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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 Arthrobacter luteus</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

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

Genome-wide Quantification of Translation in Budding Yeast by Ribosome Profiling

Translation of mRNA into proteins is a complex process involving several layers of regulation. It is often assumed that changes in mRNA transcription reflect changes in protein synthesis, but many exceptions have been observed. Recently, a technique called ribosome profiling (or Ribo-Seq) has emerged as a powerful method that allows identification, with high accuracy, which regions of mRNA are translated into proteins and quantification of translation at the genome-wide level. Here, we present a generalized protocol for genome-wide quantification of translation using Ribo-Seq in budding yeast. In addition, combining Ribo-Seq data with mRNA abundance measurements allows us to simultaneously quantify translation efficiency of thousands of mRNA transcripts in the same sample and compare changes in these parameters in response to experimental manipulations or in different physiological states. We describe a detailed protocol for generation of ribosome footprints using nuclease digestion, isolation of intact ribosome-footprint complexes via sucrose gradient fractionation, and preparation of DNA libraries for deep sequencing along with appropriate quality controls necessary to ensure accurate analysis of in vivo translation.

Translational regulation plays an important role in the control of protein abundance. Here, we describe a high-throughput method for quantitative analysis of translation in the budding yeast Saccharomyces cerevisiae.

Translation of mRNA into proteins is a complex process involving several layers of regulation. It is often assumed that changes in mRNA transcription ...

Evaluation of the Impact of Protein Aggregation on Cellular Oxidative Stress in Yeast

Protein misfolding and aggregation into amyloid conformations have been related to the onset and progression of several neurodegenerative diseases. However, there is still little information about how insoluble protein aggregates exert their toxic effects in vivo. Simple prokaryotic and eukaryotic model organisms, such as bacteria and yeast, have contributed significantly to our present understanding of the mechanisms behind the intracellular amyloid formation, aggregates propagation, and toxicity. In this protocol, the use of yeast is described as a model to dissect the relationship between the formation of protein aggregates and their impact on cellular oxidative stress. The method combines the detection of the intracellular soluble/aggregated state of an amyloidogenic protein with the quantification of the cellular oxidative damage resulting from its expression using flow cytometry (FC). This approach is simple, fast, and quantitative. The study illustrates the technique by correlating the cellular oxidative stress caused by a large set of amyloid-&#946; peptide variants with their respective intrinsic aggregation propensities.

Protein aggregation elicits cellular oxidative stress. This protocol describes a method for monitoring the intracellular states of amyloidogenic proteins and the oxidative stress associated with them, using flow cytometry. The approach is used to study the behavior of soluble and aggregation-prone variants of the amyloid-&#946; peptide.

Protein misfolding and aggregation into amyloid conformations have been related to the onset and progression of several neurodegenerative diseases. ...

A Yeast 2-Hybrid Screen in Batch to Compare Protein Interactions

Screening for protein-protein interactions using the yeast 2-hybrid assay has long been an effective tool, but its use has largely been limited to the discovery of high-affinity interactors that are highly enriched in the library of interacting candidates. In a traditional format, the yeast 2-hybrid assay can yield too many colonies to analyze when conducted at low stringency where low affinity interactors might be found. Moreover, without a comprehensive and complete interrogation of the same library against different bait plasmids, a comparative analysis cannot be achieved. Although some of these problems can be addressed using arrayed prey libraries, the cost and infrastructure required to operate such screens can be prohibitive. As an alternative, we have adapted the yeast 2-hybrid assay to simultaneously uncover dozens of transient and static protein interactions within a single screen utilizing a strategy termed DEEPN (Dynamic Enrichment for Evaluation of Protein Networks), which incorporates high-throughput DNA sequencing and computation to follow the evolution of a population of plasmids that encode interacting partners. Here, we describe customized reagents and protocols that allow a DEEPN screen to be executed easily and cost-effectively.

Batch processing of yeast 2-hybrid screens allows for direct comparison of the interaction profiles of multiple bait proteins with a highly complex set of prey fusion proteins. Here, we describe refined methods, new reagents, and how to implement their use for such screens.

Screening for protein-protein interactions using the yeast 2-hybrid assay has long been an effective tool, but its use has largely been limited to the ...

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of biological membranes, energy storage and production, cellular signaling, vesicular transport of proteins, organelle biogenesis, and regulated cell death. Because the budding yeast Saccharomyces cerevisiae is a unicellular eukaryote amenable to thorough molecular analyses, its use as a model organism helped uncover mechanisms linking lipid metabolism and intracellular transport to complex biological processes within eukaryotic cells. The availability of a versatile analytical method for the robust, sensitive, and accurate quantitative assessment of major classes of lipids within a yeast cell is crucial for getting deep insights into these mechanisms. Here we present a protocol to use liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of major cellular lipids of S. cerevisiae. The LC-MS/MS method described is versatile and robust. It enables the identification and quantification of numerous species (including different isobaric or isomeric forms) within each of the 10 lipid classes. This method is sensitive and allows identification and quantitation of some lipid species at concentrations as low as 0.2 pmol/&#181;L. The method has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. The use of alternative mobile phase additives for electrospray ionization mass spectrometry in this method can increase the efficiency of ionization for some lipid species and can be therefore used to improve their identification and quantitation.

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol using liquid chromatography coupled with tandem mass spectrometry to identify and quantify major cellular lipids in Saccharomyces cerevisiae. The described method for a quantitative assessment of major lipid classes within a yeast cell is versatile, robust, and sensitive.

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of ...

Utilization of Grafix for the Detection of Transient Interactors of Saccharomyces cerevisiae Spliceosome Subcomplexes

Pre-mRNA splicing is a very dynamic process that involves many molecular rearrangements of the spliceosome subcomplexes during assembly, RNA processing, and release of the complex components. Glycerol gradient centrifugation has been used for the separation of protein or RNP (RiboNucleoProtein) complexes for functional and structural studies. Here, we describe the utilization of Grafix (Gradient Fixation), which was first developed to purify and stabilize macromolecular complexes for single particle cryo-electron microscopy, to identify interactions between splicing factors that bind transiently to the spliceosome complex. This method is based on the centrifugation of samples into an increasing concentration of a fixation reagent to stabilize complexes. After centrifugation of yeast total extracts loaded on glycerol gradients, recovered fractions are analyzed by dot blot for the identification of the spliceosome sub-complexes and determination of the presence of individual splicing factors.

Utilization of Grafix for the Detection of Transient Interactors of Saccharomyces cerevisiae Spliceosome Subcomplexes

Here, we describe the utilization of Grafix (Gradient Fixation), a glycerol gradient centrifugation in the presence of a crosslinker, to identify interactions between splicing factors that bind transiently to the spliceosome complex.

Pre-mRNA splicing is a very dynamic process that involves many molecular rearrangements of the spliceosome subcomplexes during assembly, RNA processing, ...

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 ...

Quantitative Metabolomics of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Metabolomics is a methodology used for the identification and quantification of many low-molecular-weight intermediates and products of metabolism within a cell, tissue, organ, biological fluid, or organism. Metabolomics traditionally focuses on water-soluble metabolites. The water-soluble metabolome is the final product of a complex cellular network that integrates various genomic, epigenomic, transcriptomic, proteomic, and environmental factors. Hence, the metabolomic analysis directly assesses the outcome of the action for all these factors in a plethora of biological processes within various organisms. One of these organisms is the budding yeast Saccharomyces cerevisiae, a unicellular eukaryote with the fully sequenced genome. Because S. cerevisiae is amenable to comprehensive molecular analyses, it is used as a model for dissecting mechanisms underlying many biological processes within the eukaryotic cell. A versatile analytical method for the robust, sensitive, and accurate quantitative assessment of the water-soluble metabolome would provide the essential methodology for dissecting these mechanisms. Here we present a protocol for the optimized conditions of metabolic activity quenching in and water-soluble metabolite extraction from S. cerevisiae cells. The protocol also describes the use of liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of the extracted water-soluble metabolites. The LC-MS/MS method of non-targeted metabolomics described here is versatile and robust. It enables the identification and quantification of more than 370 water-soluble metabolites with diverse structural, physical, and chemical properties, including different structural isomers and stereoisomeric forms of these metabolites. These metabolites include various energy carrier molecules, nucleotides, amino acids, monosaccharides, intermediates of glycolysis, and tricarboxylic cycle intermediates. The LC-MS/MS method of non-targeted metabolomics is sensitive and allows the identification and quantitation of some water-soluble metabolites at concentrations as low as 0.05 pmol/&#181;L. The method has been successfully used for assessing water-soluble metabolomes of wild-type and mutant yeast cells cultured under different conditions.

Quantitative Metabolomics of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol for the identification and quantitation of major classes of water-soluble metabolites in the yeast Saccharomyces cerevisiae. The described method is versatile, robust, and sensitive. It allows the separation of structural isomers and stereoisomeric forms of water-soluble metabolites from each other.

Metabolomics is a methodology used for the identification and quantification of many low-molecular-weight intermediates and products of metabolism within ...

Analysis of Neutral Lipid Synthesis in Saccharomyces cerevisiae by Metabolic Labeling and Thin Layer Chromatography

Neutral lipids (NLs) are a class of hydrophobic, chargeless biomolecules that play key roles in energy and lipid homeostasis. NLs are synthesized de novo&#160;from acetyl-CoA and are primarily present in eukaryotes in the form of triglycerides (TGs) and sterol-esters (SEs). The enzymes responsible for the synthesis of NLs are highly conserved from Saccharomyces cerevisiae (yeast) to humans, making yeast a useful model organism to dissect the function and regulation of NL metabolism enzymes. While much is known about how acetyl-CoA is converted into a diverse set of NL species, mechanisms for regulating NL metabolism enzymes, and how mis-regulation can contribute to cellular pathologies, are still being discovered. Numerous methods for the isolation and characterization of NL species have been developed and used over decades of research; however, a quantitative and simple protocol for the comprehensive characterization of major NL species has not been discussed. Here, a simple and adaptable method to quantify the de novo&#160;synthesis of major NL species in yeast is presented. We apply 14C-acetic acid metabolic labeling coupled with thin layer chromatography to separate and quantify a diverse range of physiologically important NLs. Additionally, this method can be easily applied to study in vivo reaction rates of NL enzymes or degradation of NL species over time.

Analysis of Neutral Lipid Synthesis in Saccharomyces cerevisiae by Metabolic Labeling and Thin Layer Chromatography

Here, a protocol is presented for the metabolic labeling of yeast with 14C-acetic acid, which is coupled with thin layer chromatography for the separation of neutral lipids.

Neutral lipids (NLs) are a class of hydrophobic, chargeless biomolecules that play key roles in energy and lipid homeostasis. NLs are synthesized de ...

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in situ. Owing to the strong interaction of an electron with the matter, high-resolution cryo-ET studies are limited to specimens with a thickness of less than 200 nm, which restricts the applicability of cryo-ET only to the peripheral regions of a cell. A complex workflow that comprises the preparation of thin cellular cross-sections by cryo-focused ion beam micromachining (cryo-FIBM) was introduced during the last decade to enable the acquisition of cryo-ET data from the interior of larger cells. We present a protocol for the preparation of cellular lamellae from a sample vitrified by plunge freezing utilizing Saccharomyces cerevisiae as a prototypical example of a eukaryotic cell with wide utilization in cellular and molecular biology research. We describe protocols for vitrification of S. cerevisiae into isolated patches of a few cells or a continuous monolayer of the cells on a TEM grid and provide a protocol for lamella preparation by cryo-FIB for these two samples.

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

We present a protocol for lamella preparation of plunge frozen biological specimens by cryo-focused ion beam micromachining for high-resolution structural studies of macromolecules in situ with cryo-electron tomography. The presented protocol provides guidelines for the preparation of high-quality lamellae with high reproducibility for structural characterization of macromolecules inside the&#160;Saccharomyces cerevisiae.

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in ...

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.