We thank Laurie Destroismaisons and Sarah Peyrard for outstanding technical support and Dr.Alexandre Prat for access to the flow cytometer. We would also like to acknowledge Dr. Timothy Miller for his contribution regarding the removal of myelin from the preparations. This work was supported by the Canadian Institutes of Health Research (CIHR) Neuromuscular Research Partnership, Canadian Foundation for Innovation, ALS Society of Canada, the Frick Foundation for ALS Research, CHUM Foundation and Fonds de la Recherche en Sant&#233; du Qu&#233;bec (C.V.V.). Both C.V.V. and N.A. are Research Scholars of the Fonds de la Recherche en Sant&#233; du Qu&#233;bec and CIHR New Investigators. S.P. is supported by the Tim No&#235;l Studentship from the ALS Society of Canada.

Animals used in this study were treated in strict accordance to a protocol (N08001CVsr) approved by the Centre de Recherche du Centre Hospitalier de l&#8217;Universit&#233; de Montr&#233;al (CRCHUM) Institutional Committee for the Protection of Animals which follows national standards as outlined by the Canadian Council on Animal Care (CCAC).
Prepare all reagents required to perform this protocol (Table 1). All other details regarding equipment, supplies and suppliers can be found in List of Materials.
<img alt="Table 1" fo:content-width="5in" src="/files/ftp_upload/51887/51887table1.jpg" width="500" /> 
Table 1. Buffer Compositions.
1. Collection of Rat Spinal Cord
<ol>
	<li>Deeply anaesthetize the rat (Sprague Dawley) with 4% isoflorane. Verify anaesthetization by a lack of reflex upon pinching of the forepaw. Euthanize the rat by decapitation via guillotine. This method of euthanasia is preferred over others, which might distort the spinal cord.</li>
	<li>Cut the skin of the back to expose the spine. Cut the spinal column with bone scissors just above the pelvic bone. Visualize the opening of the vertebral column.</li>
	<li>Insert a 10 ml syringe, with a 200 &#181;l pipette tip (attached via melting slightly over flame), filled with Phosphate Buffered Saline (PBS), into the vertebral column.</li>
	<li>Flush out the spinal cord by applying a medium amount of pressure to the plunger.</li>
	<li>If any blood is present on the spinal cord, rinse with PBS before proceeding to the next step. 
	NOTE: This method has also been validated for brain and liver. If including these tissues, collect half the brain from the euthanized rat and a piece of liver equal in weight to the spinal cord. All other steps remain identical.</li>
</ol>
2. Isolation of Spinal Cord Mitochondria (Adapted from Vande Velde et al. 4)
<ol>
	<li>Collect the whole intact spinal cord and place in 5 ml glass homogenizer with 5 volumes (~3.25 ml) Homogenization Buffer (HB). For optimal recovery of isolated mitochondria, perform all steps on ice or in cold room. Homogenize tissue by hand until no large pieces of tissue remain, approximately eight strokes. Place homogenate in two (2 ml) or three (1.7 ml) microcentrifuge tubes. Centrifuge 1,300 x g for 10 min at 4 &#176;C in a benchtop microcentrifuge.</li>
	<li>Recover supernatant and place in a 5 ml ultracentrifuge tube. Add 750 &#181;l (~0.5 volumes) HB to the pellet-containing microcentrifuge tube and gently resuspend the pellet. Repeat centrifugation and resuspension steps two more times. Pool all supernatants (S1a and S1b) into the same 5 ml ultracentrifuge tube above. This step serves to remove small debris.</li>
	<li>Centrifuge pooled S1 using an ultracentrifuge equipped with a swinging bucket rotor and centrifuge at 17,000 x g for 15 min at 4 &#176;C.</li>
	<li>Keep supernatant (S2) for further processing if the cytosolic fraction is of interest (for example for Western blot analysis). Resuspend the pellet (P2), the crude mitochondrial fraction, in 4 ml HB+50 mM KCl. Centrifuge 17,000 x g for 15 min at 4 &#176;C in a swinging bucket rotor. Discard the supernatant and gently resuspend the pellet (P3) in 800 &#181;l HB. 
	NOTE: Mitochondria are washed with HB+50 mM KCl to remove any non-specific mitochondrially-associated contaminants.</li>
	<li>In a new 5 ml ultracentrifuge tube, add exactly 800 &#181;l of resuspended pellet (P3).</li>
	<li>To this tube add, 200 &#181;l of Iodixanol (density gradient medium), thereby creating a final concentration of 12% Iodixanol. Mix the contents of the tube gently, but thoroughly via pipetting with a P1000. Addition of Iodixanol first, and then resuspended pellet (P3) may be preferable to facilitate thorough mixing. Centrifuge in an ultracentrifuge equipped with a swinging bucket rotor at 17,000 x g for 15 min at 4 &#176;C.</li>
	<li>NOTE: Liver does not contain myelin and therefore this step is not necessary if only the liver is being processed. However, if liver is being processed concurrently with CNS mitochondria, it is recommended to treat all tissues equally.</li>
	<li>Aspirate the layer of myelin at the top of the tube and carefully remove and discard the supernatant. Pellet may be loose. Resuspend the pellet in 4 ml HB. Centrifuge again at 17,000 x g for 10 min at 4 &#176;C. Discard the supernatant and resuspend the pellet in 4 ml HB. Repeat the centrifugation and remove the supernatant.</li>
	<li>Resuspend the final pellet (P7) in 100-200 &#181;l HB and transfer to a 1.7 ml microcentrifuge tube. This sample contains isolated mitochondria.</li>
	<li>Proceed to protein quantification. Dilute samples and the standard curve in 2% sodium dodecyl sulfate (SDS) to ensure adequate solubilization of mitochondria during protein quantification.</li>
</ol>
3. Immunolabeling of Isolated Mitochondria for Flow Cytometry
<ol>
	<li>For each staining mix to be tested, pipette 25 &#181;g of isolated mitochondria into a 1.7 ml microcentrifuge tube. Include an unstained sample in each experiment; and for each antibody, be sure to include a sample for the appropriate isotype control.</li>
	<li>Centrifuge at 17,000 x g for 2 min at 4 &#176;C in a bench-top microcentrifuge.</li>
	<li>Remove supernatant and resuspend isolated mitochondria in 50 &#181;l Mitochondria Buffer (M Buffer) supplemented with 10% fatty-acid free BSA for 15 min at 4 &#176;C (blocking step). 
	NOTE: During labeling, perform incubations in a refrigerator at 4 &#176;C.</li>
	<li>Add primary antibody (rabbit anti-Mfn2, 20 &#181;g per ml) to tube and incubate for 30 min at 4 &#176;C. 
	NOTE: Determine optimal concentration of each antibody empirically by titration. Due to variability in concentration and/or purity, different lots of the same antibody from the same manufacturer may lead to different results; therefore titration is needed for each new lot of antibody.</li>
	<li>Wash out unbound primary antibody: Centrifuge at 17,000 x g for 2 min at 4 &#176;C. Remove the supernatant and gently resuspend the pellet in 200 &#181;l M Buffer. Centrifuge at 17,000 x g for 2 min at 4 &#176;C. Remove supernatant and resuspend the pellet in 50 &#181;l M Buffer.</li>
	<li>Add secondary antibody (Donkey anti-rabbit IgG Phycoerythrin (PE), 0.5 &#181;g per ml) to tube and incubate samples for 30 min at 4 &#176;C, protected from light.</li>
	<li>Wash out unbound secondary antibody: Centrifuge at 17,000 x g for 2 min at 4 &#176;C. Remove the supernatant and resuspend the pellet in 200 &#181;l M Buffer. Centrifuge at 17,000 x g for 2 min at 4 &#176;C. Remove supernatant and resuspend the pellet in 500 &#181;l M Buffer.</li>
	<li>To ensure events are in fact mitochondria, stain isolated mitochondria with a mitochondria specific fluorescent dye for 15 min at RT, protected from light. If staining of other functional parameters (mitochondrial transmembrane potential or superoxide production) is desired, proceed to step 4. If not, proceed to step 5 for acquisition. 
	NOTE: It is important to verify that the emission spectrum of the secondary antibody is compatible with that of the functional dyes. For example, if verifying mitochondrial purity with a commercial dye with spectral properties similar to FITC and transmembrane potential with Tetramethylrhodamine methyl ester (TMRM), a viable secondary antibody would be allophycocyanin (APC: Ex 650 nm/Em 660 nm). Add compensations controls, i.e., a sample immunolabeled or stained with a single fluorophore, when applicable.</li>
	<li>Transfer to a tube suitable for loading flow cytometer. (To facilitate the small sample size, a microtiter tube is placed inside 3 ml flow cytometer tube.) Keep samples on ice and proceed immediately to flow cytometer for acquisition.</li>
</ol>
4. Assaying Mitochondrial Transmembrane Potential and Mitochondrial Superoxide Production by Flow Cytometry
<ol>
	<li>Verify that isolated mitochondria have an intact transmembrane potential by staining with 100 nM TMRM (Ex 548 nm/Em 574 nm)5, at step 3.8, for 15 min at RT, protected from light. For comparison of transmembrane potential between samples and populations, use of lower/non-quenching concentrations of TMRM (1 to 30 nM), may be more appropriate6.
	<ol>
		<li>As a control for TMRM staining, stain isolated mitochondria with 100 nM TMRM in the presence of 100 &#181;M carbonyl cyanide m-chloro phenyl hydrazine (CCCP), a mitochondrial uncoupler that will depolarize mitochondria. The concentration of CCCP required to depolarize the mitochondria may be less if lower concentrations of dye are used.</li>
	</ol>
	</li>
	<li>Verify that isolated mitochondria produce mitochondrial superoxide by staining with an appropriate mitochondrial superoxide indicator7, also at step 3.8, for 15 min at RT, protected from light.
	<ol>
		<li>As a control for mitochondrial superoxide production, stain isolated mitochondria with dye in the presence of 10 &#181;M Antimycin A, an inhibitor of complex III of the respiratory chain that will augment mitochondrial superoxide production.</li>
	</ol>
	</li>
</ol>
5. Acquisition and Analysis of Immunolabeled Isolated Mitochondria by Flow Cytometry
<ol>
	<li>Instrument set up: Switch voltages from linear to log mode to facilitate analysis of isolated mitochondria and set voltages (FSC: 450; SSC: 250). Ensure that events are collected in FSC-A (area) mode as well as FSC-W (width) and FSC-H (height), to be able to exclude doublets (i.e., two events, passing through the detector at the same time) in analysis post-data collection. Set the number of events to be collected to 100,000. Acquire compensation controls, if applicable.</li>
	<li>Data acquisition: Before data acquisition, avoid vortexing samples. Instead mix by gently tapping tube. Initially collect events at a low pressure, during gating. Gate on total population. Adjust voltages of histograms accordingly, usually the peak of the unstained sample will correspond to the second decade (102). Once gates are established, and samples are being processed, the pressure can be switched to high.</li>
	<li>Analysis: Visualize doublets by plotting FSC-W versus FSC. Identify singlets and doublets. Gate on singlets. Select the mitochondrial population by gating on events that are stained positively with a mitochondrial-selective dye.</li>
	<li>Determine background labeling from isotype control. Using the isotype control sample, determine the percentage of the mitochondrial population labeling positive for Mfn2 antibody.</li>
</ol>

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The authors have nothing to disclose.

It is increasingly evident that mitochondria are key players in both normal physiology and disease. While immunoblotting can determine which proteins are found within mitochondria or at the mitochondrial surface in a certain condition, this method reports on the average of the entire population. This method cannot yield information about relative abundances of mitochondrial subpopulations or subsets. While it has been previously assumed that all mitochondria are created equally, the field is increasingly recognizing that...

Mitochondria are highly dynamic organelles that undergo multiple rounds of fission and fusion, are transported to sites of high energy demand and respond rapidly to physiological stimuli1. Since it is increasingly recognized that mitochondria within different tissues, even different cellular compartments, have distinct functional profiles, new methods are needed to identify these distinct mitochondrial subsets.
Microscopy provides a means whereby individual mitochondria can be visualized and the presence of a protein at or in mitochondria can be determined by immunofluorescence2. However, quantitative analysis by this method is labor intensive and is more suitable for experiments using immortalized or primary cell lines. The study of individual mitochondria derived from tissue is significantly more difficult and most methods do not allow for easy identification of mitochondrial subsets concurrently with the evaluation of mitochondrial function3.
In order to address this hurdle, a novel method to immunolabel mitochondria isolated from rodent tissues and subsequently analyzed by flow cytometry has been developed. This allows for the rapid detection and quantification of proteins localized to the mitochondrial outer membrane, which compared to analysis by microscopy, is much less labor intensive and permits the analysis of thousands of mitochondria in a single sample. This assay can be applied to monitor the fate and relative amount of mitochondrial outer membrane proteins that are thought to be constitutively present at the mitochondria, the recruitment of proteins to the mitochondrial surface, or the detection of proteins mislocalized to the mitochondria in pathological conditions. Moreover, the incorporation of conventional fluorescent indicator dyes permits the simultaneous evaluation of certain aspects of mitochondrial function in distinct mitochondrial subpopulations.

<table><tbody><tr><td>Rats</td><td>Charles River</td><td>Strain code 400</td><td>Adult (9 weeks to 18 weeks) male or female rats can be used for the isolation protocol. Weight of rats is dependent on gender and age (males between 300 to 500 g and females between 200 to 350 g) are typically used.</td></tr><tr><td>Dounce homogenizer</td><td>Kontes Glass Co.&amp;nbsp;</td><td>885450-0022</td><td>Duall 22</td></tr><tr><td>Microcentrifuge</td><td>Thermo Scientific</td><td></td><td>Sorvall Legend Micro 17 R</td></tr><tr><td>Ultra-Clear Ultracentrifuge tubes</td><td>Beckman Coulter</td><td>344057</td><td>Transparent, thin walled&amp;nbsp;</td></tr><tr><td>Sorvall Ultracentrifuge</td><td>Thermo Scientific</td><td></td><td>Sorvall WX UltraSeries</td></tr><tr><td>AH-650 rotor and buckets&amp;nbsp;&amp;nbsp;</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Opti-prep</td><td>Axis-Shield</td><td>1114542</td><td>Iodixanol, density gradient medium</td></tr><tr><td>Fatty acid free Bovine Serum Albumin</td><td>Sigma</td><td>A8806</td><td>Must be fatty acid free for mitochondria</td></tr><tr><td>Sodium succinate dibasic hexahydrate&amp;nbsp;&amp;nbsp;</td><td>Sigma</td><td>S9637</td><td></td></tr><tr><td>Rabbit anti-Mitofusin2 antibody</td><td>Sigma</td><td>M6319</td><td></td></tr><tr><td>Rabbit IgG</td><td>Jackson Immuno Research&amp;nbsp;</td><td>011-000-003</td><td></td></tr><tr><td>Anti-Rabbit IgG PE</td><td>eBioscience</td><td>12-4739-81</td><td></td></tr><tr><td>Micro titer tube</td><td>Bio-Rad</td><td>223-9391</td><td>For sample acquisition by flow cytometry&amp;nbsp;</td></tr><tr><td>MitoTrackerGreen (MTG)</td><td>Invitrogen</td><td>M7514</td><td>100 nM: Ex 490 nm/Em 516 nm</td></tr><tr><td>TMRM</td><td>Invitrogen</td><td>T668</td><td>100 nM: Ex 548 nm/Em 574 nm</td></tr><tr><td>CCCP</td><td>Sigma</td><td>C2759</td><td></td></tr><tr><td>MitoSOX Red</td><td>Invitrogen</td><td>M36008</td><td>5 &amp;micro;M: Ex 540 nm/Em 600 nm</td></tr><tr><td>Antimycin A</td><td>Sigma</td><td>A8874</td><td></td></tr><tr><td>LSR II flow cytometer</td><td>BD</td><td></td><td></td></tr><tr><td>BD FACSDiva Software</td><td>BD</td><td></td><td></td></tr><tr><td>FlowJo</td><td>TreeStar Inc.&amp;nbsp;</td><td></td><td>Software used for analysis</td></tr><tr><td>BCA protein assay kit&amp;nbsp;&amp;nbsp;</td><td>Pierce/Thermo Scientific</td><td>23225</td><td>Bradford assay is not recomended as it is not compatible with high concentrations of SDS</td></tr></tbody></table>

immunodetection of outer membrane proteins by flow cytometry of isolated mitochondria

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and pathophysiology. This protocol describes a technique where mitochondria isolated from rodent tissue are immunolabeled and analyzed by flow cytometry. Mitochondria are isolated from rodent spinal cords and subjected to a rapid enrichment step so as to remove myelin, a major contaminant of mitochondrial fractions prepared from nervous tissue. Isolated mitochondria are then labeled with an antibody of choice and a fluorescently conjugated secondary antibody. Analysis by flow cytometry verifies the relative purity of mitochondrial preparations by staining with a mitochondrial specific dye, followed by detection and quantification of immunolabeled protein. This technique is rapid, quantifiable and high-throughput, allowing for the analysis of hundreds of thousands of mitochondria per sample. It is applicable to assess novel proteins at the mitochondrial surface under normal physiological conditions as well as the proteins that may become mislocalized to this organelle during pathology. Importantly, this method can be coupled to fluorescent indicator dyes to report on certain activities of mitochondrial subpopulations and is feasible for mitochondria from the central nervous system (brain and spinal cord) as well as liver.

Mitochondria derived from rat spinal cords can be immunolabeled with an antibody targeted to Mitofusin2 (Mfn2), a protein implicated in the fusion of the outer membrane of mitochondria8. Following isolation and labeling with a Mfn2 specific antibody and a fluorescently conjugated secondary antibody, mitochondria are processed by flow cytometry (Figure 1). Following data acquisition, samples are analyzed using flow cytometry analysis software, by first visualizing all collected events on a dot ...

Watch this Scientific Journal Video about Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria at JoVE.com

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and ...

immunodetection-outer-membrane-proteins-flow-cytometry-isolated

Research

JoVE Journal

Biology

15.4K Views.  Université de Montréal, CRCHUM. Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.

Video: Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Isolation and Respiratory Measurements of Mitochondria from Arabidopsis thaliana

Mitochondria are essential organelles involved in numerous metabolic pathways in plants, most notably the production of adenosine triphosphate (ATP) from the oxidation of reduced compounds such as nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FADH2). The complete annotation of the Arabidopsis thaliana genome has established it as the most widely used plant model system, and thus the need to purify mitochondria from a variety of organs (leaf, root, or flower) is necessary to fully utilize the tools that are now available for Arabidopsis to study mitochondrial biology. Mitochondria are isolated by homogenization of the tissue using a variety of approaches, followed by a series of differential centrifugation steps producing a crude mitochondrial pellet that is further purified using continuous colloidal density gradient centrifugation. The colloidal density material is subsequently removed by multiple centrifugation steps. Starting from 100 g of fresh leaf tissue, 2 - 3 mg of mitochondria can be routinely obtained. Respiratory experiments on these mitochondria display typical rates of 100 - 250 nmol O2 min-1 mg total mitochondrial protein-1 (NADH-dependent rate) with the ability to use various substrates and inhibitors to determine which substrates are being oxidized and the capacity of the alternative and cytochrome terminal oxidases. This protocol describes an isolation method of mitochondria from Arabidopsis thaliana leaves using continuous colloidal density gradients and an efficient respiratory measurements of purified plant mitochondria.

Isolation and Respiratory Measurements of Mitochondria from Arabidopsis thaliana

As mitochondria are only a small percentage of the plant cell, they need to be purified for a range of studies. Mitochondria can be isolated from a variety of plant organs by homogenization, followed by differential and density gradient centrifugation to obtain a highly purified mitochondrial fraction.

Mitochondria are essential organelles involved in numerous metabolic pathways in plants, most notably the production of adenosine triphosphate (ATP) from ...

Biochemistry

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 ...

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Immunology and Infection

A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment elevated myocardial infarction. However, reperfusion per se can result in further cardiomyocyte death, a phenomenon known as reperfusion injury. The opening of the mitochondrial permeability transition pore (mPTP), with the decrease of the mitochondrial membrane potential (MMP), or mitochondrial depolarization, is universally recognized as the final step of reperfusion injury and is responsible for mitochondrial and cardiomyocyte death. JC-1 is a lipophilic cationic dye that accumulates in mitochondria depending on the value of MMP. The higher the MMP is, the more JC-1 accumulates in the mitochondria. The increasing amounts of JC-1 in mitochondria can be reflected by a fluorescence emission shift from green (~530 nm) to red (~590 nm). Therefore, the reduction of the red/green fluorescence intensity ratio can indicate the depolarization of mitochondria. Here, we take advantage of JC-1 to measure MMP, or the opening of mPTP in human cardiac myocytes after hypoxia/reoxygenation, detected by flow cytometry.

Here, we present a protocol that uses JC-1 dye to assess the mitochondrial membrane potential of cells after being exposed to hypoxia/reoxygenation with or without a protective agent.

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment ...

Medicine

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

Application of Flow Cytometric Analysis for Measuring Multiple Mitochondrial Parameters in 3D Brain Organoids

Mitochondrial dysfunction is a common primary or secondary contributor to many types of neurodegeneration, and changes in mitochondrial mass, mitochondrial respiratory chain (MRC) complexes, and mitochondrial DNA (mtDNA) copy number often feature in these processes. Human brain organoids derived from human induced pluripotent stem cells (iPSCs) recapitulate the brain&#39;s three-dimensional (3D) cytoarchitectural arrangement and offer the possibility to study disease mechanisms and screen new therapeutics in a complex human system. Here, we report a unique flow cytometry-based approach to measure multiple mitochondrial parameters in iPSC-derived cortical organoids. This report details a protocol for generating cortical brain organoids from iPSCs, single-cell dissociation of generated organoids, fixation, staining, and subsequent flow cytometric analysis to assess multiple mitochondrial parameters. Double staining with antibodies against the MRC complex subunit NADH: Ubiquinone Oxidoreductase Subunit B10 (NDUFB10) or mitochondrial transcription factor A (TFAM) together with voltage-dependent anion-selective channel 1 (VDAC 1) permits assessment of the amount of these proteins per mitochondrion. Since the quantity of TFAM corresponds to the amount of mtDNA, it provides an indirect estimation of the number of mtDNA copies per mitochondrial content. This entire procedure can be completed within a span of 2-3 h. Crucially, it allows for the concurrent quantification of multiple mitochondrial parameters, including both total and specific levels relative to the mitochondrial mass.

This protocol reports a unique method of using a streaming cytometer and multiple antibodies for simultaneous assessment of multiple mitochondrial functional parameters, including changes in mitochondrial volume, amounts of the mitochondrial respiratory chain (MRC) complex subunits, and mitochondrial DNA (mtDNA) replication.

Mitochondrial dysfunction is a common primary or secondary contributor to many types of neurodegeneration, and changes in mitochondrial mass, ...

Neuroscience

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between defective mitophagy and various human diseases, including neurodegenerative diseases, cancer, and metabolic diseases. In addition, recent studies have shown that mitophagy is involved in normal cellular processes, such as differentiation and development. However, the precise role of and molecular mechanisms underlying mitophagy require further study. Therefore, it is critical to develop a robust and convenient method for measuring changes in mitophagy activity. Here, we describe a detailed protocol for quantitatively assessing mitophagy activity through flow cytometry using the mitochondria-targeted fluorescent protein Keima (mt-Keima). This flow cytometry assay can analyze mitophagy activity more rapidly and sensitively than conventional microscopy- or immunoblotting-based methods. This protocol can be applied to analyze mitophagy activity in various cell types.

Mitophagy, the selective degradation of mitochondria, has been implicated in mitochondrial homeostasis and is deregulated in various human diseases. However, convenient experimental methods for measuring mitophagy activity are very limited. Here, we provide a sensitive assay for measuring mitophagy activity using flow cytometry.

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between ...

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from low-sensitivity, indirect read-outs, limited throughput, and/or population-level rather than single-cell resolution. We designed a flow cytometry-based single methodology that addresses all of these limitations: Distributed Amphifluoric FRET or DAmFRET. DAmFRET detects and quantifies protein self-assemblies by sensitized emission FRET in vivo, enables deployment across model systems-from yeast to human cells-and achieves sensitive, single-cell, high-throughput read-outs irrespective of protein localization or solubility.

This article describes a FRET-based flow cytometry protocol to quantify protein self-assembly in both S. cerevisiae and HEK293T cells.

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from ...

Improving the Accuracy of Flow Cytometric Assessment of Mitochondrial Membrane Potential in Hematopoietic Stem and Progenitor Cells Through the Inhibition of Efflux Pumps

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic homeostasis have been extensively studied by HSC researchers. Mitochondrial activity levels are reflected in their membrane potentials (&#916;&#936;m), which can be measured by cell-permeant cationic dyes such as TMRM (tetramethylrhodamine, methyl ester). The ability of efflux pumps to extrude these dyes from cells can limit their usefulness, however. The resulting measurement bias is particularly critical when assessing HSCs, as xenobiotic transporters exhibit higher levels of expression and activity in HSCs than in differentiated cells. Here, we describe a protocol utilizing Verapamil, an efflux pump inhibitor, to accurately measure &#916;&#936;m across multiple bone marrow populations. The resulting inhibition of pump activity is shown to increase TMRM intensity in hematopoietic stem and progenitor cells (HSPCs), while leaving it relatively unchanged in mature fractions. This highlights the close attention to dye-efflux activity that is required when &#916;&#936;m-dependent dyes are used, and as written and visualized, this protocol can be used to accurately compare either different populations within the bone marrow, or the same population across different experimental models.

Xenobiotic efflux pumps are highly active in hematopoietic stem and progenitor cells (HSPCs) and cause extrusion of TMRM, a mitochondrial membrane potential fluorescent dye. Here, we present a protocol to accurately measure mitochondrial membrane potential in HSPCs by TMRM in the presence of Verapamil, an efflux pump inhibitor.

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic ...

Improved Protein Quantification Using Two-Step Mitochondrial Isolation

Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification

Most physiological and disease processes, from central metabolism to immune response to neurodegeneration, involve mitochondria. The mitochondrial proteome is composed of more than 1,000 proteins, and the abundance of each can vary dynamically in response to external stimuli or during disease progression. Here, we describe a protocol for isolating high-quality mitochondria from primary cells and tissues. The two-step procedure comprises (1) mechanical homogenization and differential centrifugation to isolate crude mitochondria, and (2) tag-free immune capture of mitochondria to isolate pure organelles and eliminate contaminants. Mitochondrial proteins from each purification stage are analyzed by quantitative mass spectrometry, and enrichment yields are calculated, allowing the discovery of novel mitochondrial proteins by subtractive proteomics. Our protocol provides a sensitive and comprehensive approach to studying mitochondrial content in cell lines, primary cells, and tissues.

Author Spotlight: Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification  

We present a two-step protocol for high-quality mitochondria isolation that is compatible with protein discovery and quantification at a proteome scale. Our protocol does not require genetic engineering and is thus suitable for studying mitochondria from any primary cells and tissues.

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

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Isolation and Respiratory Measurements of Mitochondria from Arabidopsis thaliana

An Improved Method to Isolate Mitochondrial Contact Sites

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Application of Flow Cytometric Analysis for Measuring Multiple Mitochondrial Parameters in 3D Brain Organoids

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Improving the Accuracy of Flow Cytometric Assessment of Mitochondrial Membrane Potential in Hematopoietic Stem and Progenitor Cells Through the Inhibition of Efflux Pumps

Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification