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.
1. Preparation of necessary materials
2. Sporulation of yeast cells
3. Depletion of target protein from the nucleus using the anchor away technique
4. Time-lapse fluorescence microscopy
5. Analysis of chromatin segregation
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.
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.
The authors have nothing to disclose.
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 |
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 |