The authors wish to acknowledge Agence Nationale de la Recherche (ANR) and Association pour la Recherche sur le Cancer (ARC) for the PhD fellowships to J.d.D.B.T. and the Agence Nationale pour la Recherche (ANR) for financial support (grant ANR-18-CE12-0018-01). Cytometry and microscopy were performed at the Montpellier MRI BioCampus imaging facility.

1. S. cerevisiae cell culture
NOTE: The yeast strains used are listed in Table 1
NOTE: The S-phase duration can be monitored in different ways. Depending on the question to be addressed, the cells can be grown asynchronously or synchronously following G1 arrest.
<ol>
	<li>From asynchronously growing S. cerevisiae cells 
	NOTE: This method allows for the determination of the percentage of cells in the S phase in an asynchronously growing cell population. By determining the doubling time, the duration of the S phase (and the other phases) can be extrapolated.
	<ol>
		<li>Inoculate S. cerevisiae cells in 10 mL of synthetic complete (SC) medium at a low cell concentration (5 &#215; 104 cells/mL) for an overnight culture at 30 &#730;C with orbital agitation at 130 rpm. 
		NOTE: The concentration is measured with a cell counter. To efficiently calculate the doubling times, it is recommended to inoculate the culture from an overnight culture that is still in the exponential phase (i.e., ideally below 2 &#215; 107 cells/mL). Growing cells in rich medium (YPD) is not recommended, as EdU detection is not efficient.</li>
		<li>The following day, dilute the cells in 20 mL of fresh SC medium at the final concentration of 5 &#215; 105 cells/mL.</li>
		<li>Culture the cells at 30 &#730;C in a shaking water bath with horizontal shaking at 120 rpm.</li>
		<li>Measure the cell concentration every hour until it reaches 1 &#215; 107 cells/mL. 
		NOTE: This step allows for making a graphical representation of the cell concentration increase over time. The formula used to calculate the doubling times is explained in the legend of Table 2.</li>
		<li>Proceed in parallel to step 2 for EdU labeling when the cell concentration is about 2 &#215; 106-5 &#215; 106 cells/mL.</li>
	</ol>
	</li>
	<li>From G1-synchronized S. cerevisiae cells 
	NOTE: This method allows for the determination of when the S phase starts and finishes by means of flow cytometry and/or microscopy analyses.
	<ol>
		<li>Inoculate S. cerevisiae cells in 10 mL of SC medium at a low cell concentration (5 &#215; 104 cells/mL) for an overnight culture at 30 &#730;C with orbital agitation at 130 rpm. 
		NOTE: See the notes after step 1.1.1.</li>
		<li>The following day, dilute the cells in 20 mL of fresh SC medium at a final concentration of 2 &#215; 106-3 &#215; 106 cells/mL.</li>
		<li>Add 40 &#181;L of 1 mg/mL &#945;-factor diluted in water.</li>
		<li>Culture the cells at 30 &#730;C with orbital agitation at 130 rpm for 1 h.</li>
		<li>Add again 40 &#181;L of 1 mg/mL &#945;-factor diluted in water.</li>
		<li>Culture the cells at 30 &#730;C with orbital agitation at 130 rpm for 1 h.</li>
		<li>Visualize the cells under a light microscope to monitor G1 arrest. Proceed if more than 90% of the cells display a shmoo and the others are rounded, unbudded cells. 
		NOTE: Depending on the background used, it is recommended to sonicate the cells before shmoo visualization. For the W303 background, sonicate 2x, for 2 s each time, at an amplitude of 40-50.</li>
		<li>Centrifuge for 3 min at 1,500 &#215; g. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the cells in 20 mL of SC medium.</li>
		<li>Repeat steps 1.2.8-1.2.9 once. 
		NOTE: The &#945;-factor is washed away with these steps, and the cells are released in the cell cycle. Alternatively, the &#945;-factor may be washed away by filtering the yeast cells with a 1.2 &#181;m nitrocellulose filter using a funnel set on a side-arm flask connected to a vacuum pump.</li>
		<li>Collect 1 mL of cells 2x every 5 min and proceed to step 2 for EdU labeling. 
		NOTE: Pulse-label the cells from only one of the two tubes with EdU. The non-pulse-labeled cells are used to distinguish EdU-positive from EdU-negative cells on a bivariate propidium iodide (PI)-EdU graph.</li>
		<li>Add 400 &#181;L of 1 mg/mL &#945;-factor diluted in water 30 min after the release. 
		&#8203;NOTE: This high dose of &#945;-factor is required to arrest the cells in the G1 phase of the next cell cycle and to prevent the cells from re-entering the subsequent S phase.</li>
	</ol>
	</li>
</ol>
2. EdU labeling
<ol>
	<li>Transfer 1 mL of cell culture to a 2.0 mL microfuge tube containing 1 &#181;L of 10 mM EdU. Mix well by inversion. 
	NOTE: To discriminate the EdU-positive cells from the EdU-negative cells on a PI-EdU bivariate FACS, transfer another 1 mL of cell culture to a 2.0 mL microfuge tube containing 1 &#181;L of DMSO.</li>
	<li>Incubate for 3-5 min at 30 &#730;C under agitation in a shaking water bath. 
	NOTE: Three minutes are sufficient for EdU detection with a microscope; 5 min are required for optimal EdU detection on a flow cytometer.</li>
	<li>Stop the reaction with the addition of 100 &#181;L of 100% ethanol.
	<ol>
		<li>Stop the reaction with the addition of 100 &#181;L of 20% paraformaldehyde if cell size measurement is required. 
		NOTE: If the nuclear architecture of the mitotic cells is to be kept intact for further analyses following the Click reaction, we recommend fixing cells with 2% paraformaldehyde at room temperature (RT) rather than putting the cells on ice, since the latter causes microtubule depolymerization.</li>
		<li>Leave the cells for 20 min at RT under mild agitation on a variable speed rocker at 20 tilts/min to fix the cells before the addition of 100 &#181;L of 100% ethanol.</li>
	</ol>
	</li>
</ol>
3. Cell fixation and permeabilization
<ol>
	<li>Pellet the cells for 2 min at 10,000 &#215; g in a microfuge. Remove the supernatant using a vacuum pipette.</li>
	<li>Resuspend the cell pellet in 500 &#181;L of 70% ethanol. Mix well by vortexing.</li>
	<li>Leave for &#8805;1 h at RT at 20 tilts/min on a variable speed rocker to permeabilize the cells. 
	&#8203;NOTE: Cells grown in SC medium do not pellet well as they tend to stick to the microfuge walls. The addition of ethanol improves pelleting and reduces cell loss. The cells may be stored at 4 &#730;C overnight or for longer periods at &#8722;20 &#730;C.</li>
	<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
	<li>Wash the cells 2x with 500 &#181;L of 10% ethanol in PBS. 
	&#8203;NOTE: The washes are crucial to remove unincorporated EdU from the cells.</li>
</ol>
4. Click-it reaction
<ol>
	<li>For cytometry analysis
	<ol>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microfuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the pellet in 200 &#181;L of PBS containing 0.1 mg/mL RNase A and 0.2 mg/mL proteinase K.</li>
		<li>Incubate for 1-2 h at 50 &#176;C with occasional shaking (or overnight at 37 &#176;C).</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Wash the cells with 500 &#181;L of PBS.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microfuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the cell pellet in 200 &#181;L of PBS + 1% bovine serum albumin (BSA). Incubate for 30 min at RT. 
		NOTE: Longer times are not necessary and are even detrimental to the efficiency of Click reactions.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the pellet in 300 &#181;L of PBS + 1% BSA.</li>
		<li>Distribute the cells between two tubes: 200 &#181;L in a 1.5 mL microcentrifuge tube for the Click reaction and 100 &#181;L in another 1.5 mL microcentrifuge tube for Sytox Green staining.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
		<li>For Sytox Green staining 
		NOTE: We highly recommend taking an aliquot for Sytox Green staining (without Click) in order to obtain high-quality DNA content reference profiles. Indeed, the Click reaction strongly quenches intercalant fluorescence, including the Sytox Green and PI fluorescence. Consequently, the Click reaction can distort the reading of DNA content.
		<ol>
			<li>Resuspend the cell pellet in 100 &#181;L of PBS.</li>
			<li>Transfer 10-30 &#181;L (depending on the cell concentration) to a flow cytometer tube containing 300 &#181;L of 50 mM Tris-HCl, pH 7.5, and 0.5 &#181;M Sytox Green.</li>
			<li>Sonicate 2x, for 2 s each time, at an amplitude of 40-50.</li>
			<li>Leave in the dark until processing the samples on a flow cytometer. 
			NOTE: The cells can be kept at this stage at 4 &#730;C for a few days.</li>
		</ol>
		</li>
		<li>For the Click reaction
		<ol>
			<li>Prepare fresh Azide Dye buffer by mixing the reagents in the following order (quantity for one tube): 36 &#181;L of PBS, 2 &#181;L of 0.2 M CuSO4, 0.2 &#181;L of 2 mM Cy5 Azide, and 2 &#181;L of 1 M ascorbic acid. 
			NOTE: It is possible to prepare a master mix of the Azide Dye buffer. The reagents have to be mixed in the same order as that mentioned above.</li>
			<li>Resuspend the cell pellet with 40 &#181;L of freshly made Azide Dye mix. Incubate at RT in the dark for 60 min.</li>
			<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
			<li>Wash the cells 3x with 300 &#181;L of 10% ethanol in PBS. 
			NOTE: The washes are crucial to eliminate all the soluble EdU-Cy5 azide.</li>
			<li>Resuspend the cells in 100 &#181;L of 50 &#181;g/mL PI in PBS. Leave for 10 min in the dark.</li>
			<li>Transfer 10-30 &#181;L of the cell suspension (depending on the cell concentration) to a flow cytometer tube containing 300 &#181;L of 50 mM Tris-HCl, pH 7.5.</li>
			<li>Sonicate 2x, for 2 s each time, at an amplitude of 40-50.</li>
			<li>Leave in the dark until processing the samples on a cytometer. 
			NOTE: The cells can be kept at this stage at 4 &#730;C for a few days.</li>
		</ol>
		</li>
		<li>Read the Sytox Green samples using an excitation blue laser at 488 nm and a 530/30 BP filter. See Figure 1A for typical results. Read the bivariate PI-EdU samples on a dot plot using an excitation blue laser at 488 nm and a 615/20 BP filter for the PI (x-axis) and an excitation red laser at 640 nm and a 660/20 BP filter (y-axis). See Figure 1B for typical results. 
		NOTE: Figure 1C represents the typical PI-EdU bivariate FACS result for EdU-negative cells. It allows for the discrimination of the EdU-negative cells from the EdU-positive cells.</li>
	</ol>
	</li>
	<li>For microscopy analysis
	<ol>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the pellet in 200 &#181;L of PBS + 1% BSA. Incubate for 30 min at RT.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microfuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Prepare fresh Azide Dye buffer by mixing the reagents in the following order (quantity for one tube): 36 &#181;L of PBS, 2 &#181;L of 0.2 M CuSO4, 0.2 &#181;L of 2 mM Dy-530 azide, 2 &#181;L of 1 M ascorbic acid. 
		NOTE: A master mix of the Azide Dye buffer can be prepared fresh by mixing the reagents in the aforementioned order.</li>
		<li>Resuspend the pellet with 40 &#181;L of freshly made Azide Dye buffer. Incubate at RT in the dark for 60 min.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microfuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Wash the cells 2x with 300 &#181;L of 10% ethanol in PBS. 
		NOTE: The washes are crucial to eliminate all soluble Dy-530 azide.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the cells in 100 &#181;L of 0.5 &#181;g/mL 4&#39;,6-diamidino-2-phenylindole (DAPI) in PBS. Leave for 30 min in the dark at room temperature.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microcentrifuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Wash with 300 &#181;L of PBS to remove excess DAPI.</li>
		<li>Pellet the cells for 2 min at 10,000 &#215; g in a microfuge. Discard the supernatant with a vacuum pipette.</li>
		<li>Resuspend the pellet with 10-50 &#181;L of PBS depending on the cell concentration.</li>
		<li>Sonicate 2x, for 2 s each time, at an amplitude of 40-50. 
		NOTE: The cells can be kept at this stage at 4 &#730;C for a few days.</li>
		<li>Pipette 1.7 &#181;L of the cells onto a glass microscope slide and cover with a clean coverslip.</li>
		<li>Immediately observe under a fluorescence microscope with DAPI and TexasRed or Cy3 filters.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

Yeast is a prime model organism for cell cycle studies, yet the characterization of its S phase has long been hampered by its inability to incorporate exogenous nucleosides, such as BrdU, which are used as tracers of DNA replication. Equipping yeast with a high expression of herpes simplex thymidine kinase (TK) and the addition of a human nucleoside transporter (hENT) has largely solved this problem15,16. EdU is more versatile than BrdU as its detection with smal...

Genome stability through mitotic division is ensured by the transmission of a complete and equal set of chromosomes to the two produced cell progenies. This relies on the accurate completion of a series of events occurring in a given time in each phase of the cell cycle. In G1, the replication origins are licensed upon the recruitment of several licensing factors, including Cdc61. In the S phase, whole-genome duplication is initiated from multiple active replication origins and performed by replication machineries that gather in microscopically visible foci named replication factories2. In the M phase, duplicated sister chromatids are attached and bioriented on the mitotic spindle to allow their segregation to the opposite poles of the mitotic cell3. The regulation, proper completion, and duration of each phase are key to ensure genome stability. Indeed, premature exit from any of these phases leads to genome instability. For instance, a shorter G1 induced by deletion of the budding yeast CDK inhibitor Sic1 or by the overexpression of G1 cyclins will alter the subsequent S phase4,5,6. Consequently, these deregulations, associated or not with replication stress, result in chromosome breaks, rearrangements, and mis-segregation4,5,6. Therefore, monitoring the duration of the S phase and, more broadly, the duration of the other phases of the cell cycle may be crucial to identify the defects occurring in different mutants and in different stressful conditions.
A traditional method for measuring cell cycle phase duration includes simple DNA content flow cytometry (Figure 1A) and relies on a fitting algorithm (available in most cytometry software) used to separate the population into G1, S, and G2 + M phase fractions from the 1C and 2C peaks. The fractions are then multiplied by the population doubling time7. However, this method gives only estimated values, requires a homogeneous cell size distribution within a given fraction, and is not applicable to synchronized cultures. To study the S-phase duration in mammalian cells, several thymidine analogs have been developed and widely used, including EdU. Their uptake from the extracellular medium and phosphorylation by thymidine kinase (hereafter referred to as TK) make them available for DNA polymerases to incorporate them at sites of DNA synthesis (replication, recombination, repair). To bypass the absence of the TK gene in Saccharomyces cerevisiae cells, yeast strains have been engineered to allow stable and constitutive expression of the herpes simplex virus TK8 and the human equilibrative nucleoside transporter (hENT1)9. Once incorporated into DNA, EdU is detected via the selective Click reaction, which chemically couples its alkyne moiety to azide-modified fluorochromes10.
This paper provides two optimized comprehensive protocols to pulse-label asynchronous and synchronous TK-hENT1 engineered cells with EdU in order to precisely visualize and measure DNA replication duration and dynamics, as well as the duration of the other phases of the cell cycle, with high spatial and temporal resolution at both the single-cell and population levels by microscopy and flow cytometry.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#945;-factor&#160;</td>
			<td>Genescript</td>
			<td>RP01002</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Euromedex</td>
			<td>04-100--812-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper sulfate</td>
			<td>Sigma</td>
			<td>C1297</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma</td>
			<td>D9542</td>
			<td></td>
		</tr>
		<tr>
			<td>Di-sulfo-Cyanine5 azide (Cy5 azide)</td>
			<td>Interchim</td>
			<td>FP-JV6320</td>
			<td>Alternative to Alexa647-Azide</td>
		</tr>
		<tr>
			<td>Dy-530 azide</td>
			<td>Dyomics</td>
			<td>&#160;530-10</td>
			<td></td>
		</tr>
		<tr>
			<td>EdU (5-ethynyl-2&#8217;-deoxyuridine)</td>
			<td>Carbosynth</td>
			<td>NE08701</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol absolute</td>
			<td>Carlo Erba reagents</td>
			<td>P013A10D16</td>
			<td>or equivalent</td>
		</tr>
		<tr>
			<td>L- ascorbic acid</td>
			<td>Sigma</td>
			<td>A4544</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium iodide</td>
			<td>Sigma</td>
			<td>P4864</td>
			<td></td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>Euromedex</td>
			<td>EU0090</td>
			<td></td>
		</tr>
		<tr>
			<td>Rnase</td>
			<td>SIGMA</td>
			<td>R5000</td>
			<td></td>
		</tr>
		<tr>
			<td>Sytox Green</td>
			<td>Invitrogen</td>
			<td>S-7020</td>
			<td></td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cell counter</td>
			<td>OLS</td>
			<td>CASY</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow cytometer</td>
			<td>Agilent</td>
			<td>NovoSampler Pro</td>
			<td></td>
		</tr>
		<tr>
			<td>Shaking incubator</td>
			<td>Infors</td>
			<td>444-4230</td>
			<td>or equivalent</td>
		</tr>
		<tr>
			<td>Shaking water bath</td>
			<td>Julabo</td>
			<td>SW22</td>
			<td>or equivalent</td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Sonics</td>
			<td>Vibra cell</td>
			<td></td>
		</tr>
		<tr>
			<td>Wide-field microscopy</td>
			<td>Leica</td>
			<td>THUNDER Imager</td>
			<td>or equivalent</td>
		</tr>
	</tbody>
</table>

determination of s-phase duration using 5-ethynyl-2'-deoxyuridine incorporation in saccharomyces cerevisiae

Eukaryotic DNA replication is a highly regulated process that ensures that the genetic blueprint of a cell is correctly duplicated prior to chromosome segregation. As DNA synthesis defects underlie chromosome rearrangements, monitoring DNA replication has become essential to understand the basis of genome instability. Saccharomyces cerevisiae is a classical model to study cell cycle regulation, but key DNA replication parameters, such as the fraction of cells in the S phase or the S-phase duration, are still difficult to determine. This protocol uses short and non-toxic pulses of 5-ethynyl-2'-deoxyuridine (EdU), a thymidine analog, in engineered TK-hENT1 yeast cells, followed by its detection by Click reaction to allow the visualization and quantification of DNA replication with high spatial and temporal resolution at both the single-cell and population levels by microscopy and flow cytometry. This method may identify previously overlooked defects in the S phase and cell cycle progression of yeast mutants, thereby allowing the characterization of new players essential for ensuring genome stability.

To determine the S-phase duration and, more broadly, the duration of G1 and G2 + M (protocol step 1.1), S. cerevisiae W303 wild-type cells (WT, Table 1) were grown asynchronously in SC medium for 7 h. Every hour, the cell concentration was monitored to determine the doubling time (Figure 2B). In these growth conditions, the calculated doubling time was 120 min &#177; 13 min at 25 &#730;C (Table 2). When the cells were in the ex...

Watch this Scientific Journal Video about Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation in Saccharomyces cerevisiae at JoVE.com

Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation in Saccharomyces cerevisiae

We describe two complementary protocols to accurately determine S-phase duration in S. cerevisiae using EdU, a thymidine analog, which is incorporated in vivo and detected using Click chemistry by microscopy and flow cytometry. It allows for the easy characterization of the duration of DNA replication and overlooked replication defects in mutants.

Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation in Saccharomyces cerevisiae

Eukaryotic DNA replication is a highly regulated process that ensures that the genetic blueprint of a cell is correctly duplicated prior to chromosome ...

determination-s-phase-duration-using-5-ethynyl-2-deoxyuridine

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JoVE Journal

Biology

1.3K Views.  Université de Montpellier, CNRS. We describe two complementary protocols to accurately determine S-phase duration in S. cerevisiae using EdU, a thymidine analog, which is incorporated in vivo and detected using Click chemistry by microscopy and flow cytometry. It allows for the easy characterization of the duration of DNA replication and overlooked replication defects in mutants.

Video: Determination of S-Phase Duration Using 5-Ethynyl-2'-deoxyuridine Incorporation 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 ...

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 ...

Rapid Identification of Chemical Genetic Interactions in Saccharomyces cerevisiae

Determining the mode of action of bioactive chemicals is of interest to a broad range of academic, pharmaceutical, and industrial scientists. Saccharomyces cerevisiae, or budding yeast, is a model eukaryote for which a complete collection of ~6,000 gene deletion mutants and hypomorphic essential gene mutants are commercially available. These collections of mutants can be used to systematically detect chemical-gene interactions, i.e. genes necessary to tolerate a chemical. This information, in turn, reports on the likely mode of action of the compound. Here we describe a protocol for the rapid identification of chemical-genetic interactions in budding yeast. We demonstrate the method using the chemotherapeutic agent 5-fluorouracil (5-FU), which has a well-defined mechanism of action. Our results show that the nuclear TRAMP RNA exosome and DNA repair enzymes are needed for proliferation in the presence of 5-FU, which is consistent with previous microarray based bar-coding chemical genetic approaches and the knowledge that 5-FU adversely affects both RNA and DNA metabolism. The required validation protocols of these high-throughput screens are also described.

Rapid Identification of Chemical Genetic Interactions in Saccharomyces cerevisiae

Here we present a cost-effective method for defining chemical-genetic interactions in budding yeast. The approach is built on fundamental techniques in yeast molecular biology and is well suited for the mechanistic interrogation of small to medium collections of chemicals and other media environments.

Determining the mode of action of bioactive chemicals is of interest to a broad range of academic, pharmaceutical, and industrial scientists. ...

Growth-based Determination and Biochemical Confirmation of Genetic Requirements for Protein Degradation in Saccharomyces cerevisiae

Regulated protein degradation is crucial for virtually every cellular function. Much of what is known about the molecular mechanisms and genetic requirements for eukaryotic protein degradation was initially established in Saccharomyces cerevisiae. Classical analyses of protein degradation have relied on biochemical pulse-chase and cycloheximide-chase methodologies. While these techniques provide sensitive means for observing protein degradation, they are laborious, time-consuming, and low-throughput. These approaches are not amenable to rapid or large-scale screening for mutations that prevent protein degradation. Here, a yeast growth-based assay for the facile identification of genetic requirements for protein degradation is described. In this assay, a reporter enzyme required for growth under specific selective conditions is fused to an unstable protein. Cells lacking the endogenous reporter enzyme but expressing the fusion protein can grow under selective conditions only when the fusion protein is stabilized (i.e. when protein degradation is compromised). In the growth assay described here, serial dilutions of wild-type and mutant yeast cells harboring a plasmid encoding a fusion protein are spotted onto selective and non-selective medium. Growth under selective conditions is consistent with degradation impairment by a given mutation. Increased protein abundance should be biochemically confirmed. A method for the rapid extraction of yeast proteins in a form suitable for electrophoresis and western blotting is also demonstrated. A growth-based readout for protein stability, combined with a simple protocol for protein extraction for biochemical analysis, facilitates rapid identification of genetic requirements for protein degradation. These techniques can be adapted to monitor degradation of a variety of short-lived proteins. In the example presented, the His3 enzyme, which is required for histidine biosynthesis, was fused to Deg1-Sec62. Deg1-Sec62 is targeted for degradation after it aberrantly engages the endoplasmic reticulum translocon. Cells harboring Deg1-Sec62-His3 were able to grow under selective conditions when the protein was stabilized.

Growth-based Determination and Biochemical Confirmation of Genetic Requirements for Protein Degradation in Saccharomyces cerevisiae

This article describes a yeast growth-based assay for the determination of genetic requirements for protein degradation. It also demonstrates a method for rapid extraction of yeast proteins, suitable for western blotting to biochemically confirm degradation requirements. These techniques can be adapted to monitor degradation of a variety of proteins.