Name: assessment of submitochondrial protein localization in budding yeast saccharomyces cerevisiae
Uploaded: 2021-07-19T15:00:00.000+00:00
Duration: 8 min 55 s
Description: Watch this Scientific Journal Video about Assessment of Submitochondrial Protein Localization in Budding Yeast Saccharomyces cerevisiae at JoVE.com

We thank Dr. A. Tzagoloff (Columbia University) for providing antibodies raised against submitochondrial marker proteins Cyt. b2, &#945;KGD, and Sco1. We also thank Dr. Mario Henrique de Barros (Universidade de S&#227;o Paulo) for helpful discussion and comments during the establishment of this protocol.
This work was supported by research grants from Funda&#231;&#227;o de Amparo &#224; Pesquisa do Estado de S&#227;o Paulo (FAPESP) (grant 2013/07937-8).
Fernando Gomes and Helena Turano are also supported by FAPESP, grants 2017/09443-3 and 2017/23839-7, respectively. Ang&#233;lica Ramos is also supported by Coordena&#231;&#227;o de Aperfei&#231;oamento de Pessoal de N&#237;vel Superior (CAPES).

1. Growth of yeast cells
<ol>
	<li>Isolate single colonies of the strain of interest by streaking a small portion of the cells from a -80 &#176;C glycerol stock onto a YPD (1% yeast extract, 2% peptone, 2% glucose) agar plate. Incubate the plate at 30 &#176;C for 2-3 days. 
	NOTE: The S. cerevisiae strain used in this protocol is derived from BY4741 (MAT&#945;; his3&#916;1; leu2&#916;0; met15&#916;0; ura3&#916;0). With the exception of the auxotrophic marker genes, this strain does not contain any deleted gene and does not carry any plasmid. Thus, it can be successfully cultivated in a rich medium, stimulating vigorous cell growth. When working with strains transformed with plasmids, use the appropriate minimal medium for plasmid selection.</li>
	<li>Prepare a starter culture by inoculating 2-3 individual colonies from YPD agar plate in 10-20 mL of YPGal medium (1% yeast extract, 2% peptone, 2% galactose) in a 100 mL Erlenmeyer flask. Incubate at 30 &#176;C for 24 h with vigorous shaking. 
	NOTE: The choice of the growth medium depends on the yeast strain used in the protocol. Both YPD and YPGal contain fermentable carbon sources, which allow the growth of strains that do not perform mitochondrial respiration. However, since glucose represses the expression of many mitochondrial genes, it is not recommended to use this carbon source since it will produce lower amounts of mitochondria. When working with respiratory competent strains that can respire, it is also possible to use&#160;carbon sources such as glycerol and ethanol in an attempt to&#160;obtain a higher yield of mitochondria.</li>
	<li>Dilute the starter culture into 1 L of fresh YPGal medium to an OD600 less than 0.1. Cultivate the cells at 30 &#176;C with vigorous shaking until OD600 reaches 1-1.5. 
	&#8203;NOTE: Determine the growth rate (doubling time) for each yeast strain before performing the experiment. This will provide an accurate estimate of the time of incubation required for the culture to reach an OD600 of 1-1.5.</li>
</ol>
2. Isolation of highly purified mitochondria
NOTE: This protocol is adapted from17, with minor modifications.
<ol>
	<li>Harvest the cells by centrifugation at 3,000 x g for 5 min at room temperature.</li>
	<li>Wash the cells with distilled water and collect them by centrifugation at 3,000 x g for 5 min at room temperature.</li>
	<li>Determine the wet weight of the cells. 
	NOTE: The easiest way to measure the weight of the cell pellet from step 2.2 is to determine the weight of the empty centrifuge tube just before the collection of the cells. After centrifugation, discard the supernatant and measure the weight of the same tube with the cells. The weight of the cells is the difference between the two measurements.</li>
	<li>Resuspend the cells in DTT buffer (2 mL per 1 g of cells) using a Pasteur pipette or P5000 tip. See Table 1 for DDT buffer composition.</li>
	<li>Incubate the cells at 30 &#176;C for 20 min with gentle shaking (~70 rpm).</li>
	<li>Centrifuge at 3,000 x g for 5 min at room temperature to pellet the cells.</li>
	<li>Wash the cells with Zymolyase buffer without the enzyme (about 7 mL per 1 g of cells). 
	&#8203;See Table 1 for Zymolyase buffer composition.</li>
	<li>Centrifuge at 3,000 x g for 5 min at room temperature to pellet the cells.</li>
	<li>Resuspend the cells in the buffer without Zymolyase (7 mL per 1 g of cells).</li>
	<li>Transfer the cell suspension to a 250 mL Erlenmeyer flask and add Zymolyase-20T (3 mg per g wet weight).</li>
	<li>Incubate the cells at 30 &#176;C for 30-40 min with gentle shaking (~70 rpm). Check the efficiency of this process by comparing the turbidity of the cell suspension before and after Zymolyase treatment. 
	NOTE: In this step, the cells will be converted into spheroplasts due to cell-wall digestion by Zymolyase.
	<ol>
		<li>For this, add 50 &#181;L of each cell suspension to separate glass tubes containing 2 mL of water. After mixing vigorously, the turbidity of the cell suspension treated with Zymolyase should rapidly decrease due to the osmotic disruption of spheroplasts. On the other hand, the turbidity of the non-treated cell suspension should remain unchanged. 
		NOTE: The effects of turbidity can also be monitored by simple visual inspection or by measuring the OD600 of both samples. In the second case, the OD600 of the Zymolyase treated sample should be 10%-20% of the non-treated sample. An alternate method involves counting the cells in both samples by using light microscopy.</li>
		<li>If the yield of spheroplasts formation is low, add more Zymolyase and incubate for a further 15 min interval.</li>
	</ol>
	</li>
	<li>Harvest the spheroplasts by centrifugation at 2500 x g for 5 min at 4 &#176;C. 
	NOTE: All the further steps should be carried out fast and on ice or at 4 &#176;C to avoid protein degradation by hydrolytic enzymes.</li>
	<li>Wash the spheroplasts twice with ice-cold homogenization buffer (about 6.5 mL per 1 g of cells) and pellet by centrifuging at 2,500 x g for 5 min at 4 &#176;C. See Table 1 for homogenization buffer composition.</li>
	<li>Resuspend the spheroplasts in ice-cold homogenization buffer (6.5 mL per 1 g of cells) and transfer it to a pre-chilled glass Dounce homogenizer. Use a large glass homogenizer of approximately 30 mL.</li>
	<li>Homogenize the spheroplasts with 15 strokes using a pestle. 
	NOTE: The number of strokes should be adjusted depending on the pestle fitting. For tight pestle, 15 strokes are sufficient. On the other hand, if using a loose pestle, it is recommended to perform up to 25 strokes.</li>
	<li>Transfer the homogenate to a 50 mL centrifuge tube and add 1 volume of ice-cold homogenization buffer.</li>
	<li>Centrifuge the homogenate at low speed, 1,500 x g for 5 min at 4 &#176;C to pellet nuclei, cell debris, and unbroken cells.</li>
	<li>Transfer the supernatant to a new 50 mL centrifuge tube using a Pasteur pipette or P5000 tip taking care to avoid disrupting the pellet.</li>
	<li>Centrifuge at 4,000 x g for 5 min at 4 &#176;C.</li>
	<li>Transfer the supernatant to a high-speed centrifuge tube, and centrifuge at 12,000 x g for 15 min at 4 &#176;C to pellet the crude mitochondria fraction.</li>
	<li>Discard the supernatant and gently wash the crude mitochondrial pellet in 20-30 mL ice-cold homogenization buffer by gentle pipetting using a P5000 tip.</li>
	<li>Transfer the suspension to a 50 mL centrifuge tube and centrifuge at 4,000 x g for 5 min at 4 &#176;C to pellet the remaining cell debris.</li>
	<li>Transfer the supernatant to a high-speed centrifuge tube, and centrifuge at 12,000 x g for 15 min at 4 &#176;C to pellet the crude mitochondria fraction.</li>
	<li>Discard the supernatant and gently resuspend the crude mitochondrial pellet in a small volume (typically 1000 &#181;L) of ice-cold SEM buffer by gentle pipetting using a P1000 tip. See Table 1 for SEM buffer composition. 
	NOTE: Although this crude mitochondrial fraction can be used directly in some applications such as in organello protein import assays, it contains substantial amounts of other cellular components. These contaminations might lead to misinterpretations of the results when determining the submitochondrial localization of a protein. Therefore, further purification steps are required to get highly purified mitochondrial preparation, as described below.</li>
	<li>Prepare sucrose solutions in the EM buffer at concentrations of 60%, 32%, 23%, and 15% (w/v). These solutions are stable for up to 1 month at 4 &#176;C. See Table 1 for EM buffer composition.</li>
	<li>Prepare a 4-step sucrose gradient in an ultracentrifuge tube as follows: Place 1.5 mL of 60% (w/v) sucrose onto the bottom of the centrifuge tube. Next, pipette carefully stepwise: 4 mL of 32%, 1.5 mL of 23%, and 1.5 mL of 15% sucrose (w/v). Take care to avoid disrupting the phases.</li>
	<li>Carefully load the crude mitochondrial fraction on top of the sucrose gradient.</li>
	<li>Centrifuge for 1 h at 134,000 x g at 4 &#176;C in a swinging bucket rotor.</li>
	<li>Carefully keep removing the sucrose solution until the highly purified mitochondria fraction is reached which is represented by a brown band at the 60%/32% sucrose interface.</li>
	<li>Recover the purified mitochondria using a P1000 cut tip and place it into a pre-chilled high-speed centrifuge tube.</li>
	<li>Dilute the recovered mitochondria with 5-10 volumes of ice-cold SEM buffer.</li>
	<li>Centrifuge for 30 min at 12,000 x g at 4 &#176;C.</li>
	<li>Resuspend the pure mitochondria in 500 &#181;L of ice-cold SEM buffer by gentle pipetting using a P1000 cut tip.</li>
	<li>Determine the protein concentration of the highly purified mitochondrial preparation using the Bradford procedure following the manufacturer&#39;s instructions. Adjust the protein concentration to 10 mg of protein/mL with ice-cold SEM buffer. 
	&#8203;NOTE: For the submitochondrial fractionation protocol described below, it is recommended to use freshly prepared mitochondria. However, V&#246;gtle and collaborators performed a detailed quality control analysis of the mitochondrial intactness and showed that frozen organelles can also be used in this protocol13. For this, make aliquots of 40 &#181;L and freeze them in liquid nitrogen. Store at -80 &#176;C.</li>
</ol>
3. Submitochondrial fractionation protocol
NOTE: This protocol is adapted from reference18 and is composed of two steps: (1) hypotonic swelling in the presence or absence of proteinase K, and (2) sonication followed by carbonate extraction. Perform all the steps of both the protocols on ice or at 4 &#176;C to avoid protein degradation.
<ol>
	<li>Hypotonic swelling in the presence of proteinase K
	<ol>
		<li>Transfer 40 &#181;L of highly purified mitochondria at 10 mg/mL (400 &#181;g) into four 1.5 mL pre-chilled labeled microcentrifuge tubes.</li>
		<li>Add 360 &#181;L of SEM buffer in tubes 1 and 2.</li>
		<li>Add 360 &#181;L of EM buffer in tubes 3 and 4.</li>
		<li>Add 4 &#181;L of proteinase K (10 mg/mL) in tubes 2 and 4. Use the pipetting scheme listed in Table 2 to avoid mistakes. 
		NOTE: Prepare a 10 mg/mL solution of proteinase K in water immediately before use. The final concentration of proteinase K in the experiment should be around 50-100 &#181;g/mL. Please see the Discussion section for further details.</li>
		<li>Mix all the tubes gently and incubate on ice for 30 min with occasional mixing.</li>
		<li>Add 4 &#181;L of 200 mM PMSF to all the four tubes to stop proteinase K activity. 
		CAUTION: PMSF is highly toxic. Wear gloves when working with solutions containing PMSF.</li>
		<li>Centrifuge at 20,000 x g for 30 min at 4 &#176;C.</li>
		<li>Collect the supernatant and transfer it to a new 1.5 mL pre-chilled labeled microcentrifuge tube. Take care to avoid disrupting the pellet. 
		NOTE: The pellet can be directly resuspended in the sample buffer for further SDS-PAGE and western blot analysis. However, traces of proteinase K can remain active even after PMSF treatment and eventually can digest some proteins after the pellet has been dissolved in the SDS-containing sample buffer. To avoid this problem, proteinase K could be completely inactivated by treatment of the samples with trichloroacetic acid (TCA) as described below. 
		CAUTION: TCA is highly toxic. Wear gloves when working with solutions containing TCA.</li>
		<li>Resuspend the pellet from step 3.1.8 in 400 &#181;L of ice-cold SEM buffer.</li>
		<li>Precipitate the supernatant (from step 3.1.8) and the resuspended pellet (from step 3.1.9) with TCA to a final concentration of 10% (w/v).</li>
		<li>Incubate all the tubes on ice for 10 min.</li>
		<li>Centrifuge the TCA-treated samples for 10 min at 12,000 x g at 4 &#176;C.</li>
		<li>Remove the supernatant and resuspend the pellet in 200 &#181;L of sample buffer. 
		NOTE: It is possible that the bromophenol blue pH indicator turns yellow because of the acid treatment. If this happens, add small aliquots (1-5 &#181;L) of 1 M Tris base until it turns blue.</li>
		<li>Add 4 &#181;L of 200 mM PMSF to all the tubes.</li>
		<li>Store all the samples at -80 &#176;C until further analysis by SDS-PAGE and western blot.</li>
	</ol>
	</li>
	<li>Sonication and carbonate extraction 
	NOTE: In this protocol, it is not necessary to use freshly prepared mitochondria once the sonication causes the rupture of the mitochondrial membranes.
	<ol>
		<li>Transfer 200 &#181;L of highly purified mitochondria at 10 mg/mL (2 mg protein) into a 1.5 mL pre-chilled microcentrifuge tube.</li>
		<li>Dilute mitochondria one-fold with ice-cold SEM buffer.</li>
		<li>Sonicate mitochondria for 3 x 30 s on ice. Use a sonicator compatible for small volumes.</li>
		<li>Centrifuge the sample for 30 min at 100,000 x g at 4 &#176;C.</li>
		<li>Collect the supernatant and transfer it to a new 1.5 mL pre-chilled microcentrifuge tube. Keep it on ice. This sample will be named soluble protein fraction (S).</li>
		<li>Resuspend the pellet from step 3.2.4 in 400 &#181;L of ice-cold SEM buffer.</li>
		<li>Take 100 &#181;L of the resuspended pellet from step 3.2.6 and transfer it to a new 1.5 mL pre-chilled microcentrifuge tube. Keep it on ice. This sample will be named submitochondrial particles fraction (SMP).</li>
		<li>Dilute the remaining 300 &#181;L from step 3.2.6 one-fold with freshly prepared 200 mM sodium carbonate.</li>
		<li>Incubate the sample from step 3.2.8 on ice for 30 min.</li>
		<li>Centrifuge the sample for 30 min at 100,000 x g at 4 &#176;C.</li>
		<li>Collect the supernatant and transfer it to a new 1.5 mL pre-chilled microcentrifuge tube. Keep it on ice. This sample will be named carbonate supernatant fraction (CS).</li>
		<li>Resuspend the pellet from step 3.2.10 in 400 &#181;L of ice-cold SEM buffer. This sample will be named carbonate precipitated fraction (CP).</li>
		<li>Precipitate all the samples (S, SMP, CS, and CP) with TCA to a final concentration of 10% (w/v).</li>
		<li>Incubate all the tubes on ice for 10 min.</li>
		<li>Centrifuge the TCA-treated samples for 10 min at 12,000 x g at 4 &#176;C.</li>
		<li>Remove the supernatant and resuspend each pellet in the sample buffer. If the sample buffer becomes yellow, add small aliquots (1-5 &#181;L) of 1 M Tris base until it turns blue.</li>
		<li>Add 1 &#181;L of 200 mM PMSF to all the tubes.</li>
		<li>Store all the samples at -80 &#176;C until further analysis by SDS-PAGE and western blot.</li>
	</ol>
	</li>
</ol>

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M., Chojnacka, M., Chacinska, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+presence+of+disulfide+bonds+reveals+an+evolutionarily+conserved+mechanism+involved+in+mitochondrial+protein+translocase+assembly.">The presence of disulfide bonds reveals an evolutionarily conserved mechanism involved in mitochondrial protein translocase assembly.</a> Scientific Reports. 6, 27484 (2016).</li><li>Callegari, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TIM29+is+a+subunit+of+the+human+carrier+translocase+required+for+protein+transport.">TIM29 is a subunit of the human carrier translocase required for protein transport.</a> FEBS letters. 590 (23), 4147-4158 (2016).</li><li>Neupert, W., Herrmann, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translocation+of+proteins+into+mitochondria.">Translocation of proteins into mitochondria.</a> Annual Review of Biochemistry. 76, 723-749 (2007).</li><li>Meineke, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+outer+membrane+form+of+the+mitochondrial+protein+Mcr1+follows+a+TOM-independent+membrane+insertion+pathway.">The outer membrane form of the mitochondrial protein Mcr1 follows a TOM-independent membrane insertion pathway.</a> FEBS Letters. 582 (6), 855-860 (2008).</li></ol>

The authors declare no conflicts of interest.

The protocol presented here has been successfully used and continuously optimized for a long-time to determine the protein localization in the submitochondrial compartments13,14,18,21,22,23. The reliability and reproducibility of this protocol are strongly dependent on the purity and integrity of mitochondrial preparations<su...

Mitochondria are essential organelles of eukaryotic cells that play crucial roles in bioenergetics, cellular metabolism, and signaling pathways1. To properly execute these tasks, mitochondria rely on a unique set of proteins and lipids responsible for their structure and function. The budding yeast Saccharomyces cerevisiae has been widely used as a model system for investigations on mitochondrial processes, as well as for other organelles2. The mitochondrial genome codes for only eight proteins in yeast; the vast majority of mitochondrial proteins (~99%) are encoded by nuclear genes, which are translated on cytosolic ribosomes, and subsequently imported into their correct submitochondrial compartments by sophisticated protein import machineries3,4,5. Thus, mitochondrial biogenesis depends on the coordinated expression of both the nuclear and mitochondrial genomes6,7. Genetic mutations causing defects in mitochondrial biogenesis are associated with human diseases8,9,10.
In the past two decades, high-throughput proteomic studies targeting highly-purified mitochondria resulted in a comprehensive characterization of yeast mitochondrial proteome, which has been estimated&#160;to be composed of at least 900 proteins11,12,13,14. Although these studies provided valuable information, the suborganellar localization of each protein in the four mitochondrial subcompartments, namely, the outer membrane (OM), intermembrane space (IMS), inner membrane (IM), and matrix, is still required. This question was partially addressed with proteomic-wide studies of the two smaller mitochondrial subcompartments (OM and IMS)15,16. More recently, V&#246;gtle and collaborators made a major step forward by generating a high-quality global map of submitochondrial protein distribution in yeast. Using an integrated approach combining SILAC-based quantitative mass spectrometry, different submitochondrial fractionation protocols, and the data set from the OM and IMS proteomes, the authors assigned 818 proteins into the four mitochondrial subcompartments13.
Despite the advances achieved by these high-throughput proteomic studies, our knowledge about the submitochondrial proteome composition is far from being complete. Indeed, among 986 proteins reported by V&#246;gtle and collaborators as being localized into yeast mitochondria, 168 could not be assigned in any of the four submitochondrial compartments13. Moreover, the authors did not provide information about the membrane topology of proteins that were predicted to be peripherally attached to the periphery of mitochondrial membranes. For example, it is not possible to know if a protein that was assigned as peripherally attached to the inner membrane is facing the matrix or the intermembrane space. Apart from these missing data from the proteome-wide studies, there are conflicting information about the suborganellar localization of a significant number of mitochondrial proteins. One example is the protease Prd1, which has been assigned as an intermembrane space protein in the common databases such as Saccharomyces Genome Database (SGD) and Uniprot. Surprisingly, using a subfractionation protocol similar to that described here, V&#246;gtle and collaborators clearly showed that Prd1 is a genuine matrix protein13. As mentioned above, the submitochondrial localization of many mitochondrial proteins needs to be elucidated or reevaluated. Here, we provide a simple and reliable protocol to determine the suborganellar localization of yeast mitochondrial proteins. This protocol was developed and optimized by various research groups and has been routinely used to determine the submitochondrial localization, as well as the membrane topology of many mitochondrial proteins.

<table><tbody><tr><td>Bacto Peptone</td><td>BD</td><td>211677</td><td></td></tr><tr><td>Bacto Yeast extract</td><td>BD</td><td>212750</td><td></td></tr><tr><td>Beckman Ultra-Clear Centrifuge Tubes, 14 x 89 mm</td><td>Beckman Coulter</td><td>344059</td><td></td></tr><tr><td>Bovine serum albumin (BSA fatty acid free)</td><td>Sigma-Aldrich</td><td>A7030</td><td>Component of Homogenization buffer</td></tr><tr><td>DL-Dithiothreitol</td><td>Sigma-Aldrich</td><td>43815</td><td>Component of DDT buffer</td></tr><tr><td>D-Sorbitol</td><td>Sigma-Aldrich</td><td>S1876</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Sigma-Aldrich</td><td>E9884</td><td></td></tr><tr><td>Galactose</td><td>Sigma-Aldrich</td><td>G0625</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G7021</td><td></td></tr><tr><td>MOPS</td><td>Sigma-Aldrich</td><td>M1254</td><td></td></tr><tr><td>Phenylmethylsulfonyl fluoride (PMSF)</td><td>Sigma-Aldrich</td><td>P7626</td><td>Used to inactivate proteinase K</td></tr><tr><td>Potassium phosphate dibasic</td><td>Sigma-Aldrich</td><td>P3786</td><td></td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma-Aldrich</td><td>P0662</td><td></td></tr><tr><td>Proteinase K</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S8501</td><td></td></tr><tr><td>Trichloroacetic acid (TCA)</td><td>Sigma-Aldrich</td><td>T6399</td><td></td></tr><tr><td>Trizma Base</td><td>Sigma-Aldrich</td><td>T1503</td><td></td></tr><tr><td>Zymolyase-20T from &lt;em&gt;Arthrobacter luteus&lt;/em&gt;</td><td>MP Biomedicals, Irvine, CA</td><td>320921</td><td>Used to lyse living yeast cell walls to produce spheroplast</td></tr></tbody></table>

assessment of submitochondrial protein localization in budding yeast saccharomyces cerevisiae

Despite recent advances in the characterization of yeast mitochondrial proteome, the submitochondrial localization of a significant number of proteins remains elusive. Here, we describe a robust and effective method for determining the suborganellar localization of yeast mitochondrial proteins, which is considered a fundamental step during mitochondrial protein function elucidation. This method involves an initial step that consists of obtaining highly pure intact mitochondria. These mitochondrial preparations are then subjected to a subfractionation protocol consisting of hypotonic shock (swelling) and incubation with proteinase K (protease). During swelling, the outer mitochondrial membrane is selectively disrupted, allowing the proteinase K to digest proteins of the intermembrane space compartment. In parallel, to obtain information about the topology of membrane proteins, the mitochondrial preparations are initially sonicated, and then subjected to alkaline extraction with sodium carbonate. Finally, after centrifugation, the pellet and supernatant fractions from these different treatments are analyzed by SDS-PAGE and western blot. The submitochondrial localization as well as the membrane topology of the protein of interest is obtained by comparing its western blot profile with known standards.

The success of submitochondrial fractionation protocol depends on obtaining highly purified intact mitochondria. For this, it is essential that during the yeast cell lysis, the intactness of the organelles remains almost totally preserved. This is achieved by using a cell lysis protocol that combines the enzymatic digestion of the cell wall followed by physical disruption of the plasma membrane by using a Dounce homogenizer. The mitochondrial contents are then collected by differential centrifugation. This subcellular fr...

Watch this Scientific Journal Video about Assessment of Submitochondrial Protein Localization in Budding Yeast Saccharomyces cerevisiae at JoVE.com

Assessment of Submitochondrial Protein Localization in Budding Yeast Saccharomyces cerevisiae

Despite recent advances, many yeast mitochondrial proteins still remain with their functions completely unknown. This protocol provides a simple and reliable method to determine the submitochondrial localization of proteins, which has been fundamental for the elucidation of their molecular functions.

Assessment of Submitochondrial Protein Localization in Budding Yeast Saccharomyces cerevisiae

Despite recent advances in the characterization of yeast mitochondrial proteome, the submitochondrial localization of a significant number of proteins ...

assessment-submitochondrial-protein-localization-budding-yeast

Universidad San Sebastian

Research

JoVE Journal

Biochemistry

2.8K Views.  Universidade de São Paulo. Despite recent advances, many yeast mitochondrial proteins still remain with their functions completely unknown. This protocol provides a simple and reliable method to determine the submitochondrial localization of proteins, which has been fundamental for the elucidation of their molecular functions.

Video: Assessment of Submitochondrial Protein Localization in Budding Yeast Saccharomyces cerevisiae

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

Biology

Identification of protein complexes with quantitative proteomics in S. cerevisiae

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as important intracellular signalling molecules. However, unlike proteins, lipids are small hydrophobic molecules that traffic primarily by poorly described nonvesicular routes, which are hypothesized to occur at membrane contact sites (MCSs). MCSs are regions where the endoplasmic reticulum (ER) makes direct physical contact with a partnering organelle, e.g., plasma membrane (PM). The ER portion of ER-PM MCSs is enriched in lipid-synthesizing enzymes, suggesting that lipid synthesis is directed to these sites and implying that MCSs are important for lipid traffic. Yeast is an ideal model to study ER-PM MCSs because of their abundance, with over 1000 contacts per cell, and their conserved nature in all eukaryotes. Uncovering the proteins that constitute MCSs is critical to understanding how lipids traffic is accomplished in cells, and how they act as signaling molecules. We have found that an ER called Scs2p localize to ER-PM MCSs and is important for their formation. We are focused on uncovering the molecular partners of Scs2p. Identification of protein complexes traditionally relies on first resolving purified protein samples by gel electrophoresis, followed by in-gel digestion of protein bands and analysis of peptides by mass spectrometry. This often limits the study to a small subset of proteins. Also, protein complexes are exposed to denaturing or non-physiological conditions during the procedure. To circumvent these problems, we have implemented a large-scale quantitative proteomics technique to extract unbiased and quantified data. We use stable isotope labeling with amino acids in cell culture (SILAC) to incorporate staple isotope nuclei in proteins in an untagged control strain. Equal volumes of tagged culture and untagged, SILAC-labeled culture are mixed together and lysed by grinding in liquid nitrogen. We then carry out an affinity purification procedure to pull down protein complexes. Finally, we precipitate the protein sample, which is ready for analysis by high-performance liquid chromatography/ tandem mass spectrometry. Most importantly, proteins in the control strain are labeled by the heavy isotope and will produce a mass/ charge shift that can be quantified against the unlabeled proteins in the bait strain. Therefore, contaminants, or unspecific binding can be easily eliminated. By using this approach, we have identified several novel proteins that localize to ER-PM MCSs. Here we present a detailed description of our approach. 

Here we describe a new quantitative proteomics technique for identifying protein complexes in Saccharomyces cerevisiae. In this study, we have used the SILAC method together with affinity purification followed by tandem mass spectrometry to identify with high specificity the binding partners of an ER protein, Scs2p.

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as ...

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A mitochondrial genome of baker&#8217;s yeast encodes eight proteins: three subunits of the cytochrome c oxidase (Cox1p, Cox2p, and Cox3p), three subunits of the ATP synthase (Atp6p, Atp8p, and Atp9p), a subunit of the ubiquinol-cytochrome c oxidoreductase enzyme, cytochrome b (Cytb), and mitochondrial ribosomal protein Var1p. The purpose of the method described here is to specifically label these proteins with 35S methionine, separate them by electrophoresis and visualize the signals as discrete bands on the screen. The procedure involves several steps. First, yeast cells are cultured in a galactose-containing medium until they reach the late logarithmic growth stage. Next, cycloheximide treatment blocks cytoplasmic translation and allows 35S methionine incorporation only in mitochondrial translation products. Then, all proteins are extracted from yeast cells and separated by polyacrylamide gel electrophoresis. Finally, the gel is dried and incubated with the storage phosphor screen. The screen is scanned on a phosphorimager revealing the bands. The method can be applied to compare the biosynthesis rate of a single polypeptide in the mitochondria of a mutant yeast strain versus the wild type, which is useful for studying mitochondrial gene expression defects. This protocol gives valuable information about the translation rate of all yeast mitochondrial mRNAs. However, it requires several controls and additional experiments to make proper conclusions.

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Baker&#8217;s yeast mitochondrial genome encodes eight polypeptides. The goal of the current protocol is to label all of them and subsequently visualize them as separate bands.

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A ...

Isolation of Mitochondrial Contact Sites

An Improved Method to Isolate Mitochondrial Contact Sites

Mitochondria are present in virtually all eukaryotic cells and perform essential functions that go far beyond energy production, for instance, the synthesis of iron-sulfur clusters, lipids, or proteins, Ca2+ buffering, and the induction of apoptosis. Likewise, mitochondrial dysfunction results in severe human diseases such as cancer, diabetes, and neurodegeneration. In order to perform these functions, mitochondria have to communicate with the rest of the cell across their envelope, which consists of two membranes. Therefore, these two membranes have to interact constantly. Proteinaceous contact sites between the mitochondrial inner and outer membranes are essential in this respect. So far, several contact sites have been identified. In the method described here, Saccharomyces cerevisiae mitochondria are used to isolate contact sites and, thus, identify candidates that qualify for contact site proteins. We used this method to identify the mitochondrial contact site and cristae organizing system (MICOS) complex, one of the major contact site-forming complexes in the mitochondrial inner membrane, which is conserved from yeast to humans. Recently, we further improved this method to identify a novel contact site consisting of Cqd1 and the Por1-Om14 complex.

Author Spotlight: Unveiling Mitochondrial Contact Sites and Architectural Insights 

Mitochondrial contact sites are protein complexes that interact with mitochondrial inner and outer membrane proteins. These sites are essential for the communication between the mitochondrial membranes and, thus, between the cytosol and the mitochondrial matrix. Here, we describe a method to identify candidates qualifying for this specific class of proteins.

Mitochondria are present in virtually all eukaryotic cells and perform essential functions that go far beyond energy production, for instance, the ...

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 ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Isolation of mRNAs Associated with Yeast Mitochondria to Study Mechanisms of Localized Translation

Most of mitochondrial proteins are encoded in the nucleus and need to be imported into the organelle. Import may occur while the protein is synthesized near the mitochondria. Support for this possibility is derived from recent studies, in which many mRNAs encoding mitochondrial proteins were shown to be localized to the mitochondria vicinity. Together with earlier demonstrations of ribosomes&#8217; association with the outer membrane, these results suggest a localized translation process. Such localized translation may improve import efficiency, provide unique regulation sites and minimize cases of ectopic expression. Diverse methods have been used to characterize the factors and elements that mediate localized translation. Standard among these is subcellular fractionation by differential centrifugation. This protocol has the advantage of isolation of mRNAs, ribosomes and proteins in a single procedure. These can then be characterized by various molecular and biochemical methods. Furthermore, transcriptomics and proteomics methods can be applied to the resulting material, thereby allow genome-wide insights. The utilization of yeast as a model organism for such studies has the advantages of speed, costs and simplicity. Furthermore, the advanced genetic tools and available deletion strains facilitate verification of candidate factors.

Many mRNAs encoding mitochondrial proteins are associated with the mitochondria outer membrane. We describe a subcellular fractionation procedure aimed at isolation of yeast mitochondria with its associated mRNAs and ribosomes. This protocol can be applied to cells grown under diverse conditions&#160;in order to reveal mechanisms of mRNA localization and localized translation near the mitochondria.

Most of mitochondrial proteins are encoded in the nucleus and need to be imported into the organelle. Import may occur while the protein is synthesized ...

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans is an established model for understanding the genomic regulation of biological processes of interest. This worm&#8217;s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo in any other organism.

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Skeletal muscle is essential for locomotion and is the bodies&#8217; main protein store. Muscle health measurements within C. elegans are described. Prospective changes to muscle structure and function are assessed using localized GFP and cationic dyes.

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age ...

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.

Regulated protein degradation is crucial for virtually every cellular function. Much of what is known about the molecular mechanisms and genetic ...

Protein Purification Technique that Allows Detection of Sumoylation and Ubiquitination of Budding Yeast Kinetochore Proteins Ndc10 and Ndc80

Post-translational Modifications (PTMs), such as phosphorylation, methylation, acetylation, ubiquitination, and sumoylation, regulate the cellular function of many proteins. PTMs of kinetochore proteins that associate with centromeric DNA mediate faithful chromosome segregation to maintain genome stability. Biochemical approaches such as mass spectrometry and western blot analysis are most commonly used for identification of PTMs. Here, a protein purification method is described that allows the detection of both sumoylation and ubiquitination of the kinetochore proteins, Ndc10 and Ndc80, in Saccharomyces cerevisiae. A strain that expresses polyhistidine-Flag-tagged Smt3 (HF-Smt3) and Myc-tagged Ndc10 or Ndc80 was constructed and used for our studies. For detection of sumoylation, we devised a protocol to affinity purify His-tagged sumoylated proteins by using nickel beads and used western blot analysis with anti-Myc antibody to detect sumoylated Ndc10 and Ndc80. For detection of ubiquitination, we devised a protocol for immunoprecipitation of Myc-tagged proteins and used western blot analysis with anti-Ub antibody to show that Ndc10 and Ndc80 are ubiquitinated. Our results show that epitope tagged-protein of interest in the His-Flag tagged Smt3 strain facilitates the detection of multiple PTMs. Future studies should allow exploitation of this technique to identify and characterize protein interactions that are dependent on a specific PTM.

This manuscript describes the detection of sumoylation and ubiquitination of kinetochore proteins, Ndc10 and Ndc80, in the budding yeast Saccharomyces cerevisiae.

Assessment of Submitochondrial Protein Localization in Budding Yeast Saccharomyces cerevisiae

Summary

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Chapters in this video

A high-throughput method to globally study the organelle morphology in S. cerevisiae

Identification of protein complexes with quantitative proteomics in S. cerevisiae

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

An Improved Method to Isolate Mitochondrial Contact Sites

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

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

Isolation of mRNAs Associated with Yeast Mitochondria to Study Mechanisms of Localized Translation

Methods to Assess Subcellular Compartments of Muscle in C. elegans

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

Protein Purification Technique that Allows Detection of Sumoylation and Ubiquitination of Budding Yeast Kinetochore Proteins Ndc10 and Ndc80