We would like to thank the kind volunteers who donated blood for this project. We also extend our sincere appreciation to Dr. Ellen Schur and her team for providing us with additional samples from their study. We would also like to thank Andrew Kirsh for reviewing the manuscript and editing it for legibility. This work was supported by the following funding sources: P01AG001751, R01AG078279, P30AR074990, P30DK035816, P30DK017047, R01DK089036, K01HL154761, T32AG066574.

The collection of blood samples for data and methods development presented herein was approved by the Internal Review Board of the University of Washington. Representative results also include data from male C57BL/6J mice (5-7 months old) purchased from Jackson Laboratories. All animal procedures were approved by the University of Washington Office of Animal Welfare. The protocol overview is pictured in Figure 1. Reagent preparation for this protocol can be found in Supplementary File 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66863/66863fig01.jpg" /> 
Figure 1: Overview of the protocol. Workflow using high-resolution fluorespirometry to assess changes in mitochondrial membrane potential in isolated monocytes (CD14+) and T-cells (CD3+) from fresh human blood samples. Abbreviations: TMRM, tetramethylrhodamine methyl ester; SUIT, substrate-uncoupler-inhibitor titrations; ADP, adenosine diphosphate; Dig, digitonin; Mal, malate; Pyr, pyruvate; Glut, glutamate; D1-10, 10 consecutive ADP titrations. <a href="https://www.jove.com/files/ftp_upload/66863/66863fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Separation of buffy coat from whole blood
NOTE: Cell isolation is modified from Kramer et al.27.
<ol>
	<li>Allow RPMI, density gradient, and centrifuge to reach room temperature. Sterilize the biosafety cabinet and materials before starting.</li>
	<li>Collect venous blood into three 10 mL K2EDTA tubes. Invert the tubes at least 3 times.</li>
	<li>Centrifuge the tubes at 500 x g 10 min (22 &#176;C, 9 acceleration [acc], 2 deceleration [dec]).</li>
	<li>Remove 1 mL of plasma from each tube and store at -80 &#176;C for future analyses.</li>
	<li>Transfer the plasma and half the red blood cell layer from each tube into a single 50-mL conical tube. Add RPMI up to the 40 mL mark. Invert at least 3 times.</li>
	<li>Slowly layer 10 mL of the plasma solution into four 15 mL conical tubes containing 3 mL of the density gradient.</li>
	<li>Centrifuge at 700 x g for 30 min (22 &#176;C, 5 acc, 2 dec).</li>
	<li>Collect all the plasma and the buffy coat containing the peripheral mononuclear cells (PBMCs) without disrupting the red blood cells.</li>
	<li>Centrifuge at 500 x g for 10 min (22 &#176;C, 5 acc, 5 dec) and aspirate the supernatant.</li>
	<li>Wash the PBMC pellet 1x-2x by resuspending it in 10 mL of RPMI and centrifuging at 500 x g for 10 min (22 &#176;C, 5 acc, 5 dec).</li>
</ol>
2. Magnetic separation of CD14+ and CD3+ cells
<ol>
	<li>Place a column in the magnetic field of a magnetic cell separator (see the Table of Materials). Wash the column with 3 mL of RP-5.</li>
	<li>Resuspend the PBMC pellet in 80 &#181;L of RP-5 and 20 &#181;L of anti-CD14 microbeads (see the Table of Materials). Incubate for 15 min at 4 &#176;C.</li>
	<li>Resuspend the cells with 1 mL of RP-5 and load the suspension onto the column. Collect unlabeled cells that flow through into a 15 mL conical tube labeled &#34;Flow-through 1&#34;. Wait until all cell suspension has gone through the column, then continue washing with 3 mL of RP-5 3x, collecting all flow-through.</li>
	<li>Carefully remove the column from the magnetic field and place it on a new 15 mL conical tube. Add 5 mL of RP-5 and immediately use the plunger to purge the column contents into a collection tube labeled &#34;CD14+&#34;.</li>
	<li>Centrifuge &#34;Flow-through 1&#34; at 500 x g for 10 min (22 &#176;C, 5 acc, 5 dec) and aspirate the supernatant.</li>
	<li>Using cells from Flow-through 1, repeat steps 2.2-2.5 using anti-CD3 microbeads (see the Table of Materials) to isolate T-cells.</li>
	<li>Centrifuge tubes containing T-cells (CD3+) and monocytes (CD14+) at 300 x g for 5 min. Aspirate the supernatant and resuspend the pellet in 1 mL of RP-5.</li>
	<li>Determine cell concentration using a hemocytometer or automatic cell counter. 
	NOTE: Cells can be counted by adding 10 &#181;L of a cell dilution of 1:10 or 1:20 to a hemocytometer. One may refer to previously published protocols to count cells using a hemocytometer36.</li>
	<li>Pipette 2.5 million monocytes or 5 million T-cells into a new centrifuge tube. Centrifuge for 30 s at 2000 x g, aspirate the supernatant and resuspend the cells in MiR05 for a total volume of 20 &#181;L and final concentration of 125 million monocytes or 250 million T-cells per mL. 
	NOTE: The final concentration was selected to inject 2.5 million monocytes or 5 million T-cells in a volume of 20 &#181;L. Mouse T-cells isolated from the spleen have also been tested using this method. The procedure is found in Supplementary File 1.</li>
</ol>
3. High-resolution fluorespirometry - Oxygen and TMRM fluorescence calibration
NOTE: This method was adapted from previous work done on permeabilized fibers by Pharaoh et al.11. A high, un-inhibitory concentration of TMRM is used for quench mode, where the relationship of mMP and TMRM concentration in the matrix is inverted. Thus, a decrease in mMP leads to the release of TMRM dye from the matrix and an increase in fluorescence32.
<ol>
	<li>Install 0.5 mL chambers in the O2K respirometer according to the manufacturer&#39;s instructions (see Table of Materials). Turn on the instrument and connect it to the software provided by the manufacturer for data acquisition.</li>
	<li>Adjust the temperature to 37 &#176;C and stir speed to 750 rpm.</li>
	<li>Wash the chambers with distilled water&#160;3x. Replace water with 0.54 mL of Mir05, close the stoppers fully, and remove excess buffer with the integrated suction system (ISS). Raise stoppers to allow room oxygen to equilibrate with chamber oxygen using a stopper-spacer.</li>
	<li>Once the oxygen flux is stable, perform the air oxygen calibration (R1) according to the manufacturer&#39;s instructions. 
	NOTE: It may take &#62;30 min for oxygen flux to stabilize. Oxygen sensors require the determination of zero-point oxygen (R0) and background oxygen flux from 50-200 &#181;M from separate experiments using dithionite titrations. Specific methods can be found in the manufacturer&#39;s manual.</li>
	<li>Seal the chamber by closing the stoppers. 
	NOTE: Unlike experiments using permeabilized fibers, the chambers do not require hyperoxygenation for PBMCs. Sealing the chamber after R1 calibration provides sufficient oxygen for the experiment. The oxygen levels should be maintained between 50-250 &#181;M. If oxygen concentration falls below the threshold, the chamber can be partially opened so that chamber oxygen can equilibrate with room air oxygen.</li>
	<li>TMRM calibration
	<ol>
		<li>Use the Green LED fluo-sensors (ex. 525 nm) with the AmR filter set (see Table of Materials). Set fluorometer Gain to 1000 and Intensity to 1000. Turn on the fluo-sensors and begin recording the baseline.</li>
		<li>Inject 2.5 &#181;L of 0.05 mM TMRM and allow the signal to stabilize (~2 min) before the next 2.5 &#181;L injection until a total of 4 injections are performed for a total TMRM concentration of 1 &#181;M TMRM in the chamber. Use a Hamilton syringe for all injections.</li>
		<li>Calibrate the fluo-sensor by selecting the fluorescent signal (voltage) for each injection representing 0, 0.25, 0.5, 0.75, and 1.0 &#181;M of TMRM for a five-point calibration.</li>
	</ol>
	</li>
</ol>
4. Substrate-uncoupler-inhibitor titration (SUIT) protocol
NOTE: Run blank experiments where 20 &#181;L of Mir05 is injected into the chamber instead of 20 &#181;L of cell suspension, as the TMRM signal will change in response to injections alone (discussed in representative results). Allow for the oxygen flux signal to stabilize (about 2-3 min) before the next injection for both blank and sample experiments. The following titration protocol and expected observations are in Table 1.
<ol>
	<li>Once oxygen flux is stable, select and label both the oxygen flux and TMRM signal &#34;pre-cell&#34;.</li>
	<li>Inject the cell suspension containing either 5 million T-cells or 2.5 million monocytes in ~20 &#181;L and measure for about 10 min. Select and label both the oxygen flux and TMRM signal as &#34;Cell&#34;.</li>
	<li>Permeabilize cells by injecting 2 &#181;L of 1 mg/mL digitonin (final concentration: 4 &#181;g/mL). Wait for 20 min. Select and label both the oxygen flux and TMRM signal as &#34;Dig&#34;. 
	NOTE: It is suggested to optimize the digitonin concentration in separate experiments.</li>
	<li>Add 2.5 &#181;L of 1 M succinate (final concentration: 5 mM). Select and label both the oxygen flux and TMRM signal as &#34;SUCC&#34;.</li>
	<li>Once oxygen flux is stable, add 5 &#181;L of 100 mM malate (final concentration: 1.0 mM), 5 &#181;L of 1 M glutamate (final concentration: 10 mM), and 5 &#181;L of 500 mM pyruvate (final concentration: 5 mM). Select and label both the oxygen flux and TMRM signal as &#34;MPG&#34;.</li>
	<li>Once oxygen flux is stable, titrate ADP. Select rates for each titration and label them &#34;D&#34; sequentially 1 through 10, depending on the number of titrations. Use the titration scheme in Table 2.</li>
	<li>Once oxygen flux is stable, perform a series of 1 &#181;L titrations of 0.25 mM carbonyl cyanide p-(tri-fluromethoxy) phenyl-hydrazone (FCCP) until the fluorescent signal reaches its maximum. Select and label both the oxygen flux and TMRM signal representing minimum membrane potential and label it &#34;FCCP&#34;. 
	NOTE: FCCP concentration of 0.5-1.0 &#181;M is usually required to deplete mitochondrial membrane potential. 
	CAUTION: FCCP is toxic. Refer to the safety data sheet (SDS) for proper handling.</li>
	<li>OPTIONAL: Once oxygen flux is stable, inject 1 &#181;L of 0.25 mM rotenone to inhibit complex I and determine respiratory capacity through complex II. 
	NOTE: Changes in membrane potential are no longer relevant after titrating with uncoupler. 
	CAUTION: Rotenone is toxic. Refer to SDS for proper handling.</li>
	<li>Once oxygen flux is stable, inject 1 &#181;L of 1.25 mM (final concentration: 2.5 &#181;M) of Antimycin A to inhibit mitochondrial respiration. 
	CAUTION: Antimycin A is toxic. Refer to SDS for proper handling.</li>
</ol>
5. Calculation of mitochondrial membrane potential and analysis
<ol>
	<li>Using blank experiments, record the calibrated TMRM values (micromolar TMRM) before the injection of the blank sample (&#34;pre-sample&#34;) and for each of the injections. See Figure 2.</li>
	<li>For each blank experiment, calculate the&#160;background ratio by setting the &#34;pre-sample&#34; TMRM concentration to 1.0. Calculate the subsequent proportional decrease in TMRM. Calculate the average background ratio from all blank experiments. 
	NOTE: The number of blank experiments to include may depend on the instrument&#39;s precision. See the calculation example in Table 3 from five different blank experiments, where the standard deviation of the average background ratio fell between 0 and 0.016 for each titration.</li>
	<li>Background calculation: Calculate the background for each sample experiment by multiplying &#34;pre-sample&#34; TMRM of the sample experiment times the average background ratio for each injection. See the calculation example in Table 3.</li>
	<li>Background correction:&#160;Subtract the experiment&#39;s background to the measured TMRM values of the sample. See the calculation example in Table 4.</li>
	<li>FCCP correction:&#160;Subtract the FCCP background-corrected mMP from each injection. See the calculation example in Table 4.</li>
	<li>ADP sensitivity curve: Normalize the ADP-driven decrease in mMP by setting the highest and lowest membrane potential as 100% and 0%, respectively, using the mMP values collected throughout the ADP titration. Fit the data into a non-linear fit regression model using the preferred statistical software to calculate the half-maximal inhibitory concentration (IC50) of ADP on mMP. 
	NOTE: The curve fits the [Inhibitor] vs. normalized response - Variable slope in Prism.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66863/66863fig02.jpg" /> 
Figure 2: Calculating mitochondrial membrane potential (mMP) and ADP sensitivity from TMRM fluorescence. Steps for calculating mitochondrial membrane potential (mMP) and ADP sensitivity from measurements of TMRM fluorescence by high-resolution fluorespirometry of one sample of T-cells (n = 1). Step 1: TMRM fluorescence is measured in blank samples as done in the biological sample. Step 2: Determine the ratio in the TMRM signal with each titration relative to the signal prior to the sample for each blank experiment. Calculate the average for each titration of all blank experiments. Step 3: Calculate the background for each sample experiment by multiplying the &#34;pre-sample&#34; fluorescence by the average background ratio for each titration. Step 4: Calculate the difference between background and sample TMRM fluorescence for each titration to express data as mMP or mitochondrial TMRM uptake. Step 5: Correct mMP so that full uncoupling with FCCP reflects zero mMP. Step 6: Perform non-linear regression to graph changes in mMP with increasing ADP concentrations. Measurements were performed in 0.5 mL chambers, one containing 5 million T-cells from a healthy volunteer. Averaged data are expressed as mean &#177; SEM. Single data points of a single replicate are expressed without error bars. <a href="https://www.jove.com/files/ftp_upload/66863/66863fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

High-Resolution Analysis of Mitochondrial Function in Human PBMCs

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<li>Williams, A. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Disruption+of+Acetyl-Lysine+turnover+in+muscle+mitochondria+promotes+insulin+resistance+and+redox+stress+without+overt+respiratory+dysfunction%5BTitle%5D)+AND+%22Cell+Metab%22%5BJournal%5D)">Disruption of Acetyl-Lysine turnover in muscle mitochondria promotes insulin resistance and redox stress without overt respiratory dysfunction.</a> Cell Metab. 31 (1), 131-147 (2020).</li>
<li>Krumschnabel, G., Eigentler, A., Fasching, M., Gnaiger, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Use+of+safranin+for+the+assessment+of+mitochondrial+membrane+potential+by+high-resolution+respirometry+and+fluorometry%5BTitle%5D)+AND+%22Methods+Enzymol%22%5BJournal%5D)">Use of safranin for the assessment of mitochondrial membrane potential by high-resolution respirometry and fluorometry.</a> Methods Enzymol. 542, 163-181 (2014).</li>
<li>Chowdhury, S. R., Djordjevic, J., Albensi, B. C., Fernyhough, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Simultaneous+evaluation+of+substrate-dependent+oxygen+consumption+rates+and+mitochondrial+membrane+potential+by+TMRM+and+safranin+in+cortical+mitochondria%5BTitle%5D)+AND+%22Biosci+Rep%22%5BJournal%5D)">Simultaneous evaluation of substrate-dependent oxygen consumption rates and mitochondrial membrane potential by TMRM and safranin in cortical mitochondria.</a> Biosci Rep. 36 (1), 00286(2015).</li>
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<li>Perry, S. W., Norman, J. P., Barbieri, J., Brown, E. B., Gelbard, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mitochondrial+membrane+potential+probes+and+the+proton+gradient%3A+a+practical+usage+guide%5BTitle%5D)+AND+%22Biotechniques%22%5BJournal%5D)">Mitochondrial membrane potential probes and the proton gradient: a practical usage guide.</a> Biotechniques. 50 (2), 98-115 (2011).</li>
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<li>Geltink, R. I. K., Kyle, R. L., Pearce, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Unraveling+the+complex+interplay+between+T+cell+metabolism+and+function%5BTitle%5D)+AND+%22Annu+Rev+Immunol%22%5BJournal%5D)">Unraveling the complex interplay between T cell metabolism and function.</a> Annu Rev Immunol. 36, 461-488 (2018).</li>
<li>Sukumar, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Inhibiting+glycolytic+metabolism+enhances+CD8%2B+T+cell+memory+and+antitumor+function%5BTitle%5D)+AND+%22J+Clin+Invest%22%5BJournal%5D)">Inhibiting glycolytic metabolism enhances CD8+ T cell memory and antitumor function.</a> J Clin Invest. 123 (10), 4479-4488 (2013).</li>
<li>Gerriets, V. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Foxp3+and+Toll-like+receptor+signaling+balance+T%5BTitle%5D)+AND+%22Nat+Immunol%22%5BJournal%5D)">Foxp3 and Toll-like receptor signaling balance T.</a> Nat Immunol. 17 (12), 1459-1466 (2016).</li>
<li>MacIver, N. J., Michalek, R. D., Rathmell, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Metabolic+regulation+of+T+lymphocytes%5BTitle%5D)+AND+%22Annu+Rev+Immunol%22%5BJournal%5D)">Metabolic regulation of T lymphocytes.</a> Annu Rev Immunol. 31, 259-283 (2013).</li>
<li>Desdín-Micó, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((T+cells+with+dysfunctional+mitochondria+induce+multimorbidity+and+premature+senescence%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">T cells with dysfunctional mitochondria induce multimorbidity and premature senescence.</a> Science. 368 (6497), 1371-1376 (2020).</li>
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</ol>

The authors declare no conflicts of interest.

This protocol uses high-resolution fluorespirometry to measure the sensitivity of the mitochondrial response to energy demand by measuring the dissipation of mMP in response to increasing levels of ADP in PBMCs, monocytes, and T-cells. This is done by adding complex I and II substrates to maximize the mitochondrial membrane potential and titrating ADP to gradually stimulate ATP-synthase to use the proton gradient for ATP generation.
Critical steps in the protocol include setting the gain and intensity of the fluorophore to 1000 and making sure a TMRM fluorescent signal is acquired during the TMRM titration. Because TMRM fluorescence declines following each titration (a limitation of this method), it is imperative to run background experiments using blank samples. We have also found that DMSO has an inhibitory effect on mitochondrial respiration and membrane potential and, therefore, recommend diluting the working solution of TMRM in Mir05 (Supplementary Figure 1). 
Some modifications that may be used when trying this protocol are adjusting cell concentrations and using the standard 2 mL chamber. However, the 0.5 mL chamber is preferred for T-cells and monocytes because of the high concentration of cells needed for optimal response in membrane potential and oxygen flux. A lower concentration of cells may be optimal when testing cells with greater respiratory capacity, like macrophages.
Additional limitations of the method presented here include the requirement for at least 5 million T-cells and 2.5 million monocytes. We can often obtain enough cells from ~20 mL of blood from healthy participants, but these numbers can vary by health status, age, and sex26. In addition, as in most methods assessing mitochondrial capacity, the cells need to be freshly isolated. However, this method could be tried in cryopreserved cells in the future. In comparison with the yield from human blood, the T-cell yield from spleens of healthy mice is high enough to conduct this assay.
Circulating T-cells, particularly long-lived memory (TM) and regulatory (Treg) cells, rely on oxidative phosphorylation for energy37. While their energy demand and oxygen consumption are low (e.g., compared with that of resting muscle), their survival is essential for an effective immune response to reinfection and cancer38,39,40. A reduction in T-cell oxidative phosphorylation results in impaired proliferative capacity and promotes T-cell exhaustion and senescence5,41. Additionally, mitochondrial hyperpolarization promotes a sustained production of cytokines (IL-4 and IL-21) by effector CD4 T-cells during activation42. Upon infection, the energy requirement for activation and proliferation of immune cells can be as high as 25%-30% of the basal metabolic rate43. Therefore, immune cells function in a wide and extreme range of energy demands, and this protocol can test mitochondrial responses within that range.
Chronic inflammation is a common feature of obesity, diabetes, and aging. Dysregulated levels of circulating hormones, lipids, and glucose have systemic impacts and can thus affect how mitochondria respond to an energetic challenge. Here, we have presented a method to assess mitochondrial ADP sensitivity in circulating PBMCs. Further studies are needed to determine how ADP sensitivity may be modulated in metabolic disease and how it impacts health status.

A cell&#39;s ability to function and survive in a period of physiological stress is largely dependent on its ability to meet the energetic requirement to restore homeostasis1,2. Energy demand rises in response to a variety of stimuli. For instance, increased muscle contraction during exercise increases the utilization of ATP and glucose by skeletal muscle, and a rise in protein synthesis following infection increases the utilization of ATP by immune cells for cytokine production and proliferation3,4,5,6. A spike in energy demand triggers a series of bioenergetic processes to restore the ATP/ADP ratio. As ATP is consumed, ADP levels rise and stimulate F1F0&#160;ATP-synthase (complex V), which requires a protonmotive force to drive its mechanical rotation and catalytic conversion of ADP to ATP within the mitochondrion7. The protonmotive force is an electrochemical gradient created by the pumping of protons during the transfer of electrons from substrates to oxygen through the electron transport system (ETS) within the inner mitochondrial membrane. The resulting difference in proton concentration (delta pH) and electrical potential (membrane potential) creates the protonmotive force that drives ATP synthesis and oxygen consumption in response to energy demand reducing the ATP/ADP ratio or raising ADP levels. The affinity of mitochondria to ADP can be determined by the calculation of the Km or EC50 of ADP-stimulated respiration of isolated mitochondria or permeabilized cells8,9. This method has shown that permeabilized muscle fibers from older humans require a greater concentration of ADP to stimulate 50% of their maximal oxidative phosphorylation capacity than those of younger subjects9. Similarly, aging mouse skeletal muscle requires more ADP to lower the production of mitochondrial reactive oxygen species (ROS)10,11. Additionally, ADP sensitivity is reduced in permeabilized muscle fibers of mice with diet-induced obesity relative to controls and is enhanced in the presence of insulin and following nitrate consumption12,13. Thus, the capacity of mitochondria to respond to energy demand varies under different physiological conditions, but this has not been previously explored in the context of immune cells.
Peripheral blood mononuclear cells (PBMCs) are commonly used to investigate cellular bioenergetics in human subjects14,15,16,17,18,19,20. This is largely due to cells being easily obtainable from uncoagulated blood samples in clinical studies, the responsiveness of cells to metabolic perturbations, and the methods developed by various groups to interrogate mitochondrial metabolism by using inhibitors and uncouplers to determine the maximal and minimal capacity of mitochondrial respiration21,22. These methods have led to an appreciation of the roles of bioenergetics in aging, metabolic disease, and immune function14,20,23,24. Mitochondrial respiratory capacity is often reduced in skeletal muscle and PBMCs under conditions of heart failure18,25. PBMC bioenergetics are also correlated with cardiometabolic risk factors in healthy adults17 and are responsive to treatments such as nicotinamide riboside18. PBMCs include neutrophils, lymphocytes (B-cells and T-cells), monocytes, natural killer cells, and dendritic cells, which all contribute to PBMC mitochondrial capacity26,27,28. In addition, cellular bioenergetics play a crucial role in immune cell activation, proliferation, and renewal23. However, a limitation of these methods is that the cells are not functioning under a physiological range of substrates. Additional methods are therefore required to interrogate mitochondrial function in substrate concentrations that are more relevant to what cells experience in vivo.
Mitochondrial membrane potential (mMP) is the major component of a protonmotive force and is essential for a variety of mitochondrial processes beyond ATP production, such as regulation of respiratory flux, reactive oxygen species production, protein and ion import, autophagy, and apoptosis. mMP can be assessed with electrochemical probes or fluorescent dyes sensitive to changes in membrane polarization like JC-1, Rhod123, DiOC6, tetramethyl rhodamine (TMRE) or methyl ester (TMRM), and safranin. The latter two are lipophilic cationic dyes that have been successfully used in high-resolution fluorespirometry of tissue homogenates, isolated mitochondria, and permeabilized tissue11,29,30,31,32,33. In this technique, TMRM is used in quench mode, where cells are exposed to a high concentration of TMRM that accumulates in the mitochondrial matrix when polarized (high mMP and protonmotive force), resulting in the quenching of cytosolic TMRM fluorescence. When mitochondria depolarize in response to ADP or uncouplers, the dye is released from the matrix, increasing the TMRM fluorescent signal34,35. The purpose of this method is to simultaneously measure changes in mitochondrial respiration and mMP in response to ADP titrations in human-derived PBMCs, circulating monocytes, and T-cells, and it can also be applied to mouse splenic T-cells.

<table><tbody><tr><td>Adenosine Diphosphate</td><td>Sigma-Aldrich</td><td>A5285</td><td>Fluorespirometry</td></tr><tr><td>Antimycin A</td><td>Sigma-Aldrich</td><td>A8674</td><td>Fluorespirometry</td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma-Aldrich</td><td>A6003</td><td>Mir05 buffer</td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma-Aldrich</td><td>A6003</td><td>Cell isolation</td></tr><tr><td>Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone</td><td>Sigma-Aldrich</td><td>C2920</td><td>Fluorespirometry</td></tr><tr><td>Cell strainers</td><td>Fisher Scientific</td><td>22-363-548</td><td>Isolation of T-cells from mouse spleen protocol</td></tr><tr><td>CD14 Microbeads, human</td><td>Miltenyi Biotec</td><td>130-050-201</td><td>Cell isolation</td></tr><tr><td>CD3 Microbeads, human</td><td>Miltenyi Biotec</td><td>130-050-101</td><td>Cell isolation</td></tr><tr><td>DatLab</td><td>Oroboros</td><td>Version 8</td><td></td></tr><tr><td>Digitonin</td><td>Sigma-Aldrich</td><td>D141</td><td>Fluorespirometry</td></tr><tr><td>D-Sucrose</td><td>Sigma-Aldrich</td><td>84097</td><td>Mir05 buffer</td></tr><tr><td>Ethylene glycol-bis(&amp;beta;-aminoethyl ether)-N,N,N&amp;prime;,N&amp;prime;-tetraacetic acid (EGTA)</td><td>Sigma-Aldrich</td><td>E4378</td><td>Mir05 buffer</td></tr><tr><td>Filter Set AmR</td><td>Oroboros</td><td>44321-01</td><td></td></tr><tr><td>HBSS (10x)</td><td>Gibco</td><td>12060-040</td><td></td></tr><tr><td>HEPES&amp;nbsp;sodium salt</td><td>Sigma-Aldrich</td><td>H7523</td><td>Mir05 buffer</td></tr><tr><td>Histopaque 1077</td><td>Sigma-Aldrich</td><td>10771</td><td>Cell isolation</td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;EDTA blood collection tubes</td><td>BD Vacutainer</td><td>366643</td><td>Cell isolation</td></tr><tr><td>Lactobionic acid</td><td>Sigma-Aldrich</td><td>153516</td><td>Mir05 buffer</td></tr><tr><td>L-Glutamic acid</td><td>Sigma-Aldrich</td><td>G1626</td><td>Fluorespirometry</td></tr><tr><td>L-Malic Acid</td><td>Sigma-Aldrich</td><td>M1000</td><td>Fluorespirometry</td></tr><tr><td>LS Columns</td><td>Miltenyi Biotec</td><td>130-042-401</td><td>Cell isolation</td></tr><tr><td>Magnesium Chloride&amp;nbsp;(MgCl&lt;sub&gt;2&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>M9272</td><td>Mir05 buffer</td></tr><tr><td>Multi-MACS stand and MidiMACS Separator</td><td>Miltenyi Biotec</td><td>130-042-301</td><td>Cell isolation</td></tr><tr><td>O2k-Fluo Smart-Module</td><td>Oroboros</td><td>12100-03</td><td></td></tr><tr><td>O2k-FluoRespirometer series J</td><td>Oroboros</td><td>10201-03</td><td></td></tr><tr><td>O2k-sV-Module (0.5 chamber)</td><td>Oroboros</td><td>11200-01</td><td></td></tr><tr><td>Oligomycin</td><td>Sigma-Aldrich</td><td>04876</td><td>Fluorespirometry</td></tr><tr><td>Pan T Cell Isolation Kit II, mouse</td><td>Miltenyi</td><td>130095130</td><td>Isolation of T-cells from mouse spleen protocol</td></tr><tr><td>Potassium dihydrogen phosphate (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>P0662</td><td>Mir05 buffer</td></tr><tr><td>Potassium Hydroxide&amp;nbsp;(KOH)</td><td>Sigma-Aldrich</td><td>221473</td><td>Mir05 buffer</td></tr><tr><td>Prism</td><td>GraphPad</td><td>Version 10</td><td></td></tr><tr><td>Rotenone</td><td>Sigma-Aldrich</td><td>R8875</td><td>Fluorespirometry</td></tr><tr><td>RPMI Buffer</td><td>Corning</td><td>17-105-CV</td><td>Cell isolation</td></tr><tr><td>Sodium Pyruvate</td><td>Sigma-Aldrich</td><td>P2256</td><td>Fluorespirometry</td></tr><tr><td>Succinate disodium salt</td><td>Sigma-Aldrich</td><td>S2378</td><td>Fluorespirometry</td></tr><tr><td>Taurine</td><td>Sigma-Aldrich</td><td>T0625</td><td>Mir05 buffer</td></tr><tr><td>Tetramethyrhodamine methyl ester perchlorite</td><td>Sigma-Aldrich</td><td>T5428</td><td>Fluorespirometry</td></tr></tbody></table>

high-resolution fluorespirometry to assess dynamic changes in mitochondrial membrane potential in human immune cells

Peripheral mononuclear cells (PBMCs) exhibit robust changes in mitochondrial respiratory capacity in response to health and disease. While these changes&#160;do not always reflect what occurs in other tissues, such as skeletal muscle, these cells are an accessible and valuable source of viable mitochondria from human subjects. PBMCs are exposed to systemic signals that impact their bioenergetic state. Thus, expanding our tools to interrogate mitochondrial metabolism in this population will elucidate mechanisms related to disease progression. Functional assays of mitochondria are often limited to using respiratory outputs following maximal substrate, inhibitor, and uncoupler concentrations to determine the full range of respiratory capacity, which may not be achievable in vivo. The conversion of adenosine diphosphate (ADP) to adenosine triphosphate (ATP) by ATP-synthase results in a decrease in mitochondrial membrane potential (mMP) and an increase in oxygen consumption. To provide a more integrated analysis of mitochondrial dynamics, this article describes the use of high-resolution fluorespirometry to measure the simultaneous response of oxygen consumption and mitochondrial membrane potential (mMP) to physiologically relevant concentrations of ADP. This technique uses tetramethylrhodamine methylester (TMRM) to measure mMP polarization in response to ADP titrations following maximal hyperpolarization with complex I and II substrates. This technique can be used to quantify how changes in health status, such as aging and metabolic disease, affect the sensitivity of mitochondrial response to energy demand in PBMCs, T-cells, and monocytes from human subjects.

To illustrate the differences in optimal cell concentration for the assay, 5 million T-cells were loaded into one 0.5 mL chamber (10 million cells/mL), and 1.25 million cells were loaded into another chamber (2.5 million cells/mL) containing 1 &#181;M TMRM (Figure 3A-G). Three blank experiments were also included to calculate the TMRM background. We found that a higher concentration of T-cells resulted in a more distinguishable change in TMRM fluorescence relative to the background (Figure 3B,D). In addition, a higher cell concentration allowed us to detect the expected increase in oxygen consumption and simultaneous depletion of the mMP in response to the addition of FCCP (Figure 3E,F). Using a low concentration of cells yielded a weak change in fluorescence that paralleled the background. Since the calculation of mMP subtracts the background from the signal, a low cell concentration does not allow for the determination of changes in mMP in response to substrates and uncouplers. In addition to using the higher concentrations of cells in this assay, we recommend keeping the cell concentration constant for each cell type between experiments.
To validate the influence of ATP-synthase in the dissipation of mMP with ADP titrations, we ran parallel experiments on PBMCs and T-cells where one chamber received oligomycin before ADP titration (Figure 4). We found no dissipation of mMP in response to ADP in cells treated with oligomycin, suggesting that the gradual decrease in mMP with ADP is a result of proton flux through ATP-synthase (Figure 4A-F). We also compared ADP sensitivity between T-cells and PBMCs of the same participant and found ADP sensitivity to be lower (higher EC50) in the T-cell fraction (Figure 4G,H).
We conducted a series of blank experiments to determine the influence of time or the SUIT protocol on TMRM fluorescence. We found that the TMRM signal in blank experiments is mostly influenced by SUIT titrations (Figure 5A) as opposed to the timing of the titrations (Figure 5B).
We compared ADP-driven changes in oxygen consumption rates (OCR) and in mMP in T-cells and monocytes from 11 healthy, community-dwelling volunteers (Figure 6A-H). Similar to the results of previously published experiments using extracellular flux and enzymatic assays, monocytes exhibited a greater mitochondrial respiratory capacity than lymphocytes26,27 (Figure 6A,H). However, we did not detect a typical dose-response increase in OCR with ADP in either cell type (Figure 6C,D), contrary to what this method shows when using highly metabolic tissues like mouse liver (Figure 7A-H). On the other hand, the use of TMRM allowed us to detect a gradual decline in mMP with ADP in human immune cells (Figure 6E-G) and in splenic T-cells from mice (Figure 7E-H). While we did not directly compare human and mouse T-cells using the same titration protocol, we did find that the IC50 of mouse T cells was lower by a factor of 10 compared with that of circulating T-cells from human subjects.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66863/66863fig03v2.jpg" /> 
Figure 3: High-resolution fluorespirometry experiments. (A-D) Trace of high-resolution fluorespirometry experiments using T-cell concentrations of 10 million cells/mL and 2.5 million cells/mL in 0.5 mL chambers. (A) 10 million cells/mL in 0.5 mL chambers. (C) 2.5 million cells/mL in 0.5 mL chambers. Oxygen flux (pmol/s/mL) is shown in the top panel (red), and the calibrated TMRM signal is shown in the bottom panel (black). Changes in TMRM throughout the SUIT for the sample and its calculated background were plotted for the chambers containing (B) 10 million cells/mL and (D) 2.5 million cells/mL. (E) For each cell concentration, oxygen flux (pmol/s/million cells) and (F) mitochondrial membrane potential were calculated. (G) ADP sensitivity curve was plotted and fit to a non-linear regression model (solid lines). Abbreviations: mMP, mitochondrial membrane potential; TMRM, tetramethylrhodamine methyl ester; SUIT, substrate-uncoupler-inhibitor titrations; ADP, adenosine diphosphate; Dig, digitonin; Mal, malate; Pyr, pyruvate; Glut, glutamate; D1-11, 11 consecutive ADP titrations; U, uncoupler FCCP of 0.5 and 1.0 &#956;M; AMA, antimycin A. <a href="https://www.jove.com/files/ftp_upload/66863/66863fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66863/66863fig04v2.jpg" /> 
Figure 4: ATP-synthase drives ADP-driven decrease in membrane potential in T-cells and PBMCs. (A-H) The protocol described here was tested in PBMCs and T-cells. Two O2K chambers were injected with PBMCs, and two chambers of an additional O2K were injected with T-cells from the same participant. After injecting substrates malate, pyruvate, and glutamate in all chambers, one chamber of PBMCs and T-cells received oligomycin. Oligomycin prevented any ADP-driven rise in respiration in (A) PBMCs and (D) T-cells or decline in mitochondrial membrane potential in (B,C) PBMCs and (E,F) T-cells. (G,H) ADP sensitivity was greater in PBMCs compared to T-cells. <a href="https://www.jove.com/files/ftp_upload/66863/66863fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66863/66863fig05v2.jpg" /> 
Figure 5: Blank experiments show the change in TMRM fluorescence in response to time and titrations of substrates, uncouplers, and inhibitors (SUIT). (A) Change in TMRM fluorescence in response to titration. (B) Change in TMRM fluorescence in response to time. Experiments were conducted in 0.5 mL chambers filled with Mir05 containing 1 &#956;M TMRM. One chamber did not receive any SUIT titrations (no injection); two chambers in two different instruments received a standard suit protocol (standard injection); one chamber received the same SUIT titrations but with a delay between each injection (delayed injection). <a href="https://www.jove.com/files/ftp_upload/66863/66863fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66863/66863fig06v2.jpg" /> 
Figure 6: Differences in ADP sensitivity between T-cells and monocytes using OCR and mMP. (A) Trace of high-resolution fluorespirometry experiment from a subject&#39;s monocyte and T-cell sample. (B) Oxygen consumption in monocytes (n= 11) and T-cells (n= 13) from the blood of healthy volunteers. (C,D) Non-linear regression fitting of the plotted rise in respiration with ADP titrations to calculate an EC50. (E) Simultaneous measurement of mitochondrial membrane potential. (F,G) Non-linear regression fitting of the plotted decline in mitochondrial membrane potential with ADP titrations to calculate an IC50. (H) Parameters of respiratory capacity of monocytes and T-cells. Data are expressed as mean &#177; SEM for line graphs and mean &#177; SD for bar graphs. Statistically significant differences following t-tests are expressed as *p &#60; 0.05. **p &#60; 0.01, and ****p &#60; for 0.0001. <a href="https://www.jove.com/files/ftp_upload/66863/66863fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/66863/66863fig07v2.jpg" /> 
Figure 7: Comparing ADP response in respiration and mitochondrial membrane potential (mMP) in permeabilized mouse splenic T-cells and liver. (A-D) Response in respiration in permeabilized mouse splenic T-cells and liver. (E-H) Response in mMP in permeabilized mouse splenic T-cells and liver. Fresh liver and spleen were dissected from three mice following cervical dislocation. Splenic Pan T-cells were isolated using antibody-conjugated magnetic bead separation. Both samples underwent the same SUIT protocol in the presence of 1 &#956;M TMRM. (I,J) Comparison of EC50 calculated from the increase in oxygen consumption (OCR) and IC50 from the decrease in mMP in response to ADP. N = 3 per group. Data are expressed as mean &#177; SEM. <a href="https://www.jove.com/files/ftp_upload/66863/66863fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Example SUIT protocol to assess mitochondrial membrane potential in freshly isolated T-cells and monocytes using the 0.5 mL chambers. <a href="https://www.jove.com/files/ftp_upload/66863/Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2: Recommended ADP titration for 0.5 mL chamber. <a href="https://www.jove.com/files/ftp_upload/66863/Table 2.xlsx" target="_blank">Please click here to download this Table.</a>
Table 3: Calculating the average background ratio using five independent blank experiments. <a href="https://www.jove.com/files/ftp_upload/66863/Table 3.xlsx" target="_blank">Please click here to download this Table.</a>
Table 4: Calculating mitochondrial membrane potential (mMP) from sample experiment. <a href="https://www.jove.com/files/ftp_upload/66863/Table 4.xlsx" target="_blank">Please click here to download this Table.</a>
Supplementary Figure 1: Effect of Mir05 and DMSO on mitochondrial respiration and membrane potential. <a href="https://www.jove.com/files/ftp_upload/66863/Supplementray Figure 1.zip" target="_blank">Please click here to download this File.</a>
Supplementary File 1: Reagent preparation and protocol for isolating T-cells from mouse spleen. <a href="https://www.jove.com/files/ftp_upload/66863/Supplementary FIle 1.zip" target="_blank">Please click here to download this File.</a>

Author Spotlight: New Insights into PBMC Mitochondrial Responses Using Fluorespirometry

Methods for studying mitochondrial bioenergetics under physiologically relevant substrate concentrations in immune cells are limited. We provide a detailed protocol that uses high-resolution fluorespirometry to assess changes in the response of the mitochondrial membrane potential to energy demand in human T-cells, monocytes, and peripheral mononuclear cells.

High-Resolution Fluorespirometry to Assess Dynamic Changes in Mitochondrial Membrane Potential in Human Immune Cells

Peripheral mononuclear cells (PBMCs) exhibit robust changes in mitochondrial respiratory capacity in response to health and disease. While these ...

high-resolution-fluorespirometry-to-assess-dynamic-changes

Research

JoVE Journal

Biology

951 Views.  University of Washington. Om een cel te laten functioneren en overleven na een periode van fysiologische stress, moet hij in staat zijn om te voldoen aan de energiebehoefte die nodig is om de homeostase te herstellen. In ons laboratorium proberen we vast te stellen hoe mitochondriën reageren en zich aanpassen aan voedingsstressoren om beter te begrijpen hoe mitochondriën het risico op het ontwikkelen van ziekten mediëren. De huidige methoden voor het testen van PBMC bio-energetica omvatten het meten van de ademhali...

Video: Auteur in de schijnwerpers: nieuwe inzichten in PBMC mitochondriale reacties met behulp van fluorespirometrie

A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in living cells by photo-conversion of matrix targeted photoactivatable GFP (mtPAGFP) in a subset of mitochondria. The rate at which the photoconverted molecules equilibrate across the entire mitochondrial population is used as a measure of fusion activity. Thus far measurements were performed using a single cell time lapse approach, quantifying the equilibration in one cell over an hour. Here, we scale up and automate a previously published live cell method based on using mtPAGFP and a low concentration of TMRE (15 nm). This method involves photoactivating a small portion of the mitochondrial network, collecting highly resolved stacks of confocal sections every 15 min for 1 hour, and quantifying the change in signal intensity. Depending on several factors such as ease of finding PAGFP expressing cells, and the signal of the photoactivated regions, it is possible to collect around 10 cells within the 15 min intervals. This provides a significant improvement in the time efficiency of this assay while maintaining the highly resolved subcellular quantification as well as the kinetic parameters necessary to capture the detail of mitochondrial behavior in its native cytoarchitectural environment.
Mitochondrial dynamics play a role in many cellular processes including respiration, calcium regulation, and apoptosis1,2,3,13. The structure of the mitochondrial network affects the function of mitochondria, and the way they interact with the rest of the cell. Undergoing constant division and fusion, mitochondrial networks attain various shapes ranging from highly fused networks, to being more fragmented. Interestingly, Alzheimer's disease, Parkinson's disease, Charcot Marie Tooth 2A, and dominant optic atrophy have been correlated with altered mitochondrial morphology, namely fragmented networks4,10,13. Often times, upon fragmentation, mitochondria become depolarized, and upon accumulation this leads to impaired cell function18. Mitochondrial fission has been shown to signal a cell to progress toward apoptosis. It can also provide a mechanism by which to separate depolarized and inactive mitochondria to keep the bulk of the network robust14. Fusion of mitochondria, on the other hand, leads to sharing of matrix proteins, solutes, mtDNA and the electrochemical gradient, and also seems to prevent progression to apoptosis9. How fission and fusion of mitochondria affects cell homeostasis and ultimately the functioning of the organism needs further understanding, and therefore the continuous development and optimization of how to gather information on these phenomena is necessary.
Existing mitochondrial fusion assays have revealed various insights into mitochondrial physiology, each having its own advantages. The hybrid PEG fusion assay7, mixes two populations of differently labeled cells (mtRFP and mtYFP), and analyzes the amount of mixing and colocalization of fluorophores in fused, multinucleated, cells. Although this method has yielded valuable information, not all cell types can fuse, and the conditions under which fusion is stimulated involves the use of toxic drugs that likely affect the normal fusion process. More recently, a cell free technique has been devised, using isolated mitochondria to observe fusion events based on a luciferase assay1,5. Two human cell lines are targeted with either the amino or a carboxy terminal part of Renilla luciferase along with a leucine zipper to ensure dimerization upon mixing. Mitochondria are isolated from each cell line, and fused. The fusion reaction can occur without the cytosol under physiological conditions in the presence of energy, appropriate temperature and inner mitochondrial membrane potential. Interestingly, the cytosol was found to modulate the extent of fusion, demonstrating that cell signaling regulates the fusion process 4,5. This assay will be very useful for high throughput screening to identify components of the fusion machinery and also pharmacological compounds that may affect mitochondrial dynamics. However, more detailed whole cell mitochondrial assays will be needed to complement this in vitro assay to observe these events within a cellular environment.
A technique for monitoring whole-cell mitochondrial dynamics has been in use for some time and is based on a mitochondrially-targeted photoactivatable GFP (mtPAGFP)6,11. Upon expression of the mtPAGFP, a small portion of the mitochondrial network is photoactivated (10-20%), and the spread of the signal to the rest of the mitochondrial network is recorded every 15 minutes for 1 hour using time lapse confocal imaging. Each fusion event leads to a dilution of signal intensity, enabling quantification of the fusion rate. Although fusion and fission are continuously occurring in cells, this technique only monitors fusion as fission does not lead to a dilution of the PAGFP signal6. Co-labeling with low levels of TMRE (7-15 nM in INS1 cells) allows quantification of the membrane potential of mitochondria. When mitochondria are hyperpolarized they uptake more TMRE, and when they depolarize they lose the TMRE dye. Mitochondria that depolarize no longer have a sufficient membrane potential and tend not to fuse as efficiently if at all. Therefore, active fusing mitochondria can be tracked with these low levels of TMRE9,15. Accumulation of depolarized mitochondria that lack a TMRE signal may be a sign of phototoxicity or cell death. Higher concentrations of TMRE render mitochondria very sensitive to laser light, and therefore great care must be taken to avoid overlabeling with TMRE. If the effect of depolarization of mitochondria is the topic of interest, a technique using slightly higher levels of TMRE and more intense laser light can be used to depolarize mitochondria in a controlled fashion (Mitra and Lippincott-Schwartz, 2010). To ensure that toxicity due to TMRE is not an issue, we suggest exposing loaded cells (3-15 nM TMRE) to the imaging parameters that will be used in the assay (perhaps 7 stacks of 6 optical sections in a row), and assessing cell health after 2 hours. If the mitochondria appear too fragmented and cells are dying, other mitochondrial markers, such as dsRED or Mitotracker red could be used instead of TMRE.
The mtPAGFP method has revealed details about mitochondrial network behavior that could not be visualized using other methods. For example, we now know that mitochondrial fusion can be full or transient, where matrix content can mix without changing the overall network morphology. Additionally, we know that the probability of fusion is independent of contact duration and organelle dimension, is influenced by organelle motility, membrane potential and history of previous fusion activity8,15,16,17.
In this manuscript, we describe a methodology for scaling up the previously published protocol using mtPAGFP and 15nM TMRE8 in order to examine multiple cells at a time and improve the time efficiency of data collection without sacrificing the subcellular resolution. This has been made possible by the use of an automated microscope stage, and programmable image acquisition software. Zen software from Zeiss allows the user to mark and track several designated cells expressing mtPAGFP. Each of these cells can be photoactivated in a particular region of interest, and stacks of confocal slices can be monitored for mtPAGFP signal as well as TMRE at specified intervals. Other confocal systems could be used to perform this protocol provided there is an automated stage that is programmable, an incubator with CO2, and a means by which to photoactivate the PAGFP; either a multiphoton laser, or a 405 nm diode laser.

Mitochondrial fusion was measured by tracking the equilibration of photoconverted matrix-targeted GFP across the mitochondrial network over time. Thus far, only one cell could be subjected to an hour long kinetic analysis at a time. We present a method that simultaneously measures multiple cells, thereby speeding up the data collection process.

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in ...

Real-time Monitoring of Mitochondrial Respiration in Cytokine-differentiated Human Primary T Cells

During activation, the metabolism of T cells adapts to changes that impact their fate. An increase in mitochondrial oxidative phosphorylation is indispensable for T cell activation, and the survival of memory T cells is dependent on mitochondrial remodeling. Consequently, this affects the long-term clinical outcome of cancer immunotherapies. Changes in T cell quality are often studied by flow cytometry using well-known surface markers and not directly by their metabolic state. This is an optimized protocol for measuring real-time mitochondrial respiration of primary human T cells using an Extracellular Flux Analyzer and the cytokines IL-2 and IL-15, which differently affect T cell metabolism. It is shown that the metabolic state of T cells can clearly be distinguished by measuring the oxygen consumption when inhibiting key complexes in the metabolic pathway and that the accuracy of these measurements is highly dependent on optimal inhibitor concentration and inhibitor injection strategy. This standardized protocol will help implement mitochondrial respiration as a standard for T cell fitness in monitoring and studying cancer immunotherapies.

Metabolic adaptation is fundamental for T cells as it dictates differentiation, persistence, and cytotoxicity. Here, an optimized protocol for monitoring mitochondrial respiration in ex vivo cytokine-differentiated human primary T cells is presented.

During activation, the metabolism of T cells adapts to changes that impact their fate. An increase in mitochondrial oxidative phosphorylation is ...

Immunology and Infection

Real-Time Measurement of the Mitochondrial Bioenergetic Profile of Neutrophils

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and oxidative burst, and create neutrophil extracellular traps (NETs) for microbial clearance. New insights into the metabolic activities of neutrophils challenge the early concept that they primarily rely on glycolysis. Precise measurement of metabolic activities can unfold different metabolic requirements of neutrophils, including the tricarboxylic acid (TCA) cycle (also known as the Krebs cycle), oxidative phosphorylation (OXPHOS), pentose phosphate pathway (PPP), and fatty acid oxidation (FAO) under physiological conditions and in disease states. This paper describes a step-by-step protocol and prerequirements to measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration on mouse bone marrow-derived neutrophils, human blood-derived neutrophils, and the neutrophil-like HL60 cell line, using metabolic flux analysis on a metabolic extracellular flux analyzer. This method can be used for quantifying the mitochondrial functions of neutrophils under normal and disease conditions.

We describe stepwise protocols measuring the mitochondrial respiration of mouse and human neutrophils and HL60 cells using the metabolic extracellular flux analyzer.

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and ...

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 ...

Flow Cytometric Analysis of Multiple Mitochondrial Parameters in Human Induced Pluripotent Stem Cells and Their Neural and Glial Derivatives

Mitochondria are important in the pathophysiology of many neurodegenerative diseases. Changes in mitochondrial volume, mitochondrial membrane potential (MMP), mitochondrial production of reactive oxygen species (ROS), and mitochondrial DNA (mtDNA) copy number are often features of these processes. This report details a novel flow cytometry-based approach to measure multiple mitochondrial parameters in different cell types, including human induced pluripotent stem cells (iPSCs) and iPSC-derived neural and glial cells. This flow-based strategy uses live cells to measure mitochondrial volume, MMP, and ROS levels, as well as fixed cells to estimate components of the mitochondrial respiratory chain (MRC) and mtDNA-associated proteins such as mitochondrial transcription factor A (TFAM). 
By co-staining with fluorescent reporters, including MitoTracker Green (MTG), tetramethylrhodamine ethyl ester (TMRE), and MitoSox Red, changes in mitochondrial volume, MMP, and mitochondrial ROS can be quantified and related to mitochondrial content. Double staining with antibodies against MRC complex subunits and translocase of outer mitochondrial membrane 20 (TOMM20) permits the assessment of MRC subunit expression. As the amount of TFAM is proportional to mtDNA copy number, the measurement of TFAM per TOMM20 gives an indirect measurement of mtDNA per mitochondrial volume. The entire protocol can be carried out within 2-3 h. Importantly, these protocols allow the measurement of mitochondrial parameters, both at the total level and the specific level per mitochondrial volume, using flow cytometry.

This study reports a novel approach to measure multiple mitochondrial functional parameters based on flow cytometry and double staining with two fluorescent reporters or antibodies to detect changes in mitochondrial volume, mitochondrial membrane potential, reactive oxygen species level, mitochondrial respiratory chain composition, and mitochondrial DNA.

Mitochondria are important in the pathophysiology of many neurodegenerative diseases. Changes in mitochondrial volume, mitochondrial membrane potential ...

Neuroscience

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+ ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...

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 ...

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, various mitochondrial quality control mechanisms cooperate to maintain a healthy mitochondrial network. One such pathway is mitophagy, where PTEN-induced kinase 1 (PINK1) and Parkin phospho-ubiquitination of damaged mitochondria facilitate autophagosome sequestration and subsequent removal from the cell via lysosome fusion. Mitophagy is important for cellular homeostasis, and mutations in Parkin are linked to Parkinson's disease (PD). Due to these findings, there has been a significant emphasis on investigating mitochondrial damage and turnover to understand the molecular mechanisms and dynamics of mitochondrial quality control. Here, live-cell imaging was used to visualize the mitochondrial network of HeLa cells, to quantify the mitochondrial membrane potential and superoxide levels following treatment with carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a mitochondrial uncoupling agent. In addition, a PD-linked mutation of Parkin (ParkinT240R) that inhibits Parkin-dependent mitophagy was expressed to determine how mutant expression impacts the mitochondrial network compared to cells expressing wild-type Parkin. The protocol outlined here describes a simple workflow using fluorescence-based approaches to quantify mitochondrial membrane potential and superoxide levels effectively.

Author Spotlight: Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells  

This technique describes an effective workflow to visualize and quantitatively measure mitochondrial membrane potential and superoxide levels within HeLa cells using fluorescence-based live imaging.

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, ...

Real-Time Quantification of Mitochondrial Respiration in Muscle Fibers

Measurement of Mitochondrial Respiration in Human and Mouse Skeletal Muscle Fibers by High-Resolution Respirometry

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle is linked to prevalent metabolic disorders (e.g., diabetes and obesity), muscular dystrophies, and sarcopenia. While there are many techniques to evaluate mitochondrial content and morphology, the hallmark method to assess mitochondrial function is the measurement of mitochondrial oxidative phosphorylation (OXPHOS) by respirometry. Quantification of mitochondrial OXPHOS provides insight into the efficiency of mitochondrial oxidative energy production and cellular bioenergetics. A high-resolution respirometer provides highly sensitive, robust measurements of mitochondrial OXPHOS in permeabilized muscle fibers by measuring real-time changes in mitochondrial oxygen consumption rate. The use of permeabilized muscle fibers, as opposed to isolated mitochondria, preserves mitochondrial networks, maintains mitochondrial membrane integrity, and ultimately allows for more physiologically relevant measurements. This system also allows for the measurement of fuel preference and metabolic flexibility - dynamic aspects of muscle energy metabolism. Here, we provide a comprehensive guide for mitochondrial OXPHOS measurements in human and mouse skeletal muscle fibers using a high-resolution respirometer. Skeletal muscle groups are composed of different fiber types that vary in their mitochondrial fuel preference and bioenergetics. Using a high-resolution respirometer, we describe methods for evaluating both aerobic glycolytic and fatty acid substrates to assess fuel preference and metabolic flexibility in a fiber-type-dependent manner. The protocol is versatile and applicable to both human and rodent muscle fibers. The goal is to enhance the reproducibility and accuracy of mitochondrial function assessments, which will improve our understanding of an organelle important to muscle health.

Author Spotlight: Unveiling Mitochondrial Function and Cellular Metabolic Adaptation in Metabolic Diseases

Here, we describe a comprehensive method for measuring mitochondrial oxidative phosphorylation in fresh permeabilized skeletal muscle fibers from either human or mouse muscle. This method allows for the real-time quantification of mitochondrial respiration and the assessment of fuel preference and metabolic flexibility while preserving existing mitochondrial networks and membrane integrity.

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle ...

High-Resolution Fluorespirometry to Assess Dynamic Changes in Mitochondrial Membrane Potential in Human Immune Cells

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