Name: a fluorescence microscopy assay for monitoring mitophagy in the yeast saccharomyces cerevisiae
Uploaded: 2011-07-18T12:00:00
Duration: 10 min 27 s
Description: Watch this Scientific Journal Video about A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae at JoVE.com

This work was supported by Australian Research Council Grant DP0986937.

Selecting suitable control yeast strains
The assay relies on the experimenter visually evaluating the accumulation of red fluorescence in the yeast vacuole. As such the assay is subjective and relies on the selection of suitable control strains and growth conditions. A wild type control is used to indicate the levels of autophagy normally expected. As a negative control a strain null for expression of one of the core autophagy genes (e.g., ATG3) is suitable; such strains cannot deliver material (e.g., mitochondria, nucleus) to the vacuole
1. Transformation of yeast cells with plasmid DNA
Yeast cells can be easily made competent for transformation by treating with LiAcetate. Commercial kits may be purchased (e.g., EasyComp, Invitrogen) and published protocols are available13.
In this protocol we use wild type strain BY4741 (MATa his3&Delta;1 leu2&Delta;0 met15&Delta;0 ura3&Delta;0), and some members of a comprehensive library of deletion mutant strains (e.g., &Delta;atg3) derived from it, each lacking expression of a different non-essential gene. The deletion strain library represents a valuable tool for investigating autophagy using Rosella-based probes. 
The reporter employed in this protocol is mt-Rosella for mitochondria (Fig. 2A), encoded by pAS1NB:mt-Rosella12.
The following steps are based on the use of cells made competent using the EasyComp Transformation Kit and stored frozen at -80&deg;C for convenient use.
<ol>
	<li>Equilibrate Solution III (EasyComp Transformation Kit) to room temperature 30 minutes before adding to yeast cells.</li>
	<li>Add 2-2.5 &mu;l (~100 ng/&mu;l) of plasmid DNA encoding the reporter (pAS1NB:mt-Rosella) to a sterile 1.7 ml snap-cap plastic microcentrifuge tube. </li>
	<li>Add 5 &mu;l of freshly thawed suspension of competent yeast cells and mix by gentle pipetting. </li>
	<li>Add 50 &mu;l of Solution III (EasyComp Transformation Kit) and vortex to mix. 1.5) Incubate the transformation reactions for 1 hour at 28-30&deg;C with vigorous mixing every 15 minutes using a vortex mixer or manually by flicking the tube with the finger.</li>
	<li>Carefully transfer the transformation mixture to a single YMM agar growth plate supplemented with histidine, methionine and uracil, and gently distribute the cells around the surface of the agar with a sterile glass spreader. Yeast cells are fragile at this stage and gentle handling can increase the yield of transformants.</li>
	<li>Incubate growth plates for 2 to 4 days at 28-30&deg;C or until colonies appear about 2-3 mm in diameter.</li>
</ol>
2. Confirming expression of Rosella in yeast cells 
Before embarking on detailed experiments the correct localization and efficient expression of Rosella should be confirmed.
<ol>
	<li>Using sterile toothpicks select for each strain 5 single colonies from the transformation plate and resuspend cells in 50 &mu;l of sterile water in separate sterile 1.7 ml snap-cap plastic microcentrifuge tube.</li>
	<li>Place 10 &mu;l of each cell suspension onto separate labeled glass microscope slides (76 x 26 mm) and cover the cell suspension with a square coverslip (22 x 22 mm).</li>
	<li>Immediately examine the cells for fluorescence emission using a fluorescence microscope (e.g., Olympus Fluoview FV500). Look for strong green and red fluorescence and labeling typical of mitochondrial localization (Fig. 2B and 2C).</li>
	<li>The remainder of each colony examined for fluorescence in this way should be inoculated onto a fresh YMM growth plate. </li>
	<li>Incubate plates at 28-30&deg;C for 2 days until colony are approximately 2-3 mm in diameter.</li>
</ol>
3. Cell growth and induction of autophagy
Autophagy can be induced using a number of different experimental conditions, including entry into stationary phase, change in carbon source, administration of rapamycin, or nitrogen starvation. We routinely use nitrogen starvation as the method to induce mitophagy. 
<ol>
	<li>Inoculate a single colony into 10 ml of growth medium containing ethanol and the appropriate auxotrophic supplements. </li>
	<li>Incubate yeast cultures at 28-30&deg;C with orbital shaking (200 rpm) for approximately 48 hours. The cells should be in mid-log growth phase.</li>
	<li>Pellet cells from duplicate 1ml samples of culture in sterile 1.7 ml snap-cap plastic centrifuge tubes by centrifugation at 2,000 x g for 2 minutes at room temperature. Carefully remove and discard the supernatant without disturbing the cell pellet.</li>
	<li>Wash the cells three times with 1 ml sterile distilled water by resuspension followed by centrifugation to remove residual medium.</li>
	<li>After the third wash resuspend the cells in 100 ml of growth medium or nitrogen-starvation medium.</li>
	<li>Re-inoculate the resuspended cells into 10 ml of fresh growth medium or 10 ml nitrogen-starvation medium. Incubate with orbital shaking for 6-12 hours to allow induction of mitophagy. After starvation prepare cells for viewing by fluorescence microscopy.</li>
</ol>
Note: Labeling of the vacuole with CMAC-Arg
When first establishing the assay it is useful to confirm under the fluorescence microscope delivery of Rosella to the vacuole. Although the yeast vacuole is a large organelle whose location can often easily be determined by reference to transmitted light images, in some strains of yeast and under some growth conditions the vacuole can be fragmented and difficult to locate. 
The vacuole can be readily labeled using a coumarin-based vacuole dye such as CMAC-Arg (7-amino-4-chloromethylcoumarin, L-arginine amide). The action of vacuolar proteases on the dye results in bright blue fluorescence labeling of the vacuole. The blue emission can be readily distinguished from the red and green emissions of Rosella11. Labeling with CMAC-Arg does not need to be performed routinely, but it is recommended for those who are not familiar with the location and appearance of the vacuole in yeast.
4. Labeling of the vacuole with CMAC-Arg
<ol>
	<li>Add the CMAC-Arg (100 mM final concentration) to growing or nitrogen-starved cells prior to imaging. The CMAC-Arg solution can be prepared in advance, but as the solution is light sensitive it needs to be protected from exposure to light.</li>
	<li>Incubate at 28-30&deg;C with orbital shaking (200 rpm) for 30 minutes.</li>
	<li>Pellet cells by centrifugation at 2,000 x g for 2 minutes at room temperature.</li>
	<li>Discard the supernatant.</li>
	<li>Wash the cells three times with sterile distilled water by resuspension followed by centrifugation to remove residual medium and excess CMAC-Arg dye.</li>
	<li>Mount the cells for imaging as described below. </li>
</ol>
5. Mounting live yeast cells for confocal laser scanning microscopy (CLSM)
<ol>
	<li>Melt 2 mg of low melting point agarose in separate 1 ml aliquots of growth medium and nitrogen-starvation medium by incubating at 70&deg;C. The molten agarose is maintained at 70&deg;C.</li>
	<li>Gently pellet the cells in 1 ml aliquots of each cell culture by centrifugation at 2,000 x g for 2 minutes at room temperature and discard the supernatants.</li>
	<li>Wash the cells three times with 1 ml sterile distilled water by resuspension and centrifugation as in step 5.2.</li>
	<li>Resuspend the washed cells in 100 &mu;l of sterile distilled water.</li>
	<li>To mount the cells spot 20 &mu;l of molten agarose onto a labelled glass microscope slide (76 x 26 mm).</li>
	<li>Immediately spot 10 &mu;l of the washed cell suspension onto the agarose bed.</li>
	<li>Cover the agarose bed with a coverslip (22 x 22 mm). The cells will spread across the agarose surface beneath the coverslip.</li>
	<li>To prevent dehydration and movement of the coverslip during imaging carefully seal the edges of coverslip with a thin film of nail polish. </li>
	<li>When the nail polish is completely dry (10 min) the slide is ready to view by fluorescence microscopy Beware: nail polish can cause damage to microscope objectives. </li>
	<li>Cells mounted using this technique can be observed up to 1 hour after mounting.</li>
</ol>
6. Visualizing autophagy using CLSM
The level of autophagy in a population of cells is routinely assessed by determining the number of cells showing red vacuolar fluorescence before and after the induction of autophagy. Ideally the progression of autophagy is monitored at selected time points following induction of autophagy (Fig. 3). This is best performed by visualizing through the microscope binocular. It is important that the correct controls are employed (e.g., &Delta;atg3 mutant) and that the microscope settings remain unchanged between observing different samples (e.g., use of neutral density filters, filter selection).
Yeast cells are relatively small requiring a good quality fluorescence microscope (e.g., Olympus Fluoview FV500) equipped with excitation/emission filters suitable for the separate visualization of GFP and/or pHluorin (green fluorescence emission) (FITC filter) and DsRed.T3 (red fluorescence emission) (TRITC filter).
<ol>
	<li>Using a 60x water objective view non-starved wild type cells using alternately the FITC and TRITC filters. Wild type cells should exhibit a typical cellular distribution of mitochondria at the periphery of the cells that show both red and green fluorescence. Note that this distribution of mitochondria is dependent upon the pinhole setting of the confocal microscope. We generally use a lower pinhole value of 125 &mu;m that only allows the mitochondria to be observed as a series of dots localized at the periphery of the cell (Fig. 2C) so that the vacuole is not obscured12. If a high pinhole value of (200 &mu;m) is used this allows the typical filamentous mitochondrial network distributed throughout the cell (and the changes it undergoes) to be observed11. No other structures in the cell should be fluorescently labeled.</li>
	<li>Sequentially view cells for each of the time points following transfer to starvation medium. Visualize alternately using the green and red emission filters. At the 6 h starvation time point red fluorescence, but no corresponding green emission should be discernable in the vacuole. Vacuolar localization can be separately confirmed using CMAC-Arg (Fig. 2C). View ~200 cells and count the number that show only red fluorescence in the vacuole (i.e. vacuolar uptake of mitochondria).</li>
	<li>Count vacuolar accumulation for all time points observing &gt;200 cells and then plot percentage of cells showing vacuolar accumulation. </li>
	<li>Examine autophagy-negative control cells expressing mt-Rosella before starvation and following 6 h starvation. </li>
</ol>
7. Representative results:
Here, we show typical results obtained using mt-Rosella, expressed in wild type and &Delta;atg3 mutant cells. Under growing conditions with ethanol as carbon source, both wild type and &Delta;atg3 mutant cells exhibit a cellular distribution of fluorescence (red and green) typical of mitochondria in yeast cells. Red and green fluorescence emission is not detected in the vacuole (Fig. 2B and 2C). When subjected to nitrogen starvation for 6 h and beyond then, in addition to red and green fluorescence corresponding to mitochondria, wild type cells also exhibit the accumulation of red, but not green fluorescence in the vacuolar lumen (Fig. 2B; Fig. 3). However, in &Delta;atg3 mutant cells neither red nor green fluorescence accumulates in the vacuolar lumen (Fig. 2C; Fig. 3). 
<img src="/files/ftp_upload/2779/2779fig1.jpg" alt="Figure 1" /> 
Figure 1. Top, scheme of organelles and compartments in a yeast cell. Bottom, a vacuole-centric view of a yeast cell is presented indicating autophagic degradation of different organelles and compartments. Some of the differences between the various organelle-specific types of autophagy (e.g., mitophagy, nucleophagy) include the specificity (cargo selection) of the engulfed contents, various intracellular needs and extracellular cues (e.g., starvation, damage), specific signals, ATG genes and time dependency. 
<img src="/files/ftp_upload/2779/2779fig2a.jpg" alt="Figure 2a" /> 
<img src="/files/ftp_upload/2779/2779fig2b.jpg" alt="Figure 2b" /> 
<img src="/files/ftp_upload/2779/2779fig2c.jpg" alt="Figure 2c" /> 
<img src="/files/ftp_upload/2779/2779fig2d.jpg" alt="Figure 2d" /> 
Figure 2. (A) Schematic representation of mt-Rosella. The mitochondrial leader sequence (in this case of citrate synthase) is used to target the Rosella biosensor to the mitochondrial matrix. Excitation and emission maxima for both red fluorescent protein (DsRed.T3) and green fluorescent protein (pHluorin) are indicated. At high pH (mitochondrial pH~8.2) the biosensor fluoresces both red and green, however, at low pH (vacuolar pH~5.5) it fluoresces only red.
(B) Schematic representation of expected results in wild type and &Delta;atg3 mutant cells expressing mt-Rosella. 
(C) Actual results obtained by CLSM for wild type cells under growing and nitrogen starvation conditions. From left to right: DIC (difference in contrast), RFP (DsRed.T3) fluorescence emission, GFP (pHluorin) fluorescence emission, CMAC-Arg fluorescence and merge image.
(D) Actual results obtained by fluorescence microscopy for Datg3 mutant cells under growing and nitrogen starvation conditions. From left to right: DIC (difference in contrast), RFP (DsRed.T3) fluorescence emission, GFP (pHluorin) fluorescence emission, CMAC-Arg fluorescence and merge image.
<img src="/files/ftp_upload/2779/2779fig3.jpg" alt="Figure 3" /> 
Figure 3. The percentage of wild type and &Delta;atg3 mutant cells, expressing mt-Rosella and grown under nitrogen starvation, showing red fluorescence in the vacuole over a time course of 48 hours. The percentage of cells showing accumulation of red fluorescence in the vacuole was recorded at 0, 6, 12, 24 and 48 hours after commencing nitrogen starvation.

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	<li>Levine, B., Kroemer, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagy+in+the+pathogenesis+of+disease.">Autophagy in the pathogenesis of disease.</a> Cell. 132, 27-42 (2008).</li><li>Mizushima, N., Levine, B., Cuervo, A. M., Klionsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagy+fights+disease+through+cellular+self-digestion.">Autophagy fights disease through cellular self-digestion.</a> Nature. 451, 1069-1075 (2008).</li><li>He, C., Klionsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+mechanisms+and+signaling+pathways+of+autophagy.">Regulation mechanisms and signaling pathways of autophagy.</a> Annu. Rev. Genet. 43, 67-93 (2009).</li><li>Kroemer, G., Mari&#241;o, G., Levine, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagy+and+the+integrated+stress+response.">Autophagy and the integrated stress response.</a> Mol Cell. 40, 280-293 (2010).</li><li>Camougrand, N., Kissov&#225;, I., Salin, B., Devenish, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+mitophagy+in+yeast.">Monitoring mitophagy in yeast.</a> Methods Enzymol. 451, 89-107 (2008).</li><li>Yen, W. L., Klionsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+to+live+long+and+prosper:+autophagy,+mitochondria,+and+aging.">How to live long and prosper: autophagy, mitochondria, and aging.</a> Physiology (Bethesda). 23, 248-262 (2008).</li><li>Tolkovsky, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitophagy.">Mitophagy.</a> Biochim. Biophys. Acta. 1793, 1508-1515 (2009).</li><li>Bhatia-Kissov&#225;, I., Camougrand, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitophagy+in+yeast:+actors+and+physiological+roles.">Mitophagy in yeast: actors and physiological roles.</a> FEMS Yeast Res. 10, 1023-1034 (2010).</li><li>Kanki, T., Klionsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+mechanism+of+mitochondria+autophagy+in+yeast.">The molecular mechanism of mitochondria autophagy in yeast.</a> Mol. Microbiol. 75, 795-800 (2010).</li><li>Nowikovsky, K., Reipert, S., Devenish, R. J., Schweyen, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mdm38+protein+depletion+causes+loss+of+mitochondrial+K+/H++exchange+activity,+osmotic+swelling+and+mitophagy.">Mdm38 protein depletion causes loss of mitochondrial K+/H+ exchange activity, osmotic swelling and mitophagy.</a> Cell Death Differ. 14, 1647-1656 (2007).</li><li>Devenish, R. J., Prescott, M., Turcic, K., Mijaljica, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+organelle+turnover+in+yeast+using+fluorescent+protein+tags.">Monitoring organelle turnover in yeast using fluorescent protein tags.</a> Methods Enzymol. 451, 109-131 (2008).</li><li>Rosado, C. J., Mijaljica, D., Hatzinisiriou, I., Prescott, M., Devenish, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rosella:+a+fluorescent+pH-biosensor+for+reporting+vacuolar+turnover+of+cytosol+and+organelles+in+yeast.">Rosella: a fluorescent pH-biosensor for reporting vacuolar turnover of cytosol and organelles in yeast.</a> Autophagy. 4, 205-213 (2008).</li><li>Gietz, R. D., Woods, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transformation+of+yeast+by+lithium+acetate/single-stranded+carrier+DNA/polyethylene+glycol+method.">Transformation of yeast by lithium acetate/single-stranded carrier DNA/polyethylene glycol method.</a> Methods Enzymol. 350, 87-96 (2002).</li></ol>

No conflicts of interest declared.

In the representative images presented in Fig. 2B and 2C it can be clearly seen that the use of organelle-specific fluorescent reporters and fluorescence microscopy provides evidence of mitochondrial localization/distribution in both growing and nitrogen-starved cells, and allows mitophagy to be visualized. 
It is to be emphasized that this method is not restricted to following mitophagy. Autophagy of other cellular components or organelles (e.g., ribosomes, lipid droplets, peroxisomes, endoplasmic reticulum and nucleus) may be monitored with suitable labeling. Details of the Rosella biosensor construct, cell culture conditions and typical results for turnover of the nucleus (nucleophagy) have been reported11,12 illustrating the broader application of the method.
Use of organelle-specific fluorescent reporters for monitoring autophagy requires several considerations of construct design to be kept in mind. These include: (1) selection and fusion to the appropriate targeting signal to achieve organelle-specific targeting; (2) selection of a suitable fluorescent protein(s); (3) strong fluorescence emission; (4) correct organelle localization and/or distribution within the cell. Once the organelle-specific reporter has been successfully expressed in the chosen yeast host strain a second group of considerations become: (1) yeast growth conditions; (2) cell sample preparation; (3) imaging parameters as it is important that the parameters selected for CLSM (e.g., laser intensity, scan rate and mode, pinhole value, zoom value, objectives used or exposure for CCD-camera) are maintained throughout an experiment ensuring that comparisons can be made between control (growing cells) and starved samples; (3) autophagy induction and successful delivery of the reporter into the vacuole.
Overall, this fluorescent microscopy method is easy, relatively fast, and has potential application for high throughput screening for novel drugs that enhance or inhibit autophagy, and also for genes that regulate or modulate autophagy.

<table border="1">
	<tbody>
		<tr>
			<th>Name of the reagent/equipment</th>
			<th>Company</th>
			<th>Catalogue number</th>
		</tr>
		<tr>
			<td>Saccharomyces cerevisiae EasyComp Transformation Kit</td>
			<td>Invitrogen</td>
			<td>K5050-01</td>
		</tr>
		<tr>
			<td>Yeast nitrogen base without amino acids and ammonium sulfate</td>
			<td>BD Difco</td>
			<td>200-0030</td>
		</tr>
		<tr>
			<td>Low melting agarose</td>
			<td>Progen Biosciences</td>
			<td>200-0030</td>
		</tr>
		<tr>
			<td>Microscope slides (76 x 26 mm)</td>
			<td>Menzel-Gl&auml;ser</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Microscope coverslips (22 x 22 mm)</td>
			<td>Menzel-Gl&auml;ser</td>
			<td>BB022022A1</td>
		</tr>
		<tr>
			<td>Confocal Laser Scanning Microscope Fluoview FV500</td>
			<td>Olympus</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CMAC-Arg</td>
			<td>Moleular Probes-Invitrogen</td>
			<td>Y-7531</td>
		</tr>
	</tbody>
</table>
All concentrations given are w/v, unless otherwise indicated. 
Yeast minimal medium (YMM): 0.67% yeast nitrogen base without amino acids, 2% glucose, 1.5% agar (only added when making solid media) and any required auxotrophic supplements (0.01%)]. 
Saccharomyces Salts (SS): (NH4)2S04, 0.12%; KH2PO4, 0.10%; MgC12.6H2O, 0.07%; NaC1, 0.05%; CaCl2, 0.01%; FeC13, 0.0005%. 
Growth medium: SS and 2% ethanol (v/v) and appropriate auxotrophic supplements. 
Nitrogen-starvation medium: 0.17% yeast nitrogen base without amino acids and ammonium sulfate (Difco), 2% ethanol (v/v).

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.

Watch this Scientific Journal Video about A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae at JoVE.com

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.

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 ...

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Dissection of Saccharomyces Cerevisiae Asci

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

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Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

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Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

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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 ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

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 ...

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. ...

Cycloheximide Chase Analysis of Protein Degradation in Saccharomyces cerevisiae

Regulation of protein abundance is crucial to virtually every cellular process. Protein abundance reflects the integration of the rates of protein synthesis and protein degradation. Many assays reporting on protein abundance (e.g., single-time point western blotting, flow cytometry, fluorescence microscopy, or growth-based reporter assays) do not allow discrimination of the relative effects of translation and proteolysis on protein levels. This article describes the use of cycloheximide chase followed by western blotting to specifically analyze protein degradation in the model unicellular eukaryote, Saccharomyces cerevisiae (budding yeast). In this procedure, yeast cells are incubated in the presence of the translational inhibitor cycloheximide. Aliquots of cells are collected immediately after and at specific time points following addition of cycloheximide. Cells are lysed, and the lysates are separated by polyacrylamide gel electrophoresis for western blot analysis of protein abundance at each time point. The cycloheximide chase procedure permits visualization of the degradation kinetics of the steady state population of a variety of cellular proteins. The procedure may be used to investigate the genetic requirements for and environmental influences on protein degradation.

Cycloheximide Chase Analysis of Protein Degradation in Saccharomyces cerevisiae

Protein abundance reflects the rates of both protein synthesis and protein degradation. This article describes the use of cycloheximide chase followed by western blotting to analyze protein degradation in the model unicellular eukaryote, Saccharomyces cerevisiae (budding yeast).