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 &#176;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).&#160;
1. Patching of haploids in the mating efficiency grids
<ol>
	<li>Revive the haploids a and&#160;&#945; 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 &#176;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 &#176;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 &#181;L 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 &#176;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>
&#8203;2. Mixing of haploids and mating
&#8203;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 &#181;L of water in a sterile 1.5 mL vial for each of the haploids.</li>
	<li>Dilute 5 &#181;L 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 &#181;L 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 &#181;m33, 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 &#945; haploid is 0.11, the number of cells in 5 &#956;L of each haploid suspension can be calculated. To ensure 8.5 x 105 cells of each haploid type, 1.25 &#956;L of the a haploid and 1.93 &#956;L of the &#945; 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&#8211;8 &#181;L. Patch this suspension in the center grid. Incubate the plate at 30 &#176;C for 7 h, allowing the haploids sufficient time to mate&#160;(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 &#956;L of the final dilution is used for spreading.</li>
	<li>After plating, incubate the YPD plates at 30 &#176;C for 36&#8211;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).&#160;</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 &#176;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&#8722; leu&#8722;) drop-out medium was used when quantifying the mating efficiency of the SK1AM strains, and tryptophan and uracil (trp&#8722; ura&#8722;) 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&#8722; or leu&#8722; or ura&#8722;) to quantify the frequency of each of the two kinds of haploid in the population.</li>
	<li>Calculate the mating efficiency, &#951;, as follows: 
	<img alt="Equation 1" src="/files/ftp_upload/64596/64596eq01.jpg" style="margin: auto;" />&#160; &#160; 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 type...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>&#160;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 SK1AM&#945; and between ScAMa and ScAM&#945; (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...

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 ...

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2.3K Views.  Indian Institute of Technology Bombay. 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.

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 ...

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground using a cryolysis procedure in a ball mill grinder and the resulting grindate brought into solution by urea solubilization. This procedure allows for the lysis of the cells in a metabolically inactive state, preserving phosphorylation and preventing reorientation of the phosphoproteome during cell lysis.  Following reduction, alkylation, trypsin digestion of the proteins, the samples are desalted on C18 columns and the sample complexity reduced by fractionation using hydrophilic interaction chromatography (HILIC). HILIC columns preferentially retain hydrophilic molecules which is well suited for phosphoproteomics.  Phosphorylated peptides tend to elute later in the chromatographic profile than the non phosphorylated counterparts. After fractionation, phosphopeptides are enriched using immobilized metal chromatography, which relies on charge-based affinities for phosphopeptide enrichment. At the end of this procedure the samples are ready to be quantitatively analyzed by mass spectrometry.

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

Description of a quantitative phosphorylation procedure using cryolysis, urea solubilziation, HILIC fractionation and IMAC enrichment of phosphorylated peptides.

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground ...

Microarray Analysis for Saccharomyces cerevisiae

In this protocol, gene expression in yeast (Saccharomyces cerevisiae) is changed after exposure to oxidative stress induced by the addition of hydrogen peroxide (H2O2), an oxidizing agent. In the experiment, yeast is grown for 48 hours in 1/2X YPD broth containing 3X glucose. The culture is split into a control and treated group. The experiment culture is treated with 0.5 mM H2O2 in Hanks Buffered Saline (HBSS) for 1 hour. The control culture is treated with HBSS only. Total RNA is extracted from both cultures and is converted to a biotin-labeled cRNA product through a multistep process. The final synthesis product is taken back to the UVM Microarray Core Facility and hybridized to the Affymetrix yeast GeneChips. The resulting gene expression data are uploaded into bioinformatics data analysis software.

Microarray Analysis for Saccharomyces cerevisiae

In this protocol, gene expression in yeast (Saccharomyces cerevisiae) is changed after exposure to oxidative stress induced by the addition of hydrogen peroxide (H2O2), an oxidizing agent.

In this protocol, gene expression in yeast (Saccharomyces cerevisiae) is changed after exposure to oxidative stress induced by the addition of hydrogen ...

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

Expression and Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein in Saccharomyces cerevisiae

The cystic fibrosis transmembrane conductance regulator (CFTR) is a chloride channel, that when mutated, can give rise to cystic fibrosis in humans.There is therefore considerable interest in this protein, but efforts to study its structure and activity have been hampered by the difficulty of expressing and purifying sufficient amounts of the protein1-3. Like many 'difficult' eukaryotic membrane proteins, expression in a fast-growing organism is desirable, but challenging, and in the yeast S. cerevisiae, so far low amounts were obtained and rapid degradation of the recombinant protein was observed 4-9. Proteins involved in the processing of recombinant CFTR in yeast have been described6-9 .In this report we describe a methodology for expression of CFTR in yeast and its purification in significant amounts. The protocol describes how the earlier proteolysis problems can be overcome and how expression levels of CFTR can be greatly improved by modifying the cell growth conditions and by controlling the induction conditions, in particular the time period prior to cell harvesting. The reagants associated with this protocol (murine CFTR-expressing yeast cells or yeast plasmids) will be distributed via the US Cystic Fibrosis Foundation, which has sponsored the research. An article describing the design and synthesis of the CFTR construct employed in this report will be published separately (Urbatsch, I.; Thibodeau, P. et al., unpublished). In this article we will explain our method beginning with the transformation of the yeast cells with the CFTR construct - containing yeast plasmid (Fig. 1). The construct has a green fluorescent protein (GFP) sequence fused to CFTR at its C-terminus and follows the system developed by Drew et al. (2008)10. The GFP allows the expression and purification of CFTR to be followed relatively easily. The JoVE visualized protocol finishes after the preparation of microsomes from the yeast cells, although we include some suggestions for purification of the protein from the microsomes. Readers may wish to add their own modifications to the microsome purification procedure, dependent on the final experiments to be carried out with the protein and the local equipment available to them. The yeast-expressed CFTR protein can be partially purified using metal ion affinity chromatography, using an intrinsic polyhistidine purification tag. Subsequent size-exclusion chromatography yields a protein that appears to be &gt;90% pure, as judged by SDS-PAGE and Coomassie-staining of the gel.

Expression and Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein in Saccharomyces cerevisiae

Attempts to express the cystic fibrosis transmembrane conductance regulator (CFTR) in Saccharomyces cerevisiae have, until now, yielded relatively low amounts of protein. This protocol and the associated reagents distributed via the Cystic Fibrosis Foundation should allow the preparation of milligram amounts of this 'difficult' eukaryotic membrane protein.

The  cystic fibrosis transmembrane conductance regulator (CFTR) is a chloride  channel, that when mutated, can give rise to cystic fibrosis in  ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein Expressed in Saccharomyces cerevisiae

Defects in the cystic fibrosis transmembrane conductance regulator (CFTR) protein cause cystic fibrosis (CF), an autosomal recessive disease that currently limits the average life expectancy of sufferers to &#60;40 years of age. The development of novel drug molecules to restore the activity of CFTR is an important goal in the treatment CF, and the isolation of functionally active CFTR is a useful step towards achieving this goal.
We describe two methods for the purification of CFTR from a eukaryotic heterologous expression system, S. cerevisiae. Like prokaryotic systems, S. cerevisiae can be rapidly grown in the lab at low cost, but can also traffic and posttranslationally modify large membrane proteins. The selection of detergents for solubilization and purification is a critical step in the purification of any membrane protein. Having screened for the solubility of CFTR in several detergents, we have chosen two contrasting detergents for use in the purification that allow the final CFTR preparation to be tailored to the subsequently planned experiments.
In this method, we provide comparison of the purification of CFTR in dodecyl-&#946;-D-maltoside (DDM) and 1-tetradecanoyl-sn-glycero-3-phospho-(1&#39;-rac-glycerol) (LPG-14). Protein purified in DDM by this method shows ATPase activity in functional assays. Protein purified in LPG-14 shows high purity and yield, can be employed to study post-translational modifications, and can be used for structural methods such as small-angle X-ray scattering and electron microscopy. However it displays significantly lower ATPase activity.

Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein Expressed in Saccharomyces cerevisiae

Heterologous expression and purification of the cystic fibrosis transmembrane conductance regulator (CFTR) are significant challenges and limiting factors in the development of drug therapies for cystic fibrosis. This protocol describes two methods for the isolation of milligram quantities of CFTR suitable for functional and structural studies.&#160;

Defects in the cystic fibrosis transmembrane conductance regulator (CFTR) protein cause cystic fibrosis (CF), an autosomal recessive disease that ...

EPR Monitored Redox Titration of the Cofactors of Saccharomyces cerevisiae Nar1

Electron Paramagnetic Resonance (EPR) monitored redox titrations are a powerful method to determine the midpoint potential of cofactors in proteins and to identify and quantify the cofactors in their detectable redox state.
The technique is complementary to direct electrochemistry (voltammetry) approaches, as it does not offer information on electron transfer rates, but does establish the identity and redox state of the cofactors in the protein under study. The technique is widely applicable to any protein containing an electron paramagnetic resonance (EPR) detectable cofactor.
A typical titration requires 2 ml protein with a cofactor concentration in the range of 1-100 &#181;M. The protein is titrated with a chemical reductant (sodium dithionite) or oxidant (potassium ferricyanide) in order to poise the sample at a certain potential. A platinum wire and a Ag/AgCl reference electrode are connected to a voltmeter to measure the potential of the protein solution. A set of 13 different redox mediators is used to equilibrate between the redox cofactors of the protein and the electrodes. Samples are drawn at different potentials and the Electron Paramagnetic Resonance spectra, characteristic for the different redox cofactors in the protein, are measured. The plot of the signal intensity versus the sample potential is analyzed using the Nernst equation in order to determine the midpoint potential of the cofactor.

EPR Monitored Redox Titration of the Cofactors of Saccharomyces cerevisiae Nar1

The goal of this protocol is to use electron paramagnetic resonance (EPR) monitored redox titrations to identify different cofactors of Saccharomyces cerevisiae Nar1. Redox titrations offer a very robust way to obtain midpoint potentials of different redox active cofactors in enzymes and proteins.

Electron Paramagnetic Resonance (EPR) monitored redox titrations are a powerful method to determine the midpoint potential of cofactors in proteins and to ...

Measurement of mRNA Decay Rates in Saccharomyces cerevisiae Using rpb1-1 Strains

mRNA steady state levels vary depending on environmental conditions. Regulation of the steady state accumulation levels of an mRNA ensures that the correct amount of protein is synthesized for the cell&#8217;s specific growth conditions. One approach for measuring mRNA decay rates is inhibiting transcription and subsequently monitoring the disappearance of the already present mRNA. The rate of mRNA decay can then be quantified, and an accurate half-life can be determined utilizing several techniques. In S. cerevisiae, protocols that measure mRNA half-lives have been developed and include inhibiting transcription of mRNA using strains that harbor a temperature sensitive allele of RNA polymerase II, rpb1-1. Other techniques for measuring mRNA half-lives include inhibiting transcription with transcriptional inhibitors such as thiolutin or 1,10-phenanthroline, or alternatively, by utilizing mRNAs that are under the control of a regulatable promoter such as the galactose inducible promoter and the TET-off system. Here, we describe measurement of S. cerevisiae mRNA decay rates using the temperature sensitive allele of RNA polymerase II. This technique can be used to measure mRNA decay rates of individual mRNAs or genome-wide.

Measurement of mRNA Decay Rates in Saccharomyces cerevisiae Using rpb1-1 Strains

The steady state level of specific mRNAs is determined by the rate of synthesis and decay of the mRNA. Genome-wide mRNA degradation rates or the decay rates of specific mRNAs can be measured by determining mRNA half-lives. This protocol focuses on measurement of mRNA decay rates in Saccharomyces cerevisiae.

mRNA steady state levels vary depending on environmental conditions. Regulation of the steady state accumulation levels of an mRNA ensures that the ...