Name: a flow cytometry-based assay for measuring mitochondrial membrane potential in cardiac myocytes after hypoxia/reoxygenation
Uploaded: 2018-07-13T13:00:00.000+00:00
Duration: 7 min 14 s
Description: Watch this Scientific Journal Video about A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation at JoVE.com

This study was supported by grants from The National Key Research and Development Program of China (No. 2017YFC1700503), the National Basic Research Program (973 Program) of China (No.2012CB518602), the National Natural Science Foundation of China (No. 81370223 and No. 81573957), and the Postgraduates&#39; Innovative Research Foundation of Peking Union Medical College (2016-1002-01-02).

1. Preparation of Reagents and Solutions
<ol>
	<li>Prepare human cardiac myocytes (HCMs) complete medium by adding 25 mL of Myocyte Growth Medium supplement mix to 500 mL of Myocyte Growth Medium as per the manufacturer&#8217;s instructions. Store at 4 &#176;C and warm to 37 &#176;C before using.</li>
	<li>Prepare Tongxinluo (TXL) Solution by dissolving TXL ultrafine powder in serum/glucose-free Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) and adjust the concentration of TXL to 400 &#181;g/mL by adding DMEM (for more details, see11). Store at 4 &#176;C and warm to 37 &#176;C before using.</li>
	<li>Prepare 5 mL of JC-1 working solution by adding 25 &#181;L of JC-1 stock solution (200x), 4 mL of ultrapure water, and 1 mL of stock staining buffer (5x) in a 15 mL centrifuge tube in the dark. Pipette the mixture up and down several times and warm to 37 &#176;C before using.</li>
	<li>Prepare 30 mL of working staining buffer by adding 24 mL of ultrapure water to 6 mL of stock staining buffer (5x) in a 50 mL centrifuge tube. Use this solution ice-cold.</li>
</ol>
2. Establishment of Hypoxia/Reoxygenation Model
<ol>
	<li>Plate 1.0 x 105 cells per well with HCM complete medium in a 6 well plate. Grow to reach a confluency of about 70% in a cell incubator containing 5% CO2 and 95% air at 37 &#176;C.</li>
	<li>Wash HCMs with phosphate buffer saline (PBS). Expose them to different treatments (400 &#181;g/mL TXL or not) in 2 mL of serum/glucose-free DMEM for 30 min prior to hypoxia in a cell incubator containing 5% CO2 and 95% air at 37 &#176;C.</li>
	<li>Incubate HCMs in an airtight and hypoxic jar (2.5 L) fitted with a catalyst to scavenge free oxygen and induce hypoxia for 18 h (Figure 1A), and an anaerobic indicator to measure the oxygen tension in the medium11,12,13, in a cell incubator containing 5% CO2 and 95% air at 37 &#176;C. 
	Note: It takes about 1.5 h for the catalyst to scavenge free oxygen in the jar. Therefore, the HCMs are kept in the jar for 19.5 h before the jar is opened. The color of anaerobic indicator changes from light blue in normoxic environment to pale white in hypoxic condition (Figure 1B).</li>
	<li>Reoxygenate HCMs by moving the 6 well plate into an incubator set at 37 &#176;C containing 5% CO2 and 95% air for 2 h.</li>
</ol>
3. Preparation of HCMs for Flow Cytometry
<ol>
	<li>Warm the 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA) solution and PBS to 37 &#176;C.</li>
	<li>Aspirate medium from each well, rinse the cells with 1 mL of PBS and add 0.5 mL of 0.25% trypsin along the wall of the well.</li>
	<li>Incubate at 37 &#176;C until all HCMs are almost detached (about 1 min).</li>
	<li>Add 1 mL of DMEM complete medium (DMEM medium with 10% fetal bovine serum) to the wells to neutralize the trypsin. Transfer the cell suspension to a 15 mL centrifuge tube and pellet HCMs by centrifuging at 500 x g for 3 min at room temperature.</li>
	<li>Carefully aspirate the supernatant without disturbing the pellet.</li>
	<li>Add 0.5 mL of HCM complete medium and 0.5 mL of JC-1 working solution to each tube. Resuspend the HCMs by pipetting up and down.</li>
	<li>Cover the whole tubes with aluminum foil to prevent bleaching of the fluorescent indicator and incubate HCMs in normal conditions (at 37 &#176;C containing 5% CO2 and 95% air) for 20 min.</li>
	<li>Pellet HCMs by centrifuging at 500 x g for 3 min at 4 &#176;C.</li>
	<li>Rinse HCMs twice with 2 mL of ice-cold JC-1 staining buffer.</li>
	<li>Resuspend HCMs in a final volume of 300 &#181;L of ice-cold staining buffer in the sample tube (12 mm x 75 mm) and use the cells for analysis within 30 min. 
	Note: Use carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a potent inhibitor of respiratory electron transport chain, to treat HCMs for 1 h at a concentration of 20 &#181;M, for the positive control.</li>
</ol>
4. Flow Cytometer Setup
<ol>
	<li>Start the flow cytometer and ensure that the software is connected to it.</li>
	<li>Perform fluidics startup, start the stream and set up the breakoff.</li>
	<li>Open two dot plot windows in the flow cytometry software.
	<ol>
		<li>Select forward scattered light (FSC) on the X axis and side scattered light (SSC) on the Y axis in the first dot plot to represent the characteristics of the cells in terms of their size and their granularity, respectively.</li>
		<li>Select the PE (575 &#177; 26 nm) detection channel for the Y axis and FITC (530 &#177; 30 nm) for the X axis using a logarithmic scale in the second dot plot, which is used to measure the fluorescence intensity of JC-1 dye in the cells.</li>
	</ol>
	</li>
</ol>
5. Flow Cytometry Measurement and Data Analysis
<ol>
	<li>Click Unload to let out the tube support arm and position the Blank (HCMs that have not been dyed with JC-1) sample tube on it. Click Load to return the tube support arm to its working position.</li>
	<li>Click Record to collect particles from suspension in the Blank sample tube and then gate the cell population (P1) for further analysis in the first dot plot (Figure 3).</li>
	<li>Collect HCMs in other sample tubes and optimize the measurement by adjusting the voltages of fluorescence channels, to make sure that the cell dot is located in the central area of the second dot plot.</li>
	<li>Display the statistics of each sample tube and calculate the MMPs of each sample by dividing the ratio of red fluorescence intensity by that of green fluorescence intensity.</li>
</ol>

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The authors have nothing to disclose.

Here, we present a protocol that uses JC-1 dye to assess the MMP of cells after being exposed to H/R. Detected by JC-1 assay, the MMP of cells is independent of factors such as mitochondrial size, shape, and density that may influence single-component fluorescence signals14. Consequently, the results of JC-1 assay are relatively reliable. Besides, it is convenient and time-saving to perform the JC-1 assay. This assay has a low requirement for materials and reagents, and generally, it takes less than 1.5 h to do the staining, from cell detachment to MMP measurement.
A few critical steps cannot be overstated when performing the JC-1 assay if researchers want to obtain reliable and satisfactory results. First of all, the cells loaded with JC-1 should be stored in ice-cold staining buffer until they are collected by the flow cytometer. Temperature has a considerable effect on the stability of MMP and the value of one sample varies a lot after a brief time in room temperature. In addition, preliminary experiments are recommended to determine the pretreating time and concentration of CCCP, ensuring that the red/green ratio of the positive control is ~0. It is noteworthy that carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP), which can also eliminate MMP and inhibit oxidative phosphorylation, is an alternative to CCCP and can also be used to set up the positive control10.
The JC-1 assay has a major limitation in that other fluorochromes cannot be combined with JC-1, for its fluorescence emission spans the green, yellow, and part of the red wavelengths of the spectrum. Additionally, JC-1 assay must be combined with one or more methods, such as using calcein-acetoxymethylester15,16 or even genetic manipulation17,18, to determine if the opening of mPTP accounts for the decrease of MMP. This is because the opening of other mitochondrial pores like the ATP-sensitive K+ channels19,20 can also lead to the depolarization of mitochondria.
The JC-1 assay is best suited to more coarse &#34;yes/no&#34; assessments with the aim of evaluating whether mitochondria are largely polarized (e.g., apoptosis-related experiments)8. It has also been applied to measure MMP in a variety of cell types other than cardiomyocytes, including neurons21, hepatocytes22, renal cells23 and even isolated mitochondria24. In summary, we describe a quick, simple and sensitive method for the detection of MMP in HCMs after H/R.

Coronary heart disease is the leading cause of death worldwide. The treatment of choice for reducing ischemic injury and limiting infarct size in patients with ST-segment elevated myocardial infarction is timely and effective myocardial reperfusion via primary percutaneous coronary intervention (PCI)1,2. However, reperfusion causes additional damage, which can account for up to 30 percent of the final infarct size3. It is universally acknowledged that the mitochondrial permeability transition pore (mPTP) is not only central in mitochondrial damage and cell death during ischemia/reperfusion (I/R), but is also a converging target of cardioprotective signaling4,5. As the mPTP opening would bring about depolarization of the inner mitochondrial membrane potential (MMP)4, we detected mPTP opening using the 5,5&#8242;,6,6&#8242;-Tetrachloro-1,1&#8242;,3,3&#8242;-tetraethyl-imidacarbocyanineiodide (JC-1) assay.
The JC-1 assay is a cytofluorimetric method that is both qualitative and quantitative, and it has been further validated by analyzing the MMP at the level of a single mitochondria6. JC-1 exists as an aggregated form, yielding a red to orange colored emission (590 &#177; 17.5 nm) in the matrix of mitochondria with normal MMP; with the loss of MMP, JC-1 is converted to the monomeric form that yields green fluorescence with an emission of 530 &#177; 15 nm. Therefore, a decrease in the red/green fluorescence intensity ratio could indicate the reduction of MMP in conditions such as ischemia/reperfusion (I/R).
In addition to JC-1, MMP has also been studied with membrane-permeable lipophilic cations such as Rhodamine 123 and 3,3&#8242;-dihexyloxadicarbocyanine iodide [DiOC6(3)]. However, compared to these two probes, JC-1 is more reliable for analyzing MMP. Rhodamine 123 has relatively poor sensitivity (especially in quenching mode7,8) and poor specificity. The shift in rhodamine 123 sometimes is so small that it is hard for researchers or equipment to observe/detect. Besides, in a single cell, there are different mitochondrion binding sites for rhodamine 123 and so that it may have different fluorescence emissions9. DiOC6(3) is not recommended for detecting MMP either as it reacts sensitively to the depolarization of plasma membrane10.
Therefore, here we use the JC-1 assay to assess the MMP of HCMs after being exposed to hypoxia/reoxygenation with or without a protective agent.

<table><tbody><tr><td>Mitochondrial membrane potential assay kit with JC-1</td><td>Beyodtime, China</td><td>C2006</td><td>In the kit there are JC-1 stock solution (200&amp;times;), stock staining buffer (5&amp;times;) and CCCP(10mM)</td></tr><tr><td>Tongxinluo ultrafine powder</td><td>Shijiazhuang Yiling Pharmaceutical Co., China</td><td>071201</td><td></td></tr><tr><td>Annexin V-FITC/PI Kit</td><td>Becton-Dickinson, USA</td><td>556547</td><td></td></tr><tr><td>DMEM</td><td>Life Technologies, Grand Island Biological Company, USA</td><td>11966-025</td><td></td></tr><tr><td>Human cardiac myocyte</td><td>Promocell, Germany</td><td>C-12810</td><td></td></tr><tr><td>Myocyte Growth Medium&lt;br/&gt; (SupplementMix)</td><td>Promocell, Germany</td><td>C-39275</td><td></td></tr><tr><td>Myocyte Growth Medium (Ready-to-use)</td><td>Promocell, Germany</td><td>C-22070</td><td>used with Myocyte Growth Medium SupplementMix</td></tr><tr><td>GENbox</td><td>BioM&amp;eacute;rieux, Marcy l&amp;rsquo;Etoile, France</td><td>96127</td><td>2.5L</td></tr><tr><td>Catalyst (AnaeroPack)</td><td>MITSUBISHI GAS CHEMICAL&amp;nbsp;COMPANY, INC.&amp;nbsp;, Japan</td><td>&amp;nbsp;C-1</td><td></td></tr><tr><td>Anaerobic indicator</td><td>BioM&amp;eacute;rieux, Marcy l&amp;rsquo;Etoile, France</td><td>96118</td><td></td></tr><tr><td>Flow cytometer</td><td>Becton-Dickinson, USA</td><td>FACSAria 2</td><td></td></tr><tr><td>BD FACSDiva Software</td><td>Becton-Dickinson, USA</td><td></td><td>Version8.0.1</td></tr><tr><td>Sample tube</td><td>Corning science, USA</td><td>352054</td><td>12*75mm</td></tr><tr><td>PBS</td><td>Hyclone, USA</td><td>SH30256.01</td><td></td></tr></tbody></table>

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.

Before performing the JC-1 assay to evaluate the changes of MMP, it is highly recommended that experiments be carried out to confirm the conditions successfully set by the researchers. As shown by the flow cytometry results (Figure 2), compared with the normal group, hypoxia/reoxygenation (H/R) significantly induced the apoptosis of HCMs (Annexin V + /PI±), indicating that we had established a cell-based model of I/R (45.00 ± 2.13% vs. 11.50 ± 0.18% in the normal group, p &lt;0.05). Tongxinluo, a Chinese traditional medicine with cardioprotective effects, prevented the apoptosis of HCMs in condition of H/R (24.50 ± 1.13% vs. 45.00 ± 2.13% in the H/R group, p &lt;0.05).
CCCP is a protonophore that can inhibit oxidative phosphorylation in mitochondria by uncoupling the proton gradient (loss of MMP) established during the normal activity of electron carriers in the electron transport chain. Consequently, HCMs treated with CCCP was set as positive control. As shown in Figure 4, the red/green ratio in the CCCP-treated group was almost zero. Consistent with the results from the apoptosis assay, the H/R-treated group displayed a significantly lower red/green ratio compared with the normal group (0.69 ± 0.07 vs. 5.64 ± 0.21 in the normal group, p &lt;0.05) and Tongxinluo reversed such a ratio change (2.92 ± 0.22 vs. 0.69 ± 0.07 in the H/R group, p&lt;0.05).
<img alt="Anaerobic jar setup with catalyst, indicator, and GENbox; normoxic vs hypoxic conditions shown." class="xfigimg" src="/files/ftp_upload/57725/57725fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1. Establishment of hypoxia/reoxygenation (H/R) model. A. Materials needed to establish the H/R model; B. Confirmation of the hypoxic environment in the jar. The color of anaerobic indicator changes from light blue in normoxic environment to pale white in hypoxic condition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57725/57725fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Annexin-V/PI flow cytometry plots and bar graph showing apoptosis in normal, H/R, TXL cell samples." class="xfigimg" src="/files/ftp_upload/57725/57725fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2. Validation of the establishment of hypoxia/reoxygenation (H/R) model by flow cytometry. Compared with the normal group, the apoptotic rate of human cardiac myocytes was significantly higher in the H/R group, while Tongxinluo reversed this change (*p &lt;0.05 vs. Normal; # p &lt;0.05 vs. H/R; n = 3)11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57725/57725fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Flow cytometry scatter plot; FSC vs. SSC; cell population analysis result." class="xfigimg" src="/files/ftp_upload/57725/57725fig3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3. Gating cell population (P1, Red) in the SSC versus FSC dot plot for further analysis. The integrity of the cell population is assessed using side scatter (SSC) and forward scatter (FSC). Cell debris (Outside the gate P1, Black), which have reduced light scattering, are excluded. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57725/57725fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="JC-1 flow cytometry diagram; plots of control, normal, H/R, TXL; red/green ratio bar chart." class="xfigimg" src="/files/ftp_upload/57725/57725fig4.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4. Evaluation of mitochondrial membrane potential in human cardiac myocytes. Compared with the normal group, the H/R group displayed a significantly lower red/green ratio, while Tongxinluo reversed this trend (Red: cells with no fluorescence; Blue, cells with red fluorescence; Purple, cells with green fluorescence. Negative Control, blank and no staining; Positive Control, cells treated with CCCP; *p &lt;0.05 vs. Normal; #p &lt;0.05 vs. H/R; n = 3)11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57725/57725fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation at JoVE.com

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

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 ...

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Nanyang Technological University

Research

JoVE Journal

Medicine

14.2K Views.  Chinese Academy of Medical Sciences and Peking Union Medical College. Hier presenteren we een protocol dat gebruikmaakt van JC-1 kleurstof te beoordelen van de Mitochondriale membraanpotentiaal van cellen na wordt blootgesteld aan hypoxie/heroxygenatie met of zonder een beschermende agent.

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

Stable Isotope Tracing & LC-MS Analysis to Measure DHODH and SDH Activities

Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in mammalian cells. As oxygen (O2) is the most ubiquitous terminal electron acceptor for the mammalian ETC, the O2 consumption rate is frequently used as a proxy for mitochondrial function. However, emerging research demonstrates that this parameter is not always indicative of mitochondrial function, as fumarate can be employed as an alternative electron acceptor to sustain mitochondrial functions in hypoxia. This article compiles a series of protocols that allow researchers to measure mitochondrial function independently of the O2 consumption rate. These assays are particularly useful when studying mitochondrial function in hypoxic environments. Specifically, we describe methods to measure mitochondrial ATP production, de novo pyrimidine biosynthesis, NADH oxidation by complex I, and superoxide production. In combination with classical respirometry experiments, these orthogonal and economical assays will provide researchers with a more comprehensive assessment of mitochondrial function in their system of interest.

Author Spotlight: Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals  

Here, we present a compilation of assays to directly measure mitochondrial function in mammalian cells independently of their ability to consume molecular oxygen.

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in ...

Biochemistry

Respirometric Oxidative Phosphorylation Assessment in Saponin-permeabilized Cardiac Fibers

Investigation of mitochondrial function represents an important parameter of cardiac physiology as mitochondria are involved in energy metabolism, oxidative stress, apoptosis, aging, mitochondrial encephalomyopathies and drug toxicity. Given this, technologies to measure cardiac mitochondrial function are in demand. One technique that employs an integrative approach to measure mitochondrial function is respirometric oxidative phosphorylation (OXPHOS) analysis.
The principle of respirometric OXPHOS assessment is centered around measuring oxygen concentration utilizing a Clark electrode. As the permeabilized fiber bundle consumes oxygen, oxygen concentration in the closed chamber declines. Using selected substrate-inhibitor-uncoupler titration protocols, electrons are provided to specific sites of the electron transport chain, allowing evaluation of mitochondrial function. Prior to respirometric analysis of mitochondrial function, mechanical and chemical preparatory techniques are utilized to permeabilize the sarcolemma of muscle fibers. Chemical permeabilization employs saponin to selectively perforate the cell membrane while maintaining cellular architecture.
This paper thoroughly describes the steps involved in preparing saponin-skinned cardiac fibers for oxygen consumption measurements to evaluate mitochondrial OXPHOS. Additionally, troubleshooting advice as well as specific substrates, inhibitors and uncouplers that may be used to determine mitochondria function at specific sites of the electron transport chain are provided. Importantly, the described protocol may be easily applied to cardiac and skeletal tissue of various animal models and human samples.

Saponin-permeabilized fiber preparation in conjunction with respirometric oxidative phosphorylation analysis provides integrative assessment of mitochondrial function. Mitochondrial respiration in physiological and pathological states can reflect various regulatory influences including mitochondrial interactions, morphology and biochemistry.

Investigation of mitochondrial function represents an important parameter of cardiac physiology as mitochondria are involved in energy metabolism, ...

Biology

Mitochondrial Functional Assessment in Brush Lizard Liver Mitochondria

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of metabolic pathways. Therefore, experimentally assessing mitochondrial functionality can provide insight into variations among different populations or disease states. Additionally, it is valuable to assess whether isolated mitochondria are healthy enough to proceed with experiments. One characteristic often used to compare mitochondrial function in different samples is the rate of oxygen consumption. Oxygen consumption and subsequent calculation of the respiratory control ratio in either intact cells or mitochondria isolated from tissue can serve all three purposes. Using mitochondria isolated from the livers of brush lizards in conjunction with a phosphorescent probe that is sensitive to the fluctuations in oxygen concentration of a solution, we measured oxygen consumption using a fluorescent plate reader. This method is not only quick and efficient but also can be conducted with a small amount of mitochondria and without the need for specialized equipment. The step-by-step protocol described here increases the accessibility of mitochondrial functional assessment to researchers.

Assessment of oxygen consumption provides integral information about mitochondrial function. Using a phosphorescent probe with a fluorescent plate reader, accurate and reproducible data can be obtained easily without specialized equipment. This assay enables any lab to measure the oxygen consumption of isolated mitochondria and calculate respiratory control ratios.

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of ...

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with a molecular mass smaller than 1.5 kDa. Although the definitive molecular identity of the pore is still under debate, proteins such as cyclophilin D, VDAC and ANT contribute to mtPTP formation. While the involvement of mtPTP opening in cell death is well established1, accumulating evidence indicates that the mtPTP serves a physiologic role during mitochondrial Ca2+ homeostasis2, bioenergetics and redox signaling 3.
mtPTP opening is triggered by matrix Ca2+ but its activity can be modulated by several other factors such as oxidative stress, adenine nucleotide depletion, high concentrations of Pi, mitochondrial membrane depolarization or uncoupling, and long chain fatty acids4. In vitro, mtPTP opening can be achieved by increasing Ca2+ concentration inside the mitochondrial matrix through exogenous additions of Ca2+ (calcium retention capacity). When Ca2+ levels inside mitochondria reach a certain threshold, the mtPTP opens and facilitates Ca2+ release, dissipation of the proton motive force, membrane potential collapse and an increase in mitochondrial matrix volume (swelling) that ultimately leads to the rupture of the outer mitochondrial membrane and irreversible loss of organelle function.
Here we describe a fluorometric assay that allows for a comprehensive characterization of mtPTP opening in isolated mouse heart mitochondria. The assay involves the simultaneous measurement of 3 mitochondrial parameters that are altered when mtPTP opening occurs: mitochondrial Ca2+ handling (uptake and release, as measured by Ca2+ concentration in the assay medium), mitochondrial membrane potential, and mitochondrial volume. The dyes employed for Ca2+ measurement in the assay medium and mitochondrial membrane potential are Fura FF, a membrane impermeant, ratiometric indicator which undergoes a shift in the excitation wavelength in the presence of Ca2+, and JC-1, a cationic, ratiometric indicator which forms green monomers or red aggregates at low and high membrane potential, respectively. Changes in mitochondrial volume are measured by recording light scattering by the mitochondrial suspension. Since high-quality, functional mitochondria are required for the mtPTP opening assay, we also describe the steps necessary to obtain intact, highly coupled and functional isolated heart mitochondria.

A spectrofluorometric protocol for the measurement of the mitochondrial permeability transition pore opening in isolated mouse heart mitochondria is presented here. The assay involves the simultaneous measurement of mitochondria Ca2+ handling, mitochondrial membrane potential and mitochondrial volume. The procedure for obtaining high-quality and functional heart mitochondria is also described.

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with ...

Analyzing Oxygen Consumption Rate in Primary Cultured Mouse Neonatal Cardiomyocytes Using an Extracellular Flux Analyzer

Mitochondria and oxidative metabolism are critical for maintaining cardiac muscle function. Research has shown that mitochondrial dysfunction is an important contributing factor to impaired cardiac function found in heart failure. By contrast, restoring defective mitochondrial function may have beneficial effects to improve cardiac function in the failing heart. Therefore, studying the regulatory mechanisms and identifying novel regulators for mitochondrial function could provide insight which could be used to develop new therapeutic targets for treating heart disease. Here, cardiac myocyte mitochondrial respiration is analyzed using a unique cell culture system. First, a protocol has been optimized to rapidly isolate and culture high viability neonatal mouse cardiomyocytes. Then, a 96-well format extracellular flux analyzer is used to assess the oxygen consumption rate of these cardiomyocytes. For this protocol, we optimized seeding conditions and demonstrated that neonatal mouse cardiomyocytes oxygen consumption rate can be easily assessed in an extracellular flux analyzer. Finally, we note that our protocol can be applied to a larger culture size and other studies, such as intracellular signaling and contractile function analysis.

The goal of this protocol is to illustrate how to use mouse neonatal cardiomyocytes as a model system to examine how various factors can alter oxygen consumption in the heart.

Mitochondria and oxidative metabolism are critical for maintaining cardiac muscle function. Research has shown that mitochondrial dysfunction is an ...

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.

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 ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

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 ...

Measurement of Mitochondrial Mass and Membrane Potential in Hematopoietic Stem Cells and T-cells by Flow Cytometry

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production of all mature blood cells. In recent years cellular metabolism has emerged as a crucial regulator of HSC function and fate. We have previously demonstrated that modulation of mitochondrial metabolism influences HSC fate. Specifically, by chemically uncoupling the electron transport chain we were able to maintain HSC function in culture conditions that normally induce rapid differentiation. However, limiting HSC numbers often precludes the use of standard assays to measure HSC metabolism and therefore predict their function. Here, we report a simple flow cytometry assay that allows reliable measurement of mitochondrial membrane potential and mitochondrial mass in scarce cells such as HSCs. We discuss the isolation of HSCs from mouse bone marrow and measurement of mitochondrial mass and membrane potential post ex vivo culture. As an example, we show the modulation of these parameters in HSCs via treatment with a metabolic modulator. Moreover, we extend the application of this methodology on human peripheral blood-derived T cells and human tumor infiltrating lymphocytes (TILs), showing dramatic differences in their mitochondrial profiles, possibly reflecting different T cell functionality. We believe this assay can be employed in screenings to identify modulators of mitochondrial metabolism in various cell types in different contexts.

Here we describe a reliable method to measure mitochondrial mass and membrane potential in ex vivo cultured hematopoietic stem cells and T cells.

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production ...

Using Real-Time Cell Metabolic Flux Analyzer to Monitor Osteoblast Bioenergetics

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, prevent fracture. In the interest to both optimize peak bone mass and combat various musculoskeletal diseases (i.e., post-menopausal osteoporosis, anorexia nervosa, type 1 and 2 diabetes mellitus), incredible efforts have been made in the field of bone biology to fully characterize osteoblasts throughout their differentiation process. Given the primary role of mature osteoblasts to secrete matrix proteins and mineralization vesicles, it has been noted that these processes take an incredible amount of cellular energy, or adenosine triphosphate (ATP). The overall cellular energy status is often referred to as cellular bioenergetics, and it includes a series of metabolic reactions that sense substrate availability to derive ATP to meet cellular needs. Therefore, the current method details the process of isolating primary, murine bone marrow stromal cells (BMSCs) and monitoring their bioenergetic status using the Real-time cell metabolic flux analyzer at various stages in osteoblast differentiation. Importantly, these data have demonstrated that the metabolic profile changes dramatically throughout osteoblast differentiation. Thus, using this physiologically relevant cell type is required to fully appreciate how a cell's bioenergetic status can regulate the overall function.

Real-time cell metabolic flux assay measures the oxygen consumption rate and extracellular acidification rate, which corresponds to mitochondrial and glycolytic adenosine triphosphate production, using pH and oxygen sensors. The manuscript explains a method to understand the energy status of osteoblasts and the characterization and interpretation of the cellular bioenergetic status.

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, ...

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

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