This work was funded by a DBT/Wellcome Trust (India Alliance) grant (IA/S/19/2/504632) to S.S. P.N. is a Research Fellow supported by a DBT/Wellcome Trust (India Alliance) grant (IA/S/19/2/504632). A.M. is supported by the Council of Scientific and Industrial Research (CSIR), Government of India, as a Senior Research Fellow (09/087(0873)/2017-EMR-I). The authors thank Paike Jayadeva Bhat for discussions.

NOTE: The protocol broadly involves the following steps: (1) patching the haploids in the mating efficiency grids on a YPD plate, (2) mixing the haploids in equal numbers after 24 h incubation and giving the mixed haploids a few hours to mate (7 h in this study), (3) plating the mixed cells on YPD for isolating single colonies after 7 h at 30 °C, and finally, (4) determining the number of diploids formed using the auxotrophic markers. These steps are discussed in detail below (also see Figure 2). 
1. Patching of haploids in the mating efficiency grids
<ol>
	<li>Revive the haploids a and α from freezer stocks by streaking on a YPD agar plate (2% agar, 2% dextrose, 1% peptone, 0.5% yeast extract), and allow them to grow for 48 h at 30 °C to obtain isolated single colonies.</li>
	<li>Inoculate single colonies from the YPD plate in 5 mL of YPD medium (2% dextrose, 1% peptone, 0.5% yeast extract), and incubate at 30 °C for 48 h with 250 rpm shaking. The cells are in the stationary phase of growth after this incubation period.</li>
	<li>Draw a mating efficiency grid on a fresh YPD plate. Draw the grid as a 1 cm x 1.5 cm rectangle divided into three boxes such that each has a dimension of 1 cm x 0.5 cm, as shown in Figure 2A.</li>
	<li>Patch 5 µL of the YPD culture of the two mating types on the leftmost and rightmost rectangles (Figure 2B). This volume corresponds to roughly 5 x 105 haploid cells laid out in each section. Incubate the plates for 24 h at 30 °C. 
	NOTE: The purpose of the grid is to make the experimental measures precise (such as the number of cells in the experiment). The grid size is small enough that it is experimentally tractable but large enough that it can be easily manipulated (like lifting cells from a grid) and not susceptible to drift or chance events.</li>
</ol>
​2. Mixing of haploids and mating
​NOTE: After 24 h (Figure 2C), an equal number of cells of the two haploid types are scraped off the two grids, mixed, and laid in the center rectangle (Figure 2D).
<ol>
	<li>In order to mix an equal number of cells, remove about 1/3 of the patch that was laid in the outer boxes using a sterile toothpick, and resuspend in 20 µL of water in a sterile 1.5 mL vial for each of the haploids.</li>
	<li>Dilute 5 µL of this suspension in 2 mL of water. Measure the OD of this diluted suspension using a spectrophotometer at 600 nm. Mix an equal number of cells from the two strains, based on the OD value and number of cells/mL in 1 OD for that particular strain. Calculate the volume required to be mixed, and aspirate it from the remaining 15 µL of the individual haploid suspension. 
	NOTE: The number of cells patched in the center rectangle is such that the cells form a monolayer. Considering the yeast cell to be a sphere with a radius of 2.58 µm33, a rectangular box of 1 cm x 0.5 cm would need approximately 1.7 x 106 cells to form a monolayer. Care should be taken to ensure that there is no physical touching between the cells patched for mating and the two haploid cell grids. Since equal numbers of each type of haploid cells have to be mixed, 8.5 x 105 cells are added from each strain. The cell number is calculated based on OD measurements, considering 1 OD at 600 nm to be approximately equivalent to 1 x 107 cells34. For example, if the OD600 of the a haploid suspension is 0.17 and that of the α haploid is 0.11, the number of cells in 5 μL of each haploid suspension can be calculated. To ensure 8.5 x 105 cells of each haploid type, 1.25 μL of the a haploid and 1.93 μL of the α haploid suspensions are mixed.</li>
	<li>Add the required volumes of both the haploids in a fresh and sterile 1.5 mL vial, and mix well using a pipette. The final volume of this suspension is generally around 6–8 µL. Patch this suspension in the center grid. Incubate the plate at 30 °C for 7 h, allowing the haploids sufficient time to mate (Figure 2E).</li>
</ol>
3. Plating of mixed cells on YPD agar
<ol>
	<li>After the incubation period of 7 h, scrape the cells from the center rectangle using a toothpick or a pipette tip, and dilute in 2 mL of sterile water. Then, spread the cell suspension on YPD agar to obtain single colonies. To determine the dilution factor necessary to obtain single colonies, measure the OD of the first tube into which the scraped cells are added. Specific dilution factors need to be determined for each cell type/strain being used. 
	NOTE: For example, an OD600 of 0.15 corresponds to 3 x 106 cells in a 2 mL suspension (considering 1 OD = 1 x 107 cells/mL). To obtain a few hundred colonies on the YPD plate, the cell suspension is serially diluted at 1:20 twice, and then 100 μL of the final dilution is used for spreading.</li>
	<li>After plating, incubate the YPD plates at 30 °C for 36–48 h until there are single colonies. Ensure that a few hundred individual colonies are obtained from each mating experiment for screening so as to ensure statistical significance can be detected in the data (Figure 2F). </li>
</ol>
4. Screening for diploids using auxotrophic markers
<ol>
	<li>Determine what fraction of the colonies obtained are diploid. To identify the diploid colonies on the plate, transfer the single colonies by streaking them individually onto a double drop-out plate (2% glucose, 0.66% nitrogen base, 0.05% double drop-out amino acid mixture, and 2% agar) lacking the amino acids that the strains are auxotrophic for, as shown in Figure 2G. Incubate the plates at 30 °C for 48 h. 
	NOTE: The colonies can also be transferred onto the double drop-out plate using replica plating. In this study, tryptophan and leucine (trp− leu−) drop-out medium was used when quantifying the mating efficiency of the SK1AM strains, and tryptophan and uracil (trp− ura−) drop-out medium was used for the ScAM strains. Only the diploid colonies grow on the double drop-out plate as they have both the auxotrophic markers: TRP1 and LEU2 genes in the SK1AM strains and TRP1 and URA3 genes in the ScAM strains.</li>
	<li>Additionally, streak or replica plate the colonies on single drop-out media (trp− or leu− or ura−) to quantify the frequency of each of the two kinds of haploid in the population.</li>
	<li>Calculate the mating efficiency, η, as follows: 
	<img alt="Mating efficiency formula: η=Number of haploids mated/Total number of haploids; biology equation." src="/files/ftp_upload/64596/64596eq01.jpg" style="margin: auto;" data-altupdatedat="1747929244000" data-alt="Equation 1">    Eq (1) 
	where the number of haploids mated is simply equal to twice the number of diploids identified on the double drop-out plate (since each diploid resulted from the mating of two haploids). The total number of haploids equals the sum of the number of haploids streaked plus twice the number of diploids streaked. 
	NOTE: For example, if only 60 colonies grow after streaking/replica plating 100 colonies on a double drop-out media, the mating efficiency can be quantified as 75% (as 60 x 2 haploids mated to form the 60 diploids, and 40 haploids did not mate).</li>
</ol>

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The authors declare that they have no competing interests in this work. The authors are happy to share the SK1-derived strains for all non-profit use.

The quantification of the mating efficiency in S. cerevisiae is essential for carrying out studies related to the genes involved in mating pathways or studying the influence of the external environment on mating behavior. In the past two decades, S. cerevisiae has also become a popular model to address questions related to speciation14,36,37,38. The presence of two mating types and the ease of genetic manipulation and maintenance in laboratory environments has made it an apt organism for studying the evolution of reproductive barriers in real time. The quantification of mating efficiency is essential as it gives a measure of the pre-zygotic reproductive barrier. Hence, a convenient protocol with high reproducibility is required.
Using the above protocol, the mating efficiency of two different strains that show completely different mating behavior was quantified. Hence, the protocol can be applied to any strain, as most lab yeast strains carry auxotrophies that can be used for selection39. However, there are a few important considerations while using this protocol. The time duration required for the initial YPD cultures to reach the stationary phase is strain dependent. A slow-growing strain may need more than 48 h incubation at 30 &#730;C. The volume of haploids to be mixed is calculated based on OD measurements in a spectrophotometer. The value reported in the literature is in the order of 107 cells/mL for 1 OD34; however, it is better to characterize this value for a particular strain using a plot of OD versus the number of cells. The volume required to ensure the mixing of equal numbers of cells can be calculated based on the OD value and the number of cells/mL in 1 OD, which are obtained from the characterization of the strain. One should ensure that the cells being patched in the center grid after mixing the haploids roughly form a monolayer that is not very thick. One also needs to make sure that the volume being mixed is approximately 6-8 &#181;L. A higher volume can lead to overlapping with the haploid patches that are on either side of the center grid.
This method can also be used to study pre-zygotic reproductive barriers between ecological isolates of yeast. However, for such cases, the HO endonuclease gene must first be removed from the organism&#39;s genome, as strains carrying HO endonuclease form diploids automatically due to its mating-type switching activity40. The haploids isolated from the sporulated diploid can be identified as a or &#945; by identifying the sequence present at the MAT locus using the specific primers mentioned earlier41. The HO marker can be replaced with two different antibiotic resistance genes as markers to distinguish between the two haploid mating types. This detail depends on the strain construction used in the particular study.
One of the limitations of this method is that the use of auxotrophic markers requires transferring the colonies from the YPD plate to the drop-out plates. Using fluorescence markers to distinguish between haploid and diploid cells can help reduce the experimental time. However, the use of fluorescence markers can affect the growth dynamics of the cells as there is an additional cost involved in producing the fluorescent proteins, which, thus, can alter a cell&#39;s metabolic and physiological state42. Alternatively, the auxotrophic markers to identify the two haploids can be replaced by antibiotic markers.
Some of the previous methods used for the characterization of mating efficiency report the use of an excess of one of the mating types at a ratio of 10:116,21,24. From experience, the use of a biased ratio of the two haploids results in mitotic growth of the haploid in excess. Therefore, the current method uses a 1:1 mix of the two haploids.
Since the auxotrophic markers in the strains that are used in this study are inserted close to the MAT locus, recombination during meiosis does not break the association between the auxotrophic marker and the mating type. As a result, the auxotrophy and the mating type of a particular haploid do not change even if the strains are allowed to undergo meiosis. This allows for answering questions related to the impact of gene flow as a variable on adaptation and speciation43. Thus, overall, this protocol presents a simple and robust method to quantify the mating efficiencies between different yeast strains.

Saccharomyces cerevisiae, commonly called budding yeast, is a unicellular eukaryote. It has two mating types, a and &#945;, and exhibits both asexual and sexual reproductive cycles. The a and &#945; mating types are haploids and can divide mitotically in the absence of the other mating type in the surrounding environment, which represents the asexual cycle of yeast. When the two mating types are in close proximity, they stop dividing mitotically and fuse to form a diploid cell. The diploid yeast can either divide mitotically when nutrients are present or undergo meiosis under the conditions of nitrogen starvation in the presence of a poor carbon source that is non-fermentable, such as acetate1. This results in the formation of spores, which remain dormant until there are favorable growth conditions. The life cycle is completed when these spores germinate and the two haploid types are released back to the haploid pool2,3 (Figure 1).

The mating of yeast cells includes several steps, such as agglutination, the formation of a mating projection or &#8220;shmoo&#8221;, followed by cell and nuclear fusion4,5. The two mating types a and &#945; produce a-factor and &#945;-factor, respectively, to initiate mating. These factors are polypeptide pheromones that bind to the receptors (Ste2 and Ste3) present on the cell surface of the opposite mating type5. The binding of the pheromones to the receptors initiates the pheromone response pathway, the mitogen-activated protein kinase (MAPK) signal transduction pathway6,7,8. This results in the arrest of the cell cycle in the G1 phase, leading to a metabolically active stationary phase9. The cells then stop dividing mitotically, and the proteins required for mating are synthesized. As the haploid cells cannot move toward each other, a mating projection or &#8220;shmoo&#8221; is directed toward the mating partner. When the cells come into contact, the cell wall is degraded, and the cytoplasmic contents fuse, resulting in mating to form a diploid cell10,11. The mating efficiency between haploids has been used as a measure of speciation in laboratory-evolved strains, as well as between extant species12.

Being a simple eukaryotic organism, yeast is the model of choice for a large number of research questions associated with complex eukaryotic organisms. One such question is associated with speciation and the evolution of reproductive barriers13,14. For sexually reproducing organisms, a species is defined by the biological species concept (BSC) proposed by Ernst Mayr15. According to this concept, two individuals of a population are said to belong to two different species if they cannot interbreed and are reproductively isolated. Breakdown of the sexual reproductive cycle (which involves the fusion of gametes to form a zygote, the development of the zygote into a progeny, and the attainment of sexual maturity in the progeny) leads to reproductive isolation. As shown in Figure 1, the life cycle of S. cerevisiae is comparable to the sexual reproductive cycle: a) the fusion of the two mating types a and &#945; is similar to the fusion of gametes in sexually reproducing organisms; b) the ability of the diploid to undergo mitotic division is equivalent to the zygote developing into progeny; and c) the diploid undergoing sporulation is comparable to the process of gametogenesis14.

Pre-zygotic isolation occurs when assortative mating is observed. Given an equal opportunity to mate with two genetically different a types, an &#945; type preferentially mates with one over the other or vice versa14. In the case of evolution experiments in which haploids have been evolved in different environments, the presence of a pre-mating barrier can be determined by performing a mating assay. A decrease in the mating efficiency when compared to the ancestor indicates the evolution of a pre-mating barrier. Post-zygotic isolation could arise due to the inability of the diploid to undergo effective mitotic division and/or sporulation to form haploid spores14. These can be quantified by measuring the growth rate of the diploids and calculating the sporulation efficiency, respectively. Hence, to study the evolution of reproductive barriers, robust methods for quantifying (a) the mating efficiency, (b) the mitotic growth of the diploid, and (c) the sporulation efficiency of the diploid are required. In this work, a robust method to quantify the mating efficiency of yeast strains is reported.

In laboratory experiments, one of the ways in which the occurrence of mating can be detected is by using auxotrophic markers that complement the nutritional requirements. When the two mating types are auxotrophic for two different amino acids, only the diploid cell formed by the fusion of the two mating types can grow on a medium deficient in both the amino acids. Thus, auxotrophic markers are useful for detecting mating both qualitatively and quantitatively. A qualitative test will suffice to identify the mating type of a strain after meiosis16. Quantitative tests are essential when one is interested in identifying a reduction in mating while studying the genes involved in the mating pathway17,18. In addition, with yeast being increasingly used in speciation studies, a convenient and reproducible mating assay is necessary, as the quantification of mating efficiency is a measure of the pre-zygotic barrier.

The mating efficiency between the two yeast mating types has been quantified previously16,19,20. Most of the previously used methods are similar in their design with a few variations16,21,22,23,24,25. Some of them use early log phase cultures, while a few others use mid-log phase cultures of haploid strains. There are variations in the ratios in which the two mating types are mixed. Almost all the protocols use a nitrocellulose membrane. Suspensions of both the mating types taken from previously grown cultures are mixed and filtered onto a nitrocellulose membrane placed on a YPD plate. In one of the variations of the protocol, the haploid suspension is directly patched on a YPD plate21. In experiments dealing with the genes involved in the pheromone production of the two mating types, the pheromones are externally added while making the suspensions of the two mating types24.

Following incubation for a few hours (typically around 5 h) after mixing the haploids, the cells are washed off from the membrane, diluted, and plated on selective media. In one of the earlier methods reported in 1973, the efficiency of zygote formation or mating was calculated by counting the number of budded cells, unbudded cells, and mating pairs under a microscope using a hemocytometer26. However, most methods reported later use auxotrophic markers to distinguish haploids and diploids. Mating efficiency is calculated as the percentage of diploid cells relative to the number of diploid and haploid cells in the cellular pool16,21,23.

However, despite a number of reports using yeast as a model organism to study speciation, there is no standardized protocol reported in the literature until now for calculating the efficiency of mating. Cells in the log phase may not be ideal for the quantification of mating efficiency. During mating, the cell cycle of the two haploids is arrested, and hence, the cells during mating are not dividing9. As the cell cycle is also known to be similarly arrested in cells in the stationary phase27, using such cells can make the protocol more reproducible. Stationary phase cells can be mixed and laid out on YPD plates (i.e., a nutritionally rich environment) for mating. The conventional procedures also require a nitrocellulose membrane and washing off the cells, making the process cumbersome and liable to handling errors. In addition, the protocols used to date quantify the mating efficiency in terms of one haploid. However, when measuring reproductive isolation, mating efficiency is quantified for a particular combination of haploids rather than a single haploid.

To address these issues, here, we report a robust method for the quantification of mating efficiency in yeast that is highly reproducible and easy to use. Moreover, this method and the yeast strains employed here can also be used in studies examining the effect of gene flow on the evolution of mating barriers.

Two different strains of S. cerevisiae were used in this study. One of the strains is derived from the SK1 background; this was modified in our laboratory by adding the auxotrophic markers near the MAT locus. The resulting genotypes of the haploids are provided in Table 128,29,30. In the SK1 strain, the a haploid had the TRP1 gene inserted near the MAT locus, and the &#945; haploid had the LEU2 gene inserted near the MAT locus. In the ScAM strain, the TRP1 and URA3 genes were inserted in the a and &#945; haploids, respectively. The location of insertion was in the ARS region of chromosome III (Chr III: 197378..197609). For the protocol reported here, auxotrophic markers anywhere on the genome would suffice. However, having the auxotrophic markers near the MAT locus means that these strains can also be used for studies examining the effect of gene flow on speciation31,32. The markers were added close to the MAT locus to prevent the reshuffling of the markers due to recombination. Hence, this protocol can be used to quantify mating efficiency in studies involving speciation and also to identify the alteration of mating efficiency when studying the proteins involved in the mating pathway.

<table><tbody><tr><td>Adenine</td><td>Sigma Life Science</td><td>A8626</td><td></td></tr><tr><td>Agar Powder regular grade for bacteriology</td><td>SRL</td><td>19661 (0140186)</td><td></td></tr><tr><td>Ammonium Sulphate, Hi-AR</td><td>HiMedia</td><td>GRM1273</td><td></td></tr><tr><td>D-(+)-glucose</td><td>Sigma Life Science</td><td>G8270</td><td></td></tr><tr><td>Glass Petri plates</td><td>HiMedia</td><td>PW008</td><td>&amp;nbsp;90 mm x 15 mm dimension</td></tr><tr><td>L-Arginine</td><td>Sigma Life Science</td><td>A8094</td><td></td></tr><tr><td>L-Aspartic acid</td><td>Sigma Life Science</td><td>A7219</td><td></td></tr><tr><td>L-Histidine monochloride monohydrate</td><td>Sigma Life Science</td><td>H5659</td><td></td></tr><tr><td>L-Isoleucine</td><td>Sigma Aldrich</td><td>I2752</td><td></td></tr><tr><td>L-Leucine</td><td>Sigma Life Science</td><td>L8912</td><td></td></tr><tr><td>L-Lysine</td><td>Aldrich</td><td>62840</td><td></td></tr><tr><td>L-Methionine</td><td>Sigma Life Science</td><td>M5308</td><td></td></tr><tr><td>L-Phenylalanine</td><td>Sigma Life Science</td><td>P5482</td><td></td></tr><tr><td>L-Threonine</td><td>Sigma Aldrich</td><td>T8625</td><td></td></tr><tr><td>L-Tyrosine</td><td>Sigma Life Science</td><td>T8566</td><td></td></tr><tr><td>L-Valine</td><td>Sigma Life Science</td><td>V0513</td><td></td></tr><tr><td>Mating efficiency grid</td><td></td><td></td><td>1 cm x 1.5 cm rectangular grid drawn on the Petri plate</td></tr><tr><td>Microcentrifuge tubes</td><td>Tarsons</td><td>500010</td><td></td></tr><tr><td>Peptone</td><td>HiMedia</td><td>RM001</td><td></td></tr><tr><td>Uracil</td><td>Sigma Life Science</td><td>U0750</td><td></td></tr><tr><td>Yeast Extract Powder</td><td>HiMedia</td><td>RM027</td><td></td></tr><tr><td>Yeast Nitrogen Base w/o Amino acids and Ammonium Sulphate</td><td>BD Difco</td><td>233520</td><td></td></tr></tbody></table>

determination of the mating efficiency of haploids in saccharomyces cerevisiae

Saccharomyces cerevisiae is a widely used model organism in genetics, evolution, and molecular biology. In recent years, it has also become a popular model organism to study problems related to speciation. The life cycle of yeast involves both asexual and sexual reproductive phases. The ease of performing evolution experiments and the short generation time of the organism allow for the study of the evolution of reproductive barriers. The efficiency with which the two mating types (a and &#945;) mate to form the a/&#945; diploid is referred to as the mating efficiency. Any decrease in the mating efficiency between haploids indicates a pre-zygotic barrier. Thus, to quantify the extent of reproductive isolation between two haploids, a robust method to quantify the mating efficiency is required. To this end, a simple and highly reproducible protocol is presented here. The protocol involves four main steps, which include patching the haploids on a YPD plate, mixing the haploids in equal numbers, diluting and plating for single colonies, and finally, calculating the efficiency based on the number of colonies on a drop-out plate. Auxotrophic markers are employed to clearly make the distinction between haploids and diploids.

Quantification of the mating efficiency of the two mating types 
The protocol described here was used to quantify the mating efficiency between two yeast strains-between SK1AMa and SK1AMα and between ScAMa and ScAMα (Figure 3A). In these experiments, the mating between the two haploids was repeated at least 12 times. In each of the repeats of the experiment, at least 100 colonies were streaked on double drop-out media. The robustness of the protocol enabled easy differentiation of the mating efficiency between the two strains, SK1AM and ScAM. While the SK1 strains mated with a very high efficiency, the ScAM strains mated with a relatively lower efficiency (Figure 3A). This is perhaps not surprising, as the ScAM strains originated from hybrid mating between the S. cerevisiae and S. carlsbergensis strains35.
In addition to distinguishing the mating efficiency between the strains, this method can also be used to quantify the differences in the mating efficiency of the strains across different environments. As shown in Figure 3B, the mating efficiency of the ScAM strains dropped significantly when the environment did not contain glucose as a primary carbon source. Mating is an energetically expensive process, and growth on alternate carbon sources qualitatively reduces the efficiency of mating.
Dynamics of mating efficiency 
The high repeatability of the method also allows for tracking the dynamics of mating efficiency. When allowed to mate for different time periods (Figure 3C), no mating was observed for the first 4-5 h; the first diploids appeared only after this duration. Presumably, this is the time needed for the mating process to be completed in the given environment. Thereafter, the mating efficiency increased rapidly. However, beyond 7 h, mating efficiency calculations are influenced by the fact that mitotic growth starts to happen on the plate. The identification and selection of early or late diploids in this way can allow for the design of experiments that aim to evolve rapid or delayed mating between haploids. The precise time at which the mating efficiency peaks is variable for each strain. For example, using growth kinetics, we experimentally determined that the ScAM strain exhibits a growth rate lower than that of other S. cerevisiae strains (like SK1), and this also probably influences the dynamics of mating.
<img alt="Yeast life cycle diagram; haploid-diploid stages, mitosis/meiosis processes, and mating transitions." class="xfigimg" src="/files/ftp_upload/64596/64596fig01.jpg" data-altupdatedat="1747929244000" data-alt="Figure 1"> 
Figure 1: Life cycle of Saccharomyces cerevisiae. S. cerevisiae has two mating types, a and α, and exhibits both asexual and sexual reproductive phases. The two mating types, in the absence of the other, divide mitotically. However, when they are present in the vicinity of each other, they stop dividing mitotically, and the cellular and nuclear contents fuse to form a diploid. The diploid cell can further divide mitotically. However, in the presence of unfavorable conditions (starvation), it undergoes meiosis to produce spores containing four haploids. These spores germinate under favorable conditions to release two haploids of each kind, thus completing the life cycle. The efficacy with which the two haploids a and α mate is referred to as the mating efficiency. Any decrease in this mating efficiency indicates a pre-zygotic barrier to mating. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64596/64596fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Yeast colony growth stages on YPD agar; microbiology experiment setup and result analysis." class="xfigimg" src="/files/ftp_upload/64596/64596fig02.jpg" data-altupdatedat="1747929244000" data-alt="Figure 2"> 
Figure 2: Steps involved in the mating efficiency assay. (A) A mating efficiency grid of 1 cm x 1.5 cm, which is further divided into boxes of 1 cm x 0.5 cm each, is drawn on a YPD plate. (B) Haploid cells are patched on the extreme boxes. (C) The growth of haploid cells after 24 h at 30 ˚C. (D) One-third of the haploid patches are removed using a toothpick, mixed in equal cell numbers (based on OD600), and patched in the center grid. (E) The growth of the cells patched in the center grid after 7 h at 30 ˚C. This contains the new diploids formed and the haploids that have not mated. (F) Isolated single colonies obtained on a YPD plate after the dilution and plating of the cells scraped from the center grid. This plate was incubated at 30 ˚C for 36-48 h. (G) To identify the number of diploids present on the YPD plate, the colonies are transferred to a double drop-out plate lacking tryptophan and leucine. Only diploids can grow on this plate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64596/64596fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Bar and line graphs of mating efficiency, showing variable conditions and time-dependent data." class="xfigimg" src="/files/ftp_upload/64596/64596fig03.jpg" data-altupdatedat="1747929244000" data-alt="Figure 3"> 
Figure 3: Mating efficiency of the yeast strains. (A) The mating efficiency of the two haploids a and α of the SK1 and ScAM strains on a YPD plate. (B) The mating efficiency of the two haploids a and α of the ScAM strain on YP plates containing 2% galactose and on glycerol/lactate plates.(C) The dynamics of mating in the ScAM strain in a glucose environment. The results are shown as mean ± SD from 12 independent repeats. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64596/64596fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Strains</td>
			<td>Genotype</td>
			<td>Comment</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>SK1AMa </td>
			<td>ars314::TRP1, MATa,  ho::LYS2, lys2, ura3, leu2::hisG, his3::hisG, trp1::hisG</td>
			<td>This strain has TRP1 gene and can hence grow in the absence of tryptophan in the media.</td>
			<td></td>
		</tr>
		<tr>
			<td>SK1AMα</td>
			<td>ars314::LEU2, MATalpha, ho::LYS2, lys2, ura3, leu2::hisG, his3::hisG, trp1::hisG</td>
			<td>This strain has LEU2 gene and can hence grow in the absence of leucine in the media.</td>
			<td></td>
		</tr>
		<tr>
			<td>ScAMa</td>
			<td>ars314::TRP1, MATa, ho, MEL1, ade1, ile, trp1-HIII ura3‐52</td>
			<td>This strain has TRP1 gene and can hence grow in the absence of tryptophan in the media.</td>
			<td>Derived from ScPJB644a in reference 28</td>
		</tr>
		<tr>
			<td>ScAMα</td>
			<td>ars314::URA3, MATalpha, ho, MEL1, ade1, ile, trp1-HIII, ura3‐52</td>
			<td>This strain has URA3 gene and can hence grow in the absence of uracil in the media.</td>
			<td>Derived from ScPJB644α in reference 28</td>
		</tr>
	</tbody>
</table>
Table 1: Genotypes of the S. cerevisiae strains used in this study.

Watch this Scientific Journal Video about Determination of the Mating Efficiency of Haploids in Saccharomyces cerevisiae at JoVE.com

Determination of the Mating Efficiency of Haploids in Saccharomyces cerevisiae

In this work, a robust method for the quantification of mating efficiency in the yeast Saccharomyces cerevisiae is described. This method is particularly useful for the quantification of pre-zygotic barriers in speciation studies.

Determination of the Mating Efficiency of Haploids in Saccharomyces cerevisiae

Saccharomyces cerevisiae is a widely used model organism in genetics, evolution, and molecular biology. In recent years, it has also become a popular ...

determination-mating-efficiency-haploids-saccharomyces

Nanyang Technological University

Research

JoVE Journal

Biology

2.4K Views.  Indian Institute of Technology Bombay. Dit protocol is een eenvoudige methode om de efficiëntie te kwantificeren waarmee verschillende stammen en gistsoorten met elkaar paren. De voorgestelde methode is eenvoudig, robuust en efficiënt. De methode kan worden uitgebreid tot elke giststam.De methode die in deze studie wordt beschreven, kan worden gebruikt om te begrijpen hoe soortvorming optreedt in gist. Bovendien kunnen de stammen die we beschrijven bijzonder nuttig zijn om experimenten met genstroom te ontwerpen. De krach...

Video: Determination of the Mating Efficiency of Haploids in Saccharomyces cerevisiae

Dissection of Saccharomyces Cerevisiae Asci

Yeast is a highly tractable model system that is used to study many different cellular processes. The common laboratory strain Saccharomyces cerevisiae exists in either a haploid or diploid state. The ability to combine alleles from two haploids and the ability to introduce modifications to the genome requires the production and dissection of asci. Asci production from haploid cells begins with the mating of two yeast haploid strains with compatible mating types to produce a diploid strain. This can be accomplished in a number of ways either on solid medium or in liquid. It is advantageous to select for the diploids in medium that selectively promotes their growth compared to either of the haploid strains. The diploids are then allowed to sporulate on nutrient-poor medium to form asci, a bundle of four haploid daughter cells resulting from meiotic reproduction of the diploid. A mixture of vegetative cells and asci is then treated with the enzyme zymolyase to digest away the membrane sac surrounding the ascospores of the asci. Using micromanipulation with a microneedle under a dissection microscope one can pick up individual asci and separate and relocate the four ascopores. Dissected asci are grown for several days and tested for the markers or alleles of interest by replica plating onto appropriate selective media.

Dissection of Saccharomyces Cerevisiae Asci

Micromanipulation of yeast cells is needed for meiotic genetic analysis or to select diploid zygotes. These micromanipulations are carried out using the microneedle of a dissection microscope. The microneedle is used to relocate cells and is controlled by a micromanipulator which are available with various degrees of automation.

Yeast is a highly tractable model system that is used to study many different cellular processes. The common laboratory strain Saccharomyces cerevisiae ...

Detection of Homologous Recombination Intermediates via Proximity Ligation and Quantitative PCR in Saccharomyces cerevisiae

DNA damage, including DNA double-stranded breaks and inter-strand cross-links, incurred during the S and G2 phases of the cell cycle can be repaired by homologous recombination (HR). In addition, HR represents an important mechanism of replication fork rescue following stalling or collapse. The regulation of the many reversible and irreversible steps of this complex pathway promotes its fidelity. The physical analysis of the recombination intermediates formed during HR enables the characterization of these controls by various nucleoprotein factors and their interactors. Though there are well-established methods to assay specific events and intermediates in the recombination pathway, the detection of D-loop formation and extension, two critical steps in this pathway, has proved challenging until recently. Here, efficient methods for detecting key events in the HR pathway, namely DNA double-stranded break formation, D-loop formation, D-loop extension, and the formation of products via break-induced replication (BIR) in Saccharomyces cerevisiae are described. These assays detect their relevant recombination intermediates and products with high sensitivity and are independent of cellular viability. The detection of D-loops, D-loop extension, and the BIR product is based on proximity ligation. Together, these assays allow for the study of the kinetics of HR at the population level to finely address the functions of HR proteins and regulators at significant steps in the pathway.

Detection of Homologous Recombination Intermediates via Proximity Ligation and Quantitative PCR in Saccharomyces cerevisiae

The D-loop capture (DLC) and D-loop extension (DLE) assays utilize the principle of proximity ligation together with quantitative PCR to quantify D-loop formation, D-loop extension, and product formation at the site of an inducible double-stranded break in Saccharomyces cerevisiae.

DNA damage, including DNA double-stranded breaks and inter-strand cross-links, incurred during the S and G2 phases of the cell cycle can be repaired by ...

Genetics

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

Assessing Differences in Sperm Competitive Ability in Drosophila

Competition among conspecific males for fertilizing the ova is one of the mechanisms of sexual selection, i.e. selection that operates on maximizing the number of successful mating events rather than on maximizing survival and viability 1. Sperm competition represents the competition between males after copulating with the same female 2, in which their sperm are coincidental in time and space. This phenomenon has been reported in multiple species of plants and animals 3. For example, wild-caught D. melanogaster females usually contain sperm from 2-3 males 4. The sperm are stored in specialized organs with limited storage capacity, which might lead to the direct competition of the sperm from different males 2,5.
Comparing sperm competitive ability of different males of interest (experimental male types) has been performed through controlled double-mating experiments in the laboratory 6,7. Briefly, a single female is exposed to two different males consecutively, one experimental male and one cross-mating reference male. The same mating scheme is then followed using other experimental male types thus facilitating the indirect comparison of the competitive ability of their sperm through a common reference. The fraction of individuals fathered by the experimental and reference males is identified using markers, which allows one to estimate sperm competitive ability using simple mathematical expressions 7,8. In addition, sperm competitive ability can be estimated in two different scenarios depending on whether the experimental male is second or first to mate (offense and defense assay, respectively) 9, which is assumed to be reflective of different competence attributes.
Here, we describe an approach that helps to interrogate the role of different genetic factors that putatively underlie the phenomenon of sperm competitive ability in D. melanogaster.

Assessing Differences in Sperm Competitive Ability in Drosophila

Differential sperm competitive ability among Drosophila males with distinct genotypes can be ascertained through double-mating experiments. Each of these experiments involves one of the males of interest and a reference male. Readily identifiable markers in the progeny allow inference of the fraction of individuals fathered by each male.

Competition among conspecific males for fertilizing the ova is one of the mechanisms of sexual selection, i.e. selection that operates on maximizing the ...

BEST: Barcode Enabled Sequencing of Tetrads

Tetrad analysis is a valuable tool for yeast genetics, but the laborious manual nature of the process has hindered its application on large scales. Barcode Enabled Sequencing of Tetrads (BEST)1 replaces the manual processes of isolating, disrupting and spacing tetrads. BEST isolates tetrads by virtue of a sporulation-specific GFP fusion protein that permits fluorescence-activated cell sorting of tetrads directly onto agar plates, where the ascus is enzymatically digested and the spores are disrupted and randomly arrayed by glass bead plating. The haploid colonies are then assigned sister spore relationships, i.e. information about which spores originated from the same tetrad, using molecular barcodes read during genotyping. By removing the bottleneck of manual dissection, hundreds or even thousands of tetrads can be isolated in minutes. Here we present a detailed description of the experimental procedures required to perform BEST in the yeast Saccharomyces cerevisiae, starting with a heterozygous diploid strain through the isolation of colonies derived from the haploid meiotic progeny.

Barcode Enabled Sequencing of Tetrads (BEST) replaces the manual processes of isolating, disrupting and spacing tetrads. BEST isolates tetrads by fluorescence-activated cell sorting onto agar plates, separates the spores by agitation with glass beads, and determines which randomly arrayed colonies were derived from the same original tetrad using molecular barcodes.

Tetrad analysis is a valuable tool for yeast genetics, but the laborious manual nature of the process has hindered its application on large scales. ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

Efficient Sporulation of Saccharomyces cerevisiae in a 96 Multiwell Format

During times of nutritional stress, Saccharomyces cerevisiae undergoes gametogenesis, known as sporulation. Diploid yeast cells that are starved for nitrogen and carbon will initiate the sporulation process. The process of sporulation includes meiosis followed by spore formation, where the haploid nuclei are packaged into environmentally resistant spores. We have developed methods for the efficient sporulation of budding yeast in 96 multiwell plates, to increase the throughput of screening yeast cells for sporulation phenotypes. These methods are compatible with screening with yeast containing plasmids requiring nutritional selection, when appropriate minimal media is used, or with screening yeast with genomic alterations, when a rich presporulation regimen is used. We find that for this method, aeration during sporulation is critical for spore formation, and have devised techniques to ensure sufficient aeration that are compatible with the 96 multiwell plate format. Although these methods do not achieve the typical ~80% level of sporulation that can be achieved in large-volume flask based experiments, these methods will reliably achieve about 50-60% level of sporulation in small-volume multiwell plates.

Efficient Sporulation of Saccharomyces cerevisiae in a 96 Multiwell Format

Here, sporulation of Saccharomyces cerevisiae is carried out in a 96 multiwell format.

During times of nutritional stress, Saccharomyces cerevisiae undergoes gametogenesis, known as sporulation. Diploid yeast cells that are starved for ...

High-throughput Screening for Protein-based Inheritance in S. cerevisiae

The encoding of biological information that is accessible to future generations is generally achieved via changes to the DNA sequence. Long-lived inheritance encoded in protein conformation (rather than sequence) has long been viewed as paradigm-shifting but rare. The best characterized examples of such epigenetic elements are prions, which possess a self-assembling behavior that can drive the heritable manifestation of new phenotypes. Many archetypal prions display a striking N/Q-rich sequence bias and assemble into an amyloid fold. These unusual features have informed most screening efforts to identify new prion proteins. However, at least three known prions (including the founding prion, PrPSc) do not harbor these biochemical characteristics. We therefore developed an alternative method to probe the scope of protein-based inheritance based on a property of mass action: the transient overexpression of prion proteins increases the frequency at which they acquire a self-templating conformation. This paper describes a method for analyzing the capacity of the yeast ORFeome to elicit protein-based inheritance. Using this strategy, we previously found that &gt;1% of yeast proteins could fuel the emergence of biological traits that were long-lived, stable, and arose more frequently than genetic mutation. This approach can be employed in high throughput across entire ORFeomes or as a targeted screening paradigm for specific genetic networks or environmental stimuli. Just as forward genetic screens define numerous developmental and signaling pathways, these techniques provide a methodology to investigate the influence of protein-based inheritance in biological processes.

High-throughput Screening for Protein-based Inheritance in S. cerevisiae

This protocol describes a high-throughput methodology to functionally screen for protein-based inheritance in S. cerevisiae.

The encoding of biological information that is accessible to future generations is generally achieved via changes to the DNA sequence. Long-lived ...

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

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

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

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.

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

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 ...

Determination of the Mating Efficiency of Haploids in Saccharomyces cerevisiae

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