together after meiosis II.
When Ctf19-FRB is anchored away at the time of release of prophase I exit (t = 0 h), approximately 47% of cells display more than four DNA masses upon the completion of meiosis, suggesting a defect in the attachment of kinetochores and microtubules (Figure 3C,G). With anchoring away of Ctf19-FRB either after kinetochore assembly but before meiosis I (t = 45 min) or before meiosis II (t = 3 h), approximately 16% of cells display additional DNA masses. These results support the hypothesis that Ctf19 is important for kinetochore re-assembly at prophase I release but is less important in chromosome segregation once the kinetochore has been reassembled.
The results obtained here show that time-lapse microscopy combined with cell cycle synchronization and conditional depletion of a target protein allow for the study of a protein in a specific meiotic stage. Although Ctf19 is not essential for mitosis, the meiotic kinetochore is extensively reorganized, and Ctf19 has a crucial role in kinetochore re-assembly after prophase I exit9,11,12. The results in Figure 3 show that when Ctf19 is depleted prior to kinetochore re-assembly, a large fraction of cells display additional DNA masses after meiosis II. When Ctf19 is depleted from the nucleus after kinetochore reassembly but before either meiosis I or meiosis II, there are fewer cells that display additional DNA masses. These results suggest that the most important role for Ctf19 is at the end of prophase I. This difference in chromosome segregation fidelity with depletion of Ctf19 at various stages highlights the importance of using stage-specific conditional alleles to study the function of proteins specifically in meiosis I or meiosis II. A meiotic null mutant would have displayed a severe defect and would not have provided information about the role of Ctf19 at each stage.
Additionally, although CTF19 is not an essential gene, there is a chromosome transmission fidelity (ctf) phenotype during vegetative growth of CTF19 mutants19. Use of a null allele or mutant CTF19 allele could create a population of aneuploid cells that may not undergo meiosis properly. Use of the anchor away technique and NDT80-in synchronization system circumvent this problem by precisely depleting Ctf19 from the nucleus just prior to meiosis I or meiosis II, reducing the concern that the analyzed cells were aneuploid.
Figure 1: Overview of the experiment. Cartoon of yeast cell showing a single pair of homologous chromosomes. NDT80-in cells enter meiosis and arrest at prophase I. After addition of β-estradiol, cells enter the meiotic divisions synchronously. Addition of rapamycin determines when the target protein (marked with a star) is anchored away. In this experiment, the target protein Ctf19 was anchored away at three different time points by adding rapamycin (RAP) at the same time as prophase I (RAP addition at t = 0 min), after prophase I release (RAP addition at t = 45 min), and after meiosis I chromosome segregation (RAP addition at t = 3 h). The thick black arrows indicate cell cycle stage of rapamycin addition and, therefore, target protein depletion. Abbreviations: RAP = rapamycin. Please click here to view a larger version of this figure.
Figure 2: Chamber set-up for time-lapse imaging experiments. (A) Reusable chamber (right) cut from a pipette tip holder insert (left). The pipette tip holder insert is cut with a red-hot blade to remove a 4 square x 4 square portion of the insert. The dashed lines indicate a sample section of the tip holder that can be cut to make a chamber. The remaining plastic dividers within the 4 square x 4 square cut-out are trimmed away and a hollow chamber remains (right). (B) To create the final chamber, silicone sealant is added to edges on one side of the chamber, and then placed on top of a cover slip. Please click here to view a larger version of this figure.
Figure 3: Time at which Ctf19 is anchored away affects chromatin segregation. (A-F) Representative time lapse images of cells expressing Htb2-mCherry during meiosis. Images were taken every 10 min, immediately following the addition of β-estradiol. Scale bars= 5 µm. Numbers in the upper-right corner of each frame label the time that has elapsed between each frame, and time 0 indicates the frame in which chromatin segregates in meiosis I. Only cells that completed the meiotic divisions were counted. (A) Wildtype cell (no protein tagged with FRB) in which β-estradiol was added 12 h after KAc transfer. Cell contains four DNA masses after completion of meiosis II. (B) Ctf19-FRB cell in which rapamycin was added at t = 0 (simultaneously with β-estradiol addition). This cell shows three DNA masses after meiosis II. (C) Ctf19-FRB cell in which rapamycin was added at t = 0 (simultaneously with β-estradiol addition). This cell shows five DNA masses after meiosis II. (D) Ctf19-FRB cell in which rapamycin was added at t = 0 (simultaneously with β-estradiol addition). This cell dies after completing meiosis II. (E) Ctf19-FRB cell in which rapamycin was added 45 min after β-estradiol addition. This cell shows five DNA masses after meiosis II. (F) Ctf19-FRB cell in which rapamycin was added 45 min after β-estradiol addition. After meiosis II, six DNA masses are present. (G) Quantification of the number of DNA masses in at least 100 cells for each condition (two movies analyzed for each condition). t = under each bar indicates the time at which rapamycin was added relative to β-estradiol addition. Abbreviations: MII = meiosis II. Please click here to view a larger version of this figure.
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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 release4,8. Although individual cells will vary slightly in the amount of time spent in each of the subsequent meiotic stages, most cells will maintain a high degree of synchrony throughout the meiotic divisions.
The anchor away technique is an adaptable tool for mitotic and meiotic studies. By altering the anchor away target protein and the time of rapamycin addition, this method can be adapted to study various proteins during any stage of budding yeast meiosis. Common methods for making conditional mutants to study budding yeast mitosis are not always suitable for meiotic studies. For example, use of a repressible promoter often requires a change in carbon source to alter the expression of the target protein, which may disrupt meiosis20. Temperature-sensitive mutants rely on a change in temperature to reduce or abolish the activity of the target protein. Many of these conditional mutants cannot be used in meiotic studies because shifting nutrient conditions or temperature can disrupt meiosis21,22,23. Furthermore, deletions or mutants may limit studies of meiosis because expressing a mutant protein early in meiosis might block or adversely affect later stages. The anchor away technique can circumvent these challenges because it does not rely on changes in nutrition or temperature8. Moreover, depletion of the protein can be temporally regulated such that the time of rapamycin addition determines when the protein will be anchored away.
One limitation of the anchor away technique is that it may not be suitable for non-nuclear proteins because it relies on the Rpl13A ribosomal subunit as an anchor to transport the target protein out of the nucleus. However, there are other applications of the anchor away technique by changing the protein-protein interactions that may also be useful during meiosis, such as anchoring proteins to complexes or tethering them to membranes8. Previous studies show that nuclear depletion of target proteins occurs within 30 min of rapamycin addition2,8. However, it is possible that some proteins need a longer or shorter period to be shuttled out of the nucleus. To ensure that the target protein is being shuttled out of the nucleus, the target protein can be tagged with FRB fused to GFP (FRB-GFP) to monitor the length of time between rapamycin addition and nuclear depletion. Another limitation of the anchor away system is that some proteins are sensitive to the FRB and FRB-GFP tags at the C-terminus. As such, N-terminal tagging of the target protein with FRB is an alternative strategy for anchoring away a target protein.
To monitor chromatin segregation, budding yeast cells expressed Htb2-mCherry. In this strain, the Htb2 protein is tagged at its endogenous locus, which ensures that Htb2 is expressed normally. Other fluorescently tagged proteins could also be used to monitor different aspects of meiosis. When constructing a strain for time-lapse imaging, it is important to consider that some proteins are sensitive to C-terminal fluorescent tags. Growth and sporulation assays of the strain harboring the fluorescently tagged protein ensures that the tag is not altering the function of the protein. One challenge of time-lapse fluorescence microscopy is exposing cells to enough fluorescence so that the fluorescent proteins are detected while avoiding cell death or slowing of meiosis from excess exposure. An important troubleshooting step is to find the appropriate microscope and camera settings to achieve this balance. The required exposure time will vary slightly between proteins, so multiple iterations of an experiment should be performed to determine the appropriate exposure time. When using the chamber method described here for time-lapse imaging experiments, an especially sensitive step is achieving a monolayer of cells to be imaged. Without a monolayer, cells accumulate on top of one another, which presents a challenge for proper imaging and data analysis. Using an agar pad, which helps cells adhere to the ConA and spreads cells around the chamber, is an inexpensive and efficient way to obtain monolayers. Moving the agar pad to spread cells around the chamber such that a monolayer is achieved can be challenging. Ensuring that the agar pad is making very small movements during the moving process can assist in creating a monolayer. Making a reusable chamber allows for the inexpensive generation of time-lapse imaging data. With proper cleaning and disinfecting, the plastic chambers described here can be reused indefinitely. It is best practice that chambers are removed from the coverslip immediately following time-lapse imaging and stored in 95% ethanol until the next use.
Time-lapse imaging provides information about the cell cycle that can be missed from imaging fixed cells1,2. For example, when monitoring the duration of the meiotic stages, variations between single cells can be more accurately examined when imaging live cells compared to populations of fixed cells. Because yeast proteins can be easily tagged with fluorescent proteins, coupling fluorescence time-lapse imaging with the NDT80-in system and the anchor away technique can be adapted for additional studies, such as monitoring individual chromosome segregation and the timing of specific meiotic stages. When monitoring cell cycle progression following prophase I release in NDT80-in cells, it is crucial to have a protein tagged with a fluorescent molecule that will mark the stages of meiosis. Individual chromosomes can be monitored throughout meiosis by integrating repeats of the lac operon (LacO) near the centromere of a chromosome in cells expressing LacI-GFP. LacI-GFP binds tightly to LacO and is visualized as a bright focus, which can be followed throughout the meiotic divisions by time-lapse microscopy24,25. Various proteins with fluorescent tags have been used to monitor the duration of meiotic stages. These include Tub1, the α-tubulin subunit that incorporates into microtubules; Zip1, a component of the synaptonemal complex; and Spc42, a spindle pole body component1,2,26,27.
In conclusion, this method incorporates meiotic cell cycle synchronization, conditional depletion of a target protein, and time-lapse fluorescence microscopy to study meiotic chromosome segregation. By imaging budding yeast cells during meiosis and using conditional mutants that affect only the intended stages of meiosis, this method can provide accurate data about the meiotic cell cycle.
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The authors declare no competing financial interests.
We thank the Light Microscopy Imaging Center at Indiana University. This work was supported by a grant from the National Institutes of Health (GM105755).
|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|
|Concanavialin A||Mllipore Sigma||C2010||Make as 1mg/mL in 1X PBS|
|CoolSNAP HQ2 CCD camera||Photometrics||Used in Section 4.3|
|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|
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