The development of this protocol was supported by grants from the Taiwan Ministry of Science and Technology (MOST) NSC103-2320-B-010-002-MY2 and MOST104-2628-B-010-002-MY4 to C.L. Hsu. C.W. Wei is a recipient of Excellent Thesis Award of Institute of Microbiology and Immunology, National Yang-Ming University.

The mouse tissue harvest procedure described here has been approved by the Institutional Animal Care and Use Committee (IACUC) of National Yang-Ming University.
1. Preparation of Lymphocyte Suspension from Lymphoid Organs
<ol>
	<li>Euthanize the mouse by an approved method such as CO2 inhalation in a transparent acrylic chamber followed by cervical dislocation to ensure death. 
	NOTE: Keep the flow rate of CO2 for a 20% displacement per minute of the chamber, and no higher than 5 psi (pound per square inch). It takes 2-3 min for a mouse to lose consciousness. Maintain the CO2 flow at least for 1 min after the animal has stopped breathing (5 - 7 min overall).</li>
	<li>Place the mouse on its back on a clean board (e.g. a polystyrene plastic board) and fix the limbs with tape. Rinse with 70% ethanol.</li>
	<li>To harvest spleen, cut and open the abdominal cavity with 11-cm dissecting scissors (the spleen is below the stomach and above the left kidney). Remove the spleen with forceps and clean from connective tissues (use scissors if necessary).</li>
	<li>To harvest the thymus, cut the diaphragm and the rib cage from both sides with dissecting scissors. Use forceps to lift up the sternum to reveal the entire thoracic cavity. Separate the thymus carefully from surrounding tissues with scissors, and remove any residual connective tissue on a paper towel wetted with PBS.</li>
	<li>Mechanically dissociate the tissues with a syringe plunger in a 6-cm dish containing 5 mL ice-cold FACS buffer (PBS supplemented with 2% fetal bovine serum (FBS) and 1 mM EDTA). Transfer the single-cell suspension to a 15-mL centrifuge tube; wash the 6-cm dish with additional 2 mL FACS buffer and combine the wash with cell suspension as well.</li>
	<li>Centrifuge the cell suspension (300 x g, for 5 min at 4 &#176;C) and discard the supernatant. Re-suspend the pellet with 2 mL of Ammonium-Chloride-Potassium (ACK) lysing buffer (155 mM NH4Cl, 10 mM KHCO3 and 0.1 mM Na2EDTA) for 1 min at room temperature (RT) to lyse the red blood cells. Neutralize the ACK buffer by adding 10 mL FACS buffer to the cell suspension.</li>
	<li>Centrifuge the cell suspension (300 x g, for 5 min at 4 &#176;C) and discard the supernatant. Re-suspend cells in 5 mL of complete RPMI medium (RPMI 1640 medium, supplemented with 44 mM NaHCO3, 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 100 U/mL penicillin, 100 mg/mL streptomycin, 2 mM L-glutamine, 1 mM sodium pyruvate, 1% MEM non-essential amino acids and 10% FBS).</li>
	<li>Filter cell suspension through a cell strainer (pore size 70 &#181;m) to remove cell clumps. Count cells with a hemocytometer or automated cell counter.</li>
</ol>
2. Quantification of Mitochondria and Lysosomes
NOTE: Perform the organelle-specific dye staining first because the optimal staining condition for these dyes is incubation at 37 &#176;C. The surface marker staining can be significantly reduced because of antibody capping and internalization at that temperature.
<ol>
	<li>Add 1 x 106 cells to round-bottom FACS tubes, centrifuge cells (300 x g, for 5 min at 4 &#176;C) and discard supernatant.</li>
	<li>Dilute the probe stock solutions to final working concentration (100 nM MitoTracker Green or LysoTracker Green; henceforth mitochondria- or lysosome-specific dye, respectively)&#160;in pre-warmed 37 &#176;C serum-free culture medium. This is the working solution for lysosome- or mitochondria-specific dye. 
	NOTE: Test multiple concentrations including the range of working concentrations suggested by manufacturer (20 - 200 nM for the mitochondria-specific dye and 50 - 75 nM for the lysosome-specific dye used here) in order to obtain the best staining efficiency. Choose a cell population that is known to have good expression of staining target (e.g. lysosomes) as positive control (e.g. human CD14+ monocytes). Choose the working concentration based on good separation between negative and positive staining, as well as good cell viability.</li>
	<li>Resuspend the cells in 100 &#181;L of lysosome- or mitochondria-specific dye working solution and incubate in 5% CO2 incubator at 37 &#176;C for 15 min or 30 min, respectively.</li>
	<li>Add 1 mL of ice-cold FACS buffer to stop the reaction. Pellet cells by centrifugation (300 x g, for 5 min at 4 &#176;C) and discard supernatant. Cells are now ready for surface marker staining.</li>
</ol>
3. Surface Marker Staining
<ol>
	<li>Block Fc receptors with 100 &#181;L of pre-titrated 2.4G2 hybridoma supernatant on ice for 10 min. Wash cells with 1 mL of FACS buffer and centrifuge (300 x g, for 5 min at 4 &#176;C). Discard supernatant.</li>
	<li>Incubate the cells with 50 &#181;L pre-titrated fluorescence-conjugated antibody combination in FACS buffer on ice for 20 min. Avoid light. 
	NOTE: Prepare single color-stained cells that include all the fluorescent antibodies present in the antibody combination and cells treated only with organelle-specific dye for setting voltages of each fluorescence detector and compensation adjustment on flow cytometer. Likewise, prepare the Fluorescence Minus One (FMO) as background controls by using cells stained with all the antibodies, but without staining with organelle-specific dyes.</li>
	<li>Wash the cells by adding 1 mL of FACS buffer and pellet by centrifugation (300 x g, for 5 min at 4 &#176;C). Discard supernatant.</li>
	<li>Resuspend the cells in 300 &#181;L of FACS buffer containing 1 &#181;g/mL propidium iodide (PI) and immediately analyze the samples on flow cytometer.</li>
</ol>
4. Sample Collection on Flow Cytometer
<ol>
	<li>Turn on the computer, flow cytometer, FACS acquisition software and any other accessory devices depending on the machine platform. Run PBS for around 1 min to ensure the flow line is filled with PBS and sheath buffer. Make sure the flow stream runs smoothly.</li>
	<li>Open new experiment and select cytometer parameters (fluorescent detectors) according to manufacturer&#39;s instruction. Filter cells through 70-&#181;m cell strainer and vortex prior to the acquisition of each sample to avoid clumps and doublet formation.</li>
	<li>Make dot plots of forward scatter (FSC) and side scatter (SSC), which represent cell size and granularity, respectively. Optimize the voltages of FSC and SSC to make sure the cells of interest appear in the plots. Make dot plots of FSC-A vs. FSC-W (or FSC-H) and draw a gate for the single cells.</li>
	<li>Make dot plots of each fluorescent parameter. Use unstained cells to determine background signal and use single color-stained controls to adjust voltages of each fluorescent parameter, so that positive and negative cell populations are clearly distinguishable and within the dynamic range of acquisition.</li>
	<li>After all the voltages are set, use auto-compensation if available, or manually adjust the compensation with unstained and single color-stained controls to correct the spectral spillover. Apply the compensation to samples.</li>
	<li>Make a dot plot of each parameter and draw gates according to your experimental set up. For example, FSC-A vs. FSC-H (or FSC-W) to exclude doublets, followed by FSC-A vs. SSC-A for lymphocyte gating; SSC-A vs. PI (live cells) and CD4 vs. CD8 (Figure 1).</li>
	<li>Collect enough events (total cell number) for meaningful comparison between populations. Typically, at least 1,000 events in the population of interest need to be obtained20.</li>
	<li>After all the samples are acquired, export flow data to be analyzed by flow cytometry analysis software. Save the experiment as template to preserve the cytometer setting and parameters for applying to similar experiments in the future.</li>
</ol>
5. Data Analysis
NOTE: There are many tools to analyze flow cytometric data, both commercial and free software. The acquisition software of the flow cytometers can also work for data analysis. Here we used FlowJo as an example.
<ol>
	<li>Start the software and drag the FCS files into the workspace. Click on a sample to open a graphic window. Select parameters to be presented on X and Y axis by clicking on the X and Y axes. Use the toolbar to draw gates according to differentially expressed markers or defined phenotypes of cell populations.</li>
	<li>Add gates of each parameter as shown in Figure 1. Drag gates to all samples in the workspace, so that they get identical gating.</li>
	<li>Drag the plots to layout editor to create graphical reports.</li>
	<li>Display the mean fluorescence intensity (MFI) for each population of interest by clicking the statistics button (&#931;) in a graphic window and choosing &#34;Mean&#34; and the appropriate parameter, for example, the channel used to detect organelle-specific dye. Then click &#34;Add&#34;.</li>
	<li>Click the table editor icon and drag MFI to table editor to create statistical reports. Save as Excel, Text or CSV file to calculate delta MFI (&#916;MFI) as MFI (organelle-specific dyes) - MFI (FMO). 
	NOTE: Do not calculate MFI between samples acquired with different cytometer settings (voltages, fluorescent channels or compensation). Comparisons of values derived from experiments with different settings are meaningless and biased.</li>
	<li>Export all the values from the workspace and plots from layout editor for making tables and figures.</li>
</ol>

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The authors declare no conflicts of interests.

This protocol combines organelle-specific dyes and surface marker staining to quantify the amount of mitochondria or lysosomes in different T cell populations. This method was developed to overcome the limitation of cell number and homogeneity requirements for traditional methods, such as electron microscopy and immunoblot analysis. It is especially useful in analyzing rare cell populations and simultaneously examining multiple cell types at the same time.
The duration of the procedure and the temperature of incubation are the most critical factors in this protocol. Proper organelle labeling requires accurate titration of organelle-specific dyes. Cells should also be strictly kept on ice to enhance the viability and avoid antibody capping and internalization. In addition, the fluorescence of these organelle-specific dyes is vulnerable to light exposure and temperature changes. Thus, immediate analysis of samples once the staining procedure is completed is very important. We have consistently been able to complete the whole experiment within 2 hours.
Since this protocol relies on the capability of detecting multicolor fluorescence by flow cytometer to evaluate intracellular components, the acquisition settings (e.g. PMT voltages) and the compensation of spectra overlap (the spillover of fluorescence into neighboring channels) can heavily influence the intensity of the signal. This is especially relevant when the surface markers needed to identify the relevant cell types increase above six. In this situation, experience in designing multicolor flow cytometry experiments becomes very important for high quality data acquisition.
The immune system protects the body from pathogen insults and infections, and T cell activation has pivotal role in providing essential signals to other immune cells. Recent studies suggest that different T cell populations have unique metabolic programs, and mitochondrial and lysosomal contents are crucial indicators of metabolic changes. However, this phenomenon is not unique to T cells. The differentiation and activation of innate immune cells, such as macrophage and dendritic cells, are also closely linked to their metabolic status23,24. The advantage of this protocol is that it can be adapted to examine any cell populations of interest by using antibodies against lineage-specific markers. Moreover, in combination with other organelle-specific dyes, such as ER- or Cyto-ID, it has the potential to evaluate different organelles or cellular compartments in various types of cells.
The establishment of this mitochondria or lysosome quantification method is not only instrumental to study the metabolism in T cells, but also has great potential to benefit research that focuses on metabolic changes in disease settings, such as cancer or autoimmune diseases25,26.

The activation and proliferation of T cells are critical steps for mounting successful immune responses. Recent advances suggest that the cellular metabolism is tightly associated with both the development and functions of T cells. For example, naive T cells rely largely on oxidative phosphorylation (OXPHOS) to meet the energy demand during the recirculation among secondary lymphoid organs. Upon activation, naive T cells undergo drastic metabolic reprogramming, including the induction of aerobic glycolysis to increase ATP production and to fulfill the tremendous metabolic demands during cell differentiation and proliferation. The cells that fail to follow through the metabolic needs die by apoptosis1,2. During the metabolic reprogramming, mitochondria play important roles since they are the organelles largely responsible for the production of ATP to supply energy for the cell, and the cellular mitochondria content fluctuates during metabolic switches throughout T cell development and activation3. However, the accumulation of superfluous or damaged mitochondria can produce excess ROS that damage lipids, proteins, and DNA, and can eventually lead to cell death4
The excessive or damaged mitochondria resulting from metabolic alterations are removed by a specialized form of autophagy5, known as mitophagy. Mitochondria are wrapped by autophagosomes and then fused with lysosomes for degradation. These close communications between mitochondria and lysosomes have generated great interest6,7. For example, oxidative stress stimulates mitochondria to form mitochondria-derived vesicles (MDVs) that are targeted to lysosomes for degradation in a phosphatase and tensin homolog (PTEN)-induced putative kinase 1 (PINK1) and parkin (an E3 ubiquitin ligase) dependent manner8. It was also found that mitophagy is essential for beige-to-white adipocyte transition9,10. More importantly, lysosomes are not merely a degradation compartment, but also a regulator of cellular signaling. Excessive substrate accumulation due to enzyme deficiencies results in lysosomal dysfunction by disrupting lysosomal membrane permeability and affects Ca2+ homeostasis11. The T cell functional defects in lysosomal acid lipase (LAL)12 or lysosomal metabolite transporter knockout mouse model13 further showed the importance of lysosomes in maintaining T cell homeostasis. Both mitochondria and lysosomes are inseparable parts of cellular metabolic regulation. Therefore, measurement of cellular mitochondrial content has been a crucial indicator to evaluate the T cell metabolic and functional status.
Assays commonly used to quantify cellular mitochondrial or lysosomal contents include immunoblot, electron microscopy, immunofluorescent (IF) staining, and PCR analysis of mitochondrial DNA copy numbers14,15,16. While immunoblot can quantitatively compare protein levels across different samples and electron or IF microscopy can visualize the morphological hallmarks of these organelles17, these assays bear certain technical drawbacks. For example, it is time consuming to acquire sufficient number of cell images with high magnification and resolution or to compare the expression levels of a protein across dozens of samples, making these assays considered to be low throughput methods. Moreover, these assays can only be applied to homogenous cell population, such as cell lines, but not to tissue samples that are composed of mixed populations.
It is also difficult to apply these assays to rare populations, for which the requirement of minimal cell numbers from 106 to 108 is impossible to meet. Finally, cells are usually lysed or fixed during the process, making them incompatible with other methods to extract further information. Compared with the traditional methods, fluorescent-based flow cytometry has a relatively high throughput - the information of all the cells within a sample composed of mixed cell populations can be analyzed and collected simultaneously. In addition, one can detect more than 10 parameters on the same cell and sort the cells based on desired phenotypes for further assays. Reactive fluorescent probes have been used to label lysosomes and mitochondria in live cells and can be detected by flow cytometry18,19. These organelle-specific probes are cell permeable and have physico-chemical characteristics that allow them to be concentrated in specific subcellular locations or organelles. Conveniently, these probes are available in various fluorescent formats, thus enabling their application for multicolor analysis.
This protocol describes in detail how to combine surface marker staining with lysosome- or mitochondria-specific dyes to label lysosomes or mitochondria in live cells. This is especially useful for samples generated from primary tissues and organs, which are often composed of heterogeneous cell populations. Researchers can identify cell populations of interest by their surface marker expression and specifically measure the lysosomal or mitochondrial contents through organelle-specific dyes in these cells. Here, we demonstrate the detailed procedure of flow cytometric analysis that evaluates the lysosomal or mitochondrial mass in the major splenic T cell subpopulations.

<table><tbody><tr><td>10 cm Graefe forceps (straight and serrated)</td><td>Dimeda</td><td></td><td></td></tr><tr><td>11 cm Iris scissors (straight with sharp/sharp heads)</td><td>Dimeda</td><td></td><td></td></tr><tr><td>15 mL centrifuge tube</td><td>Thermo Fisher Scientific</td><td>339650</td><td></td></tr><tr><td>5 mL syringe</td><td>TERUMO&amp;nbsp;</td><td></td><td></td></tr><tr><td>6 cm Petri dish</td><td>&amp;alpha;-plus</td><td></td><td></td></tr><tr><td>70 &amp;micro;m nylon cell strainer</td><td>SPL Lifesciences</td><td>93070</td><td></td></tr><tr><td>Ammonium chloride (NH&lt;sub&gt;4&lt;/sub&gt;Cl) &amp;nbsp;&amp;nbsp;</td><td>Sigma</td><td>A9434</td><td></td></tr><tr><td>Anti-mouse CD25</td><td>Biolegend</td><td>102007</td><td>Clone:PC61</td></tr><tr><td>Anti-mouse CD4</td><td>Biolegend</td><td>100453</td><td>Clone:GK1.5</td></tr><tr><td>Anti-mouse CD44</td><td>Biolegend</td><td>103029</td><td>Clone:IM7</td></tr><tr><td>Anti-mouse CD62L</td><td>Biolegend</td><td>104407</td><td>Clone:MEL-14</td></tr><tr><td>Anti-mouse CD8</td><td>Biolegend</td><td>100713</td><td>Clone:53-6.7</td></tr><tr><td>Anti-mouse TCR&amp;beta;</td><td>Biolegend</td><td>109233</td><td>Clone:H57-579</td></tr><tr><td>BD LSRFortessa</td><td>BD Biosciences</td><td></td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Bio basic</td><td>6381-92-6</td><td></td></tr><tr><td>FALCON 5ml Polystyrene Round-Bottom Tube (FACS tube)</td><td>BD Biosciences</td><td>352052</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Hyclone</td><td>ATD161145</td><td></td></tr><tr><td>Flowjo, LLC</td><td>BD Biosciences</td><td></td><td></td></tr><tr><td>Hydroxyethyl piperazineethanesulfonic acid (HEPES)</td><td>Sigma</td><td>H4034</td><td></td></tr><tr><td>L-glutamine (200 mM)</td><td>Gibco</td><td>A2916801</td><td></td></tr><tr><td>LysoTracker Green DND26</td><td>Thermo Fisher Scientific</td><td>L7526</td><td></td></tr><tr><td>MEM Non-essential amino acids (100X)</td><td>Gibco</td><td>11140050</td><td></td></tr><tr><td>MitoTracker Green FM</td><td>Thermo Fisher Scientific</td><td>M7514</td><td></td></tr><tr><td>Penicillin-Streptomycin (10,000 U/mL)</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>Potassium bicarbonate (KHCO3)</td><td>J.T.baker</td><td>298-14-6</td><td></td></tr><tr><td>Propidium iodide solution</td><td>Sigma</td><td>25535-16-4</td><td></td></tr><tr><td>RPMI 1640 medium (powder)</td><td>Gibco</td><td>31800089</td><td></td></tr><tr><td>Sodium bicarbonate (NaHCO3)</td><td>Sigma</td><td>S5761</td><td></td></tr><tr><td>Sodium pyruvate (100 mM)</td><td>Gibco</td><td>11360070</td><td></td></tr></tbody></table>

multicolor flow cytometry-based quantification of mitochondria and lysosomes in t cells

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

Identification of major T cell subsets in spleen and thymus
In brief, single cell suspensions from spleen and thymus were lysed of red blood cells, incubated with 2.4G2 supernatant, followed by organelle-specific dye and surface marker staining with fluorescence-conjugated antibodies (Table 1). The developmental progression of thymocytes can be described by using antibodies to the co-receptors CD8 and CD4. The most immature thymocytes do not express CD4 or CD8 and are called double negative (DN). Then, they up-regulate both CD4 and CD8 and become double positive (DP). Finally, they down-regulate CD4 or CD8 and become mature single positive (SP) cells ready to leave the organ. The earliest stages of T cell development, when the cells do not express the co-receptors CD4 and CD8, can be further classified on the basis of CD44 and CD25 expression. The earliest progenitors are CD44+ CD25- (DN1), and then they start expressing CD25 and become CD44+CD25+ (DN2) cells. After that, the cells down-regulate CD44 and become CD44-CD25+ (DN3) cells and, finally, they down-regulate CD25 as well to become CD44-CD25- (DN4) cells that are on the way to expressing CD4 and CD8 and becoming double-positive (DP) cells.
In the spleen, the T cells can be divided into cytotoxic CD8+ and helper CD4+ T cells. Both of these populations can be further divided into naive, effector and memory subsets based on CD62L and CD44 staining. The detailed gating strategy is shown in Figure 1.
Quantification of mitochondria mass in different T cell subsets
Mitochondria-specific dyes were used to measure the amounts of mitochondria in T cell populations. We found that the mitochondrial contents were the lowest in DN1, peaked at DN2-DN3, and then slightly decreased in DN4 and DP thymocytes and were even lower in CD4 and CD8 SP thymocytes (Figure 2A). However, CD4 or CD8 SP thymocytes had higher mitochondria staining MFI than splenic CD4 or CD8 T cells (Figure 2B). These observations suggest that mitochondrial content fluctuates during T cell development with immature thymocytes containing more mitochondria than their mature counterparts do, and the naive T cells that recirculate in oxygen-rich blood have even lower mitochondrial mass. Interestingly, this reduction of mitochondrial contents was more prominent in the CD8 than the CD4 T lineage (Figure 2B), and T cell activation further decreased it. Among activated T cells, CD4+ memory T cells had slightly more mitochondria compared to effectors; however, the opposite was the case for CD8+ T cells: CD8+ memory T cells had the lowest mitochondrial mass among all T cell subsets (Figure 2B).
Lysosomal content measurement in various subpopulations of T cells
Using lysosome-specific dye, we observed relatively low, but detectable, lysosomal contents in all T cell populations, with more prominent presence of lysosomes in DN1 thymocytes (Figure 3A). No significant difference was found among the thymocyte subsets from DN1 onward. In the periphery, a relatively high number of lysosomes was found in memory and effector CD8+ T cells. This phenomenon is in line with previous studies showing that activated CD8+ T cells increased their expression of lysosomal-associated membrane protein 1 (LAMP-1)21,22, reflecting their possession of lysosomal-cytotoxic granules (Figure 3B)
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58844/58844fig1.jpg" /> 
Figure 1: Gating strategy of splenic lymphocytes and thymocytes. Cells were pre-gated on FSC/SSC and PI- to obtain live singlets. (A) Total thymocytes from mice were stained with CD4, CD8, CD44, and CD25. CD4 and CD8 expressions were used to distinguish CD4+CD8- SP, CD4-CD8+ SP, CD4+CD8+ DP, and CD4-CD8- DN subsets. The CD4-CD8- DN subset was further divided into CD44+CD25- DN1, CD44+CD25+ DN2, CD44-CD25+ DN3, and CD44-CD25- DN4 subpopulations. (B) Total splenocytes from mice were stained with TCR&#946;, CD4, CD8, CD44, and CD62L. The TCR&#946;+ cells were separated into CD4+ and CD8+ T cells, which were further divided into naive (CD44-CD62L+), memory (CD44+CD62L+), and effector (CD44+CD62L-) populations. The results are representative of three independent experiments with n = 3-4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58844/58844fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58844/58844fig2.jpg" /> 
Figure 2: Representative histograms showing mitochondria staining in each cell population. Cells were stained with mitochondria-specific dye for 15 min at 37 &#730;C, followed by surface marker staining. Fluorescent signal of stained cells was acquired by flow cytometer and analyzed on. (A) Mitochondria staining of DN and DP thymocytes. (B) Mitochondria staining of CD4SP and CD8SP thymocytes and splenic T cell populations. &#916;MFI = MFI (Mitochondria staining) - MFI (FMO). The solid red line represents mitochondria staining, the solid blue line shows FMO control, and the dashed red line represents mean fluorescence intensity of mitochondria staining in each population. The results are representative of three independent experiments with n = 3-4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58844/58844fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58844/58844fig3.jpg" /> 
Figure 3:&#160;Representative histograms showing lysosome staining in splenic lymphocytes and thymocytes. Cells were stained with lysosome-specific dye for 30 min at 37 &#730;C, followed by surface marker staining. Fluorescent signal of stained cells was acquired by flow cytometer and analyzed. (A) Lysosome staining of DN and DP thymocytes. (B) Lysosome staining of CD4SP and CD8SP thymocytes and splenic T cell populations. &#916;MFI = MFI (Lysosome staining) - MFI (FMO). The solid green line represents lysosome staining, the solid blue line shows FMO, and the dashed red line represents mean fluorescence intensity of lysosome staining in each population. The results are representative of one experiment with n = 5. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58844/58844fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Table 1" src="/files/ftp_upload/58844/58844table1.jpg" /> 
Table 1: Multicolor flow cytometry panel designs. The fluorochrome and concentration of antibodies/organelle dyes in the staining mixtures are listed here. It is strongly recommended to titrate and test each antibody when setting up new staining panels.

Watch this Scientific Journal Video about Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells at JoVE.com

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

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

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

multicolor-flow-cytometry-based-quantification-mitochondria-lysosomes

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

12.9K Views.  National Yang-Ming University. This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

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

Flow Cytometric Analysis of Mitochondrial Reactive Oxygen Species in Murine Hematopoietic Stem and Progenitor Cells and MLL-AF9 Driven Leukemia

We present a flow cytometric approach for analyzing mitochondrial ROS in various live bone marrow (BM)-derived stem and progenitor cell populations from healthy mice as well as mice with AML driven by MLL-AF9. Specifically, we describe a two-step cell staining process, whereby healthy or leukemia BM cells are first stained with a fluorogenic dye that detects mitochondrial superoxides, followed by staining with fluorochrome-linked monoclonal antibodies that are used to distinguish various healthy and malignant hematopoietic progenitor populations. We also provide a strategy for acquiring and analyzing the samples by flow cytometry. The entire protocol can be carried out in a timeframe as short as 3-4 h. We also highlight the key variables to consider as well as the advantages and limitations of monitoring ROS production in the mitochondrial compartment of live hematopoietic and leukemia stem and progenitor subpopulations using fluorogenic dyes by flow cytometry. Furthermore, we present data that mitochondrial ROS abundance varies among distinct healthy HSPC sub-populations and leukemia progenitors and discuss the possible applications of this technique in hematologic research.

We describe a method for using multiparameter flow cytometry to detect mitochondrial reactive oxygen species (ROS) in murine healthy hematopoietic stem and progenitor cells (HSPCs) and leukemia cells from a mouse model of acute myeloid leukemia (AML) driven by MLL-AF9.

We present a flow cytometric approach for analyzing mitochondrial ROS in various live bone marrow (BM)-derived stem and progenitor cell populations from ...

Cancer Research

Cell-based Flow Cytometry Assay to Measure Cytotoxic Activity

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells to kill CD4+ T cells loaded with their cognate peptide. Target CD4+ T cells are divided into two populations, labeled with two different concentrations of CFSE. One population is pulsed with the peptide of interest (CFSE-low) while the other remains un-pulsed (CFSE-high). Pulsed and un-pulsed CD4+ T cells are mixed at an equal ratio and incubated with an increasing number of purified CD8+ T cells. The specific killing of autologous target CD4+ T cells is analyzed by flow cytometry after coculture with CD8+ T cells containing the antigen-specific effector CD8+ T cells detected by peptide/MHCI tetramer staining. The specific lysis of target CD4+ T cells measured at different effector versus target ratios, allows for the calculation of lytic units, LU30/106 cells. This simple and straightforward assay allows for the accurate measurement of the intrinsic capacity of CD8+ T cells to kill target CD4+ T cells.

This protocol describes a sensitive, cell-based cytotoxicity assay. By enumerating the decrease in frequency of live target CD4+ T cells in the presence of an increasing number of effector CD8+ T cells, this assay allows for the direct assessment of cytolytic activity of antigen-specific CD8+ T cells.

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells ...

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 ...

Biology

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

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

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

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

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

Assessing Autophagic Flux by Measuring LC3, p62, and LAMP1 Co-localization Using Multispectral Imaging Flow Cytometry

Autophagy is a catabolic pathway in which normal or dysfunctional cellular components that accumulate during growth and differentiation are degraded via the lysosome and are recycled. During autophagy, cytoplasmic LC3 protein is lipidated and recruited to the autophagosomal membranes. The autophagosome then fuses with the lysosome to form the autolysosome, where the breakdown of the autophagosome vesicle and its contents occurs. The ubiquitin-associated protein p62, which binds to LC3, is also used to monitor autophagic flux. Cells undergoing autophagy should demonstrate the co-localization of p62, LC3, and lysosomal markers. Immunofluorescence microscopy has been used to visually identify LC3 puncta, p62, and/or lysosomes on a per-cell basis. However, an objective and statistically rigorous assessment can be difficult to obtain. To overcome these problems, multispectral imaging flow cytometry was used along with an analytical feature that compares the bright detail images from three autophagy markers (LC3, p62 and lysosomal LAMP1) and quantifies their co-localization, in combination with LC3 spot counting to measure autophagy in an objective, quantitative, and statistically robust manner.

Here, multispectral imaging flow cytometry with an analytical feature that compares bright detail images of 3 autophagy markers and quantifies their co-localization, along with LC3 spot counting, was used to measure autophagy in an objective, quantitative, and statistically robust manner.

Autophagy is a catabolic pathway in which normal or dysfunctional cellular components that accumulate during growth and differentiation are degraded via ...

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

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

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

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

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper (cTfh) Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the lymph node microenvironment in B-cell non-Hodgkin&#39;s lymphoma is analyzed. Subsets of circulating T-follicular helper cells (cTfh), the circulating memory compartment of Tfh cells in the blood, are also perturbed in disease and therefore represent potential novel predictive biomarkers. Peripheral blood-based testing is advantageous because it is relatively non-invasive and allows simple serial monitoring.This article describes a method for isolating CD4+ T-cells from human blood, and further analysis by flow-cytometry to enumerate cTfh cells and the proportions of their various subsets (cTfhPD-1-/+/hi, cTfh1,2,17 and cTfh1/17). The level of these subsets was then compared between normal subjects and patients with lymphoma. We found that the method was robust enough to obtain reliable results from routinely collected patient material. The technique we describe for the analysis can be easily adapted to cell sorting and downstream applications such as RT-PCR.

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Here, a protocol for the isolation and characterization of CD4+ T-cell subsets from human peripheral blood is described. Purified CD4+ T-cells are analyzed by flow cytometry to determine proportions of T-follicular helper cell subsets.

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the ...

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 ...